Murine Left Anterior Descending (LAD) Coronary Artery Ligation: An Improved and Simplified Model for Myocardial Infarction

Podsumowanie

We provide a reliable method for left anterior descending artery (LAD) ligation in a mouse model. This method is comparatively less invasive than other methods, involving endotracheal intubation, a left-sided thoracotomy approach, and thoracentesis. This method can be used as a model for both acute and chronic myocardial infarction (MI).

Streszczenie

Ischemic heart disease (IHD), or acute coronary syndrome (ACS), is one of the leading causes of death in the United States. IHD is characterized by reduced blood supply to the heart, resulting in the loss of oxygen to and the ensuing necrosis of the heart muscle. The MI model has gained popularity for its use as a short-term ischemia-reperfusion model and a long-term permanent ligation model. Below, we describe a reliable method for the permanent ligation of the LAD. With mouse genetic engineering technology becoming more advanced, and with an increasing availability of quality murine surgical instruments, the mouse has become a popular model for MI surgeries. Our surgical model incorporates the use of an easily reversible anesthetic for the rapid recovery of the mouse; a minimally invasive endotracheal intubation without involving a tracheotomy; and a thoracentesis through the original thoracotomy site without creating an additional incision in the chest, as is done in some other methods, to effectively remove excess blood and air from the chest cavity. This method is comparatively less invasive than other methods, which dramatically reduces surgical and post-surgical complications and mortality and improves reproducibility.

Wprowadzenie

Coronary disease, or ACS, is the most prevalent cardiovascular event and will be considered the main cause of morbidity and mortality worldwide in 2020¹. The cause of ACS is the presence of a myocardial thrombosis due to the rupture of a coronary atherosclerotic plaque that blocks or reduces blood flow to the heart tissue². Therefore, there are clinical signs consistent with the presence of acute myocardial ischemia, such as myocardial infarction (MI)^3,4. MI leads to a loss in mass of the cardiomyocytes and a progression to pathological ventricular remodeling, which can lead to ventricular dysfunction and heart failure^5,6.

One of the most effective ways to study IHD has been to mimic human myocardial infarction in an animal model. This is achieved by occluding the LAD in mice. Using this model, we study how the heart can be protected from the damage resulting from IHD.

Over the last decade, researchers have shifted from using larger animal models to smaller animals, including the shift from rats to mice. The smaller mouse model is starting to be preferred for many reasons, including their small size, large litter size, low cost to maintain, and short gestation period, as well as for the expansive availability of transgenic and gene knockout models⁷. Although mice are small in size, new surgical instruments specifically designed for them have aided in this development. Our method utilizes these new surgical instruments.

While several methods implement an invasive tracheotomy, we use a less invasive method of endotracheal intubation. Using overhead illumination of the oropharynx, we intubate without creating any incisions, providing a safer and less traumatic experience for the animal. The mouse is then placed on a ventilator and kept on isoflurane during the entire procedure. Due to the short duration of anesthesia produced by the drug, it only takes a few minutes for the animal to recover from the anesthetic once it is discontinued. Our surgical model also includes a minimally invasive thoracentesis. The careful removal of blood and excess air from the chest cavity using thoracentesis through the original thoracotomy incision has addressed a common post-operative complication of the LAD ligation: the tension pneumothorax. This method, which eliminates the need for the two additional incisions used in other methods-one for the tracheotomy and another for the thoracentesis-has yielded fewer post-surgical complications and has drastically reduced mortality.

Protokół

This animal protocol has been reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) at Rhode Island Hospital.

1. Anesthesia and Intubation

1. Weigh the mouse to calculate the dosage of post-operative pain medication.
2. Place the mouse in an induction chamber and deliver 4% isoflurane for 9 - 10 min, monitoring the animal throughout. Turn on a hot bead sterilizer so that the apparatus can preheat to approximately 250 °C. Preheating will take 15 - 20 min.
3. Once the mouse reaches a deep plane of anesthesia, with a breathing rate of approximately 32 breaths/min, place the mouse supine on a Styrofoam board and use an elastic band secured under the top incisors to hold the mouth open. Confirm sedation by performing a toe pinch. Position a high-intensity illuminator above the mouse so that the oropharynx can be visualized.
4. Use curved forceps to open the jaw and another pair of forceps to lift the tongue out of the way. Be sure to intubate while positioned at or slightly below eye-level with the body of the mouse. The use of surgical loupes is recommended.
5. Visualize the opening and closing of the vocal cords. When open, insert a 20-gauge, 1-in intravenous (IV) catheter with a blunt-tip needle introducer. Use the needle to guide the catheter to the tracheal opening, but avoid inserting the needle into the trachea. Verification of correct placement can be done using a plastic transfer pipette.
6. Transfer the intubated mouse to an operating surface equipped with a heating device. Connect the mouse to a small rodent ventilator set to a stroke volume of 150 µL/stroke and a stroke rate of 130 strokes/min.
7. Deliver 2.5% isoflurane. Verify the intubation by checking for bilateral chest rise. Verify anesthesia by performing a toe pinch. The mouse may need 5 - 10 min on the ventilator to become fully anesthetized.

2. Preparing the Mouse

1. Tape down the intubation tube at the connecting site between the ventilator and the IV catheter. Tape down the extremities. Place sterile lubricating drops on the eyes.
2. Trim the ventral left side of the thorax with an electric razor. Dust off the shaved fur with dry wipes and apply a small layer of hair removal cream using a sterile cotton swab. The cream should remain in contact with the hair follicles for approximately 30 - 45 s.
3. While the cream processes, place three sterile cotton swabs in three 1.5-mL tubes filled with Betadine to soak. Using wipes moistened with distilled water, gently wipe away the cream and fur.
4. Clean the surgical field three times, alternating Betadine and sterile 70% isopropanol prep pads, cleaning in a circular motion moving from center to periphery. Place a sterile drape with a quarter-sized hole over the surgical field of the mouse.
5. Clean the area surrounding the mouse with 70% ethanol. Verify anesthesia once more with a toe pinch.

3. LAD Ligation

1. Place the autoclaved surgical instruments in hot bead sterilizer preheated to 250 °C for approximately 20 s. Place the sterilized instruments on sterile autoclaved surgical drape. Don surgical gloves.
2. Use fine-tip forceps to gently lift the skin at a point approximately 5 mm to the left of the prominent xiphoid cartilage. Use a surgical scalpel with a No. 10 blade to create a vertical incision in the skin from this point upwards, to the level of the manubrium.
3. Use curved forceps to gently separate the skin and muscle layers. Open the muscle layer, following the skin incision. Insert two 5-0 polypropylene sutures through the muscle layer, one on either side of the incision, and secure the sutures temporarily with clamps to hold the muscle layer open.
4. Identify and make an incision in the third intercostal space, following the natural angle of the ribcage. Remove the tape from the left extremities of the mouse and secure its left rear foot to its right rear foot with tape. Cut a longer piece of tape and secure its left front foot to the operating surface in a slightly elevated position. Clean the gloves with 70% ethanol.
5. Use a retractor to gently spread apart the 3^rd and 4^th ribs. Cut a small section of sterile gauze, approximately 1 in x ½ in, and dip it in sterile 0.9% saline. Squeeze out the excess saline and use forceps to gently insert the gauze against the left lung to prevent accidental lung damage during the procedure.
6. Gently remove the thin pericardium with forceps.
7. Tear a small amount of cotton off a sterile cotton swab and roll it into a small ball. Dip this cotton ball into sterile 0.9% saline and gently swab over the surface of the heart to appreciate the arteries. Gently push the left auricle upwards and locate the coronary arteries underneath.
8. Identify the LAD and pass an 8-0 nylon suture under the LAD; complete two throws to secure the ligation. If the ligation is successful, the left ventricle distal from the ligature will blanch.
9. Using forceps, remove the gauze inserted earlier, and then gently remove the retractor. Insert a 6-in, 25-gauge flexible tube attached to a 25-gauge needle into the chest cavity through the thoracotomy opening. Advance approximately 1 - 2 in of tubing into the space above the left lung. Return the mouse to a supine position and clean the gloves with 70% ethanol.
10. Use 5-0 polypropylene sutures in a simple interrupted pattern to close the ribcage, keeping the chest tube in place. Remove the two sutures holding the muscle layer open. Use 5-0 polypropylene sutures in a simple continuous pattern to close the muscle layer, again keeping the chest tube in place.
11. Attach a 1-mL syringe to the 25-gauge needle on the chest tube. Gently pull upwards on the plunger while simultaneously gradually extracting the chest tube from the chest cavity with forceps. Extract the tubing slowly, as this step removes excess air and blood, which would otherwise become trapped in the chest cavity and result in a pneumothorax.
12. Once the syringe is full, detach the syringe from the needle and dispose of the waste in a waste beaker or sink. Continue this process until the chest tube is completely extracted. Ensure that the chest is tightly sealed.
13. Decrease the isoflurane to 1.5%. Close the skin with 4-0 polypropylene sutures in a simple interrupted pattern. Turn the isoflurane vaporizer off.
14. Administer 0.1 mg/mL buprenorphine in 0.9% saline via an intraperitoneal (IP) injection. Topically apply 2 mg/mL Lidocaine with 2 mg/mL Bupivacaine in 0.9% saline to the incision. Administer between 200 - 500 µL of 0.9% saline via a subcutaneous injection, scaling the saline amount to the weight of the mouse.
15. Wait 5 min after administering the pain medications to remove the mouse from the intubation tube. This aids in the transition off the ventilator.
    1. If the mouse does not have a bilateral chest rise once off the ventilator, perform needle decompression. To do this, introduce a 25-gauge sterile needle and a 1-mL syringe between the 3^rd and 4^th ribs until it enters the thoracic cavity, denoted by a sudden decrease in resistance. Pull up gently on the plunger to remove excess air.
16. When the mouse demonstrates an adequate bilateral breathing rate and depth and responds to a toe pinch, place the mouse in a clean recovery cage under a heat lamp. Provide the mouse with moist food and a water bottle, monitoring in a laminar flow hood for 15 - 20 min. Monitor for an exaggerated breathing effort, excessive bleeding, or other potentially life-threatening complications.
17. For the next three days, administer 0.1 mg/mL buprenorphine pain medication via an IP injection twice daily. Monitor the mouse daily.

