Left Coronary Artery Ligation: A Surgical Murine Model of Myocardial Infarction

Summary

Presented here is a surgical procedure for permanent ligation of the left coronary artery in mice. This model can be used to investigate the pathophysiology and associated inflammatory response after myocardial infarction.

Abstract

Ischemic heart disease and subsequent myocardial infarction (MI) is one of the leading causes of mortality in the United States and around the world. In order to explore the pathophysiological changes after myocardial infarction and design future treatments, research models of MI are required. Permanent ligation of the left coronary artery (LCA) in mice is a popular model to investigate cardiac function and ventricular remodeling post MI. Here we describe a less invasive, reliable, and reproducible surgical murine MI model by permanent ligation of the LCA. Our surgical model comprises of an easily reversible general anesthesia, endotracheal intubation that does not require a tracheotomy, and a thoracotomy. Electrocardiography and troponin measurement should be performed to ensure MI. Echocardiography at day 28 after MI will discern heart function and heart failure parameters. The degree of cardiac fibrosis can be evaluated by Masson's trichrome staining and cardiac MRI. This MI model is useful for studying the pathophysiological and immunological alterations after MI.

Introduction

Cardiovascular disease is a major public health concern that claims 17.9 million lives each year, accounting for 31 percent of global mortality¹. The most prevalent type of cardiovascular anomaly is coronary heart disease, and myocardial infarction (MI) is one of the major manifestations of coronary heart disease². MI is usually caused by thrombotic occlusion of a coronary artery due to the rupture of a vulnerable plaque³. The resulting ischemia causes profound ionic and metabolic changes in the affected myocardium, as well as a rapid decrease in systolic function. MI results in the death of cardiomyocytes, which can further lead to ventricular dysfunction and heart failure⁴.

Research on MI in patients is limited due to the scarcity of tissues obtained from patients with MI⁵. As such, murine models of MI are useful in both studying disease mechanisms as well as developing potential therapeutic targets. Currently available murine models of MI includeirreversible ischemia models (LCA and ablation methods) and reperfusion models (ischemia/reperfusion, I/R)⁶. Permanent ligation of the left coronary artery (LCA) in mice is the most used method, and it imitates the pathophysiology and immunology of MI in patients^7,8,9. Permanent MI can also be induced by ablation methods, which involve electrical damage or cryoinjury. Ablation methods are able to generate uniform-sized infarction at the precise location¹⁰. On the other hand, scar formation, infarct morphology, and molecular signaling mechanisms may vary among the ablation methods^10,11. The murine I/R method is another important MI model as it represents the clinical scenario of reperfusion therapy¹². The I/R model is associated with challenges such as a variable infarct size, difficulty in distinguishing responses of initial injury, and reperfusion⁶.

Although widely used, LCA ligation methods are associated with low survival rates and post-operative pain¹³. This protocol demonstrates the murine surgical MI model of LCA ligation that involves the preparation and intubation of mice, LCA ligation, post-operative care, and validation of MI. Rather than using an invasive tracheotomy¹⁴, this method employs endotracheal intubation. The animal is intubated by illuminating the oropharynx using a laryngoscope, making the procedure easier, safer, and less traumatic¹⁵. The mouse is kept on ventilator support and under isoflurane anesthesia throughout the procedure. Further, echocardiography and Masson's trichrome staining are performed to evaluate heart function and cardiac fibrosis after MI, respectively. Overall, this method provides a reliable and reproducible surgical murine model of MI that can be used to study pathophysiology and inflammation after MI.

Protocol

The present study protocol was reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Pittsburgh. Eight (sham n = 4 and MI n = 4) 1-year-old female C57BL/6J mice weighing 24-30 g were used for these experiments. Approximately 100% and at least 80% of mice survived in the first 24 h and 28 days, respectively.

1. Preparation and endotracheal intubation of the mice

1. Preheat a bead sterilizer (see Table of Materials) to 250 °C and place autoclaved surgical instruments in it for a few minutes.
2. Anesthetize the mouse in an induction chamber with 3% isoflurane and 1 L/min oxygen for 5 min.
3. Ensure the depth of anesthesia in the mouse by checking the response to a firm toe pinch.
4. Weigh the mouse to estimate the dosage of the pre-operative analgesic drug, buprenorphine (0.1 mg/kg). Inject the drug intraperitonially.
5. Trim the fur on the left side of the thorax using an electric razor.
6. Disinfect the surgical site with povidone-iodine and 70% ethanol thereafter three times.
7. Place the mouse in the supine position on an inclined board. Secure the head and limbs of the mouse using an elastic band attached to the upper incisors and adhesive tape, respectively. Apply sterile ophthalmic lubricant on the eyes to prevent dryness while under anesthesia.
8. Open the jaw and gently pull the tongue out of the oral cavity.
9. Identify the opening of the larynx by illuminating the oropharynx using a laryngoscope (see Table of Materials).
10. Cut off about 0.5 cm from a 24 G catheter needle and insert the blunt needle into the plastic shield. Direct the blunt needle with the plastic shield into the trachea. Take out the needle, leaving the plastic shield into the trachea.
11. Set the ventilator (see Table of Materials) to a respiratory rate of 137 beats per min (optimized for the mice used in this study) and tidal volume 0.18 cc. Connect the respirator tubes to the catheter shield and confirm correct intubation by looking for a synchronized chest movement with the ventilator.
12. Disconnect the respirator tube from the catheter shield and place the animal in the supine position on a preheated temperature-controlled surgical board. Reconnect the mouse to the ventilator.

2. Permanent ligation of the left coronary artery

1. Disinfect the surgical site with povidone-iodine and 70% alcohol. Apply a sterile drape having a quarter-sized hole in the center to secure the surgical site. Gently lift the skin using a pair of forceps and make a small (1.5-2 cm) cutaneous transverse incision along the line between the left pectoralis major and minor muscles using a pair of surgical scissors.
   NOTE: Scissors were used to make the incision as it provides the required control over the depth and direction of the cut.
2. Separate the underlying pectoralis muscles with forceps and dissecting scissors. The muscles were separated using retractors attached to elastic bands.
3. Make an incision in the third intercostal space with a pair of micro scissors following the natural angle of the ribcage. At this phase, extreme caution must be exercised to prevent injury to the heart and lungs.
4. Gently stretch the ribs apart using retractors to expose the left ventricle. Move the pericardial fat aside and locate the LCA, which runs from the edge of the left atrium towards the apex of the heart.
5. Pass an 8-0 nylon suture under the LCA with the help of a needle holder. Ligate the LCA with a double knot followed by a second knot (a modified surgeon's knot).
   NOTE: Blanching of the lower left ventricle confirms a successful LCA ligation.In addition to this, troponin measurement, ECG monitoring (ST elevation), echo/in vivo cardiac-gated MRI, or micro-CT images are also advised to confirm the comparable MI lesions.
6. Remove the retractors and insert a 22 G catheter needