Transverse Aortic Arch Banding in Mouse Model: A Minimally Invasive Technique in Mice for Induction of High Pressure in the Left Ventricle

Overview

In this video, we demonstrate an experimental protocol for minimally invasive transverse aortic constriction in mice to examine pressure overload-induced heart failure.

Protocol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Operation Site Preparation

1. Disinfect the operation site with 75% isopropyl alcohol. Adjust the heating pad temperature to 37 °C to avoid changes in heart and respiratory rates of the subject animal.
2. Use surgical instruments sterilized in a gravity displacement sterilizer at 121 °C for 30 min or by other appropriate methods.
3. Take 2–4 30-gauge needles and curve them with a needle holder, as seen in Figure 1, to make a tool for ligation and tissue separation and for use as a retractor. Mount the now-curved needle on a cotton applicator for use in surgery.
4. Blunt a 27-gauge needle and curve it 90º with the needle holder. Smooth the tip by rubbing a hard surface to make a 0.4 mm spacer in ligation step.

2. Animal Preparation

1. Anesthetize a 2-month-old, male, 22–25 g C57BL/6N mouse with an appropriate ketamine and xylazine mixture in relation to body weight: 100 mg/kg + 10 mg/kg i.p. Confirm the anesthetization with the negative tail pinch reflex.
2. Remove the hair on the anterior neck and chest with a 2 to 3 min application of depilatory cream. Remove the cream with warm water and a cotton applicator.
3. Apply ophthalmic ointment or gel to the eyes of the animal to prevent the drying out of the cornea.
4. Place the animal on a heating pad in the supine position and conduct intra-operative monitoring, being sure to keep appropriate documentation at every 15 min interval.
5. Use surgical paper tape to secure the limbs to the heating pad in order to keep the animal in the correct position during the surgery.
6. Disinfect the skin alternately with alcohol and povidone-iodine solution three times.
7. Use a sterile drape with an exposed operation field to prevent contamination. Use new sterile gloves for each mouse and as necessary.

3. Ligation Procedure

1. Open the skin of the mouse at the midline position of the neck and chest with a scalpel.
2. Pull the thyroid gland towards the head by gently separating connective tissues with blunt scissors.
3. Separate the muscle layer on the trachea at the midline towards both sides with the curved needles.
4. Using the blunt scissors, cut the sternum to the second rib (approximately 5 mm). Open the cut with a retractor or curved forceps.
5. Separate the thymus lobes from one another and the lower chest wall by separating the connective tissue with the curved needle; the transverse aortic arch and two carotid arteries will be clearly visible at this point.
6. Place the curved needle under the arch and perforate between the vessel wall and connective tissue on the other side by pushing the curved head out.
7. Using the curved needle, pull the 6-0 monofilament suture under the aortic arch.
8. Place the spacer in the loop and fix the suture in place with a double-knot. Remove the spacer gently.
9. Confirm a successful constriction with the knot position and cut the ends of the suture.
10. Close the chest wall using 6/0 silk suture with a simple interrupted suture pattern. Close the skin with a 6/0 monofilament suture in a continuous suture pattern.

4. Post-operative Care

1. Apply povidone-iodine solution to the suture site and place the animal in a pre-warmed cage for recovery and post-operative monitoring.
2. Inject buprenorphine (0.05–0.1 mg/kg s.c. every 12 h) after the animal has regained consciousness to relieve pain and provide the mouse with appropriate soft food.
3. Return the animal to a 12 h light/dark cycle room after full recovery.

Representative Results

Figure 1: Process of forming the desired needle hooks by bending a 1-in, 30-gauge needle tip. (A) The original 1-in, 30-gauge needle tip. (B) Grasp the needle tip with the needle holder and bend approximately halfway along its length. Bend the needle into a loop, using the needle holder and the hands, as necessary, to form the loop pictured in the upper-right of the figure. (C) Cover the needle with a polyurethane tube in order to hold the bent segment in place. (D) Using the needle holder, bend the tip of the loop into the desired shape. In this case, a needle tip with two 45–60° bends has been shown for illustrative purposes.

Materials

Name	Company	Catalog Number	Comments
1 inch 30-gauge Needles	BD, Franklin Lakes, NJ 07417, USA	305128	Curved as in the procedure and Figure 1
27-gauge Needle	BD, Franklin Lakes, NJ 07417, USA	301629	Make blunt and smooth for spacer
6/0 Dafilon	B. Braun, 34212 Melsungen, Hessen, Germany	C0933066
6/0 Silkam	B. Braun, 34212 Melsungen, Hessen, Germany	C0762067
Sterile gloves	A.R. Medicom, Inc (Asia), Hong Kong
Cotton Applicator	Mannings, Quarry Bay, Hong Kong		Local Shopping Center
Depilatory Cream	Veet (Hong Kong), Kwun Tong, Kowloon, Hong Kong		Local Shopping Center
Lexer-Baby Scissor	FST, British Columbia V7H 0A6, Canada	14078-10
Curved Iris Forcep	FST, British Columbia V7H 0A6, Canada	11065-07
Micro Olsen-Hegar Needle Holder	WPI, Sarasota, FL 34240, USA	501989
Stereo Microscope	WPI, Sarasota, FL 34240, USA	PZMIII-BS
GenTeal Eye Gel	Novartis, East Hanover, NJ 07936, USA		Local Pharmacy
Heating Pad	Kent Scientific Corporation, Torrington, CT 06790, USA	DCT-20
Surgical tape	Laboratory Animal Unit
Ketamine and Xylazine	Laboratory Animal Unit
Betadine	Laboratory Animal Unit
Buprenorphine	Laboratory Animal Unit