Wyniki

The mice are euthanized twenty-eight days after surgery, and the hearts are harvested and examined. The mice are anesthetized with 50 - 75 mg/kg ketamine and 5 - 10 mg/kg xylazine. When the animal is under adequate anesthesia, the thoracic cavity is opened, and using a 23-gauge needle, cold potassium chloride (KCL, 30 mM) is injected into the posterior basal region of the heart. The heart is arrested in diastole. For further validation of the ligation, the heart is removed from the animal...

Dyskusje

With an increasing use of the MI model in laboratories, the described procedure seeks to increase the efficiency and survival rate of the mice while minimizing their post-operative pain and discomfort. This protocol strives to minimize mortality by making numerous improvements to various aspects of the LAD ligation procedure. There are a few distinctions. Some murine intubation studies that utilize ketamine and xylazine along with isoflurane for induction, due to the benefit of their longer duration of a...

Materiały

Name	Company	Catalog Number	Comments
High-Intensity Light Source	Harvard Apparatus	72-0215
SurgiSuite Operating Platform	Kent Scientific Corporation	SurgiSuite	Uses a rechargeable, battery-operated far infrared warming pad. Charge overnight before surgery.
SurgiSuite LED Lighting Kit	Kent Scientific Corporation	SURGI-5003
Hot Bead Sterilizer	Fine Science Tools	18000-45	Preheating takes 15 - 20 min. Instruments take 20 s to sterilize.
Small Rodent Anesthesia System	VetEquip Inc.	901810
Isofluorane	Piramal Enterprises	66794-017-10
Buprenorphine	Rhode Island Hospital Pharmacy	NDC 12496-0757-1, 12496-0757-5
Surgical Loupes	Roboz	RS-6687
Small Rodent Ventilator	Harvard Apparatus	73-0043
Lubricating Drops	Thermo Fisher Scientific	19-898-350
Electric Razor	Kent Scientific Corporation	CL 9990-1201
Hair Removal Cream	Nair
Medical Tape	Thermo Fisher Scientific	18-999-380
Betadine	Thermo Fisher Scientific	19-027136
70% Isopropanol Wipes	Thermo Fisher Scientific	22-363-750
Surgical Drapes	Braintree	SP-TS
Surgical Gloves	Thermo Fisher Scientific	18999102D
5-0 Polypropylene Sutures	Ethicon	8630G
8-0 Nylon Sutures	Fine Science Tools	12051-08
Platinum-Cured Tubing	Harvard Apparatus	72-1042	0.3 mm inside diameter x 0.6 mm outside diameter
0.9% Saline	Thermo Fisher Scientific	19-310-207
4-0 Polypropylene Sutures	Ethicon	8631G
1 CC Syringe with 25-Gauge Needle	Thermo Fisher Scientific	14-826-100
Scissors	Kent Scientific Corporation	INSS600225
Forceps	Kent Scientific Corporation	INS700100
Cotton Swabs	Thermo Fisher Scientific	23-400-118
IV Catheter, 20-Gauge	Thermo Fisher Scientific	NC9892181
Retractor	Kent Scientific Corporation	INS 750369
Forceps	Fine Science Tools	11003-12
Dissecting Forceps, Straight	Kent Scientific Corporation	INS 700101
Dissecting Forceps, Curved	Kent Scientific Corporation	INS 700103
Hemostatic Forceps, Straight	Kent Scientific Corporation	INS 750451
Hemostatic Forceps, Curved	Kent Scientific Corporation	INS 750452
Tissue Forceps	Kent Scientific Corporation	INS 700131
Needle Holder	Kent Scientific Corporation	INS 600109
Scissors	Kent Scientific Corporation	INS 600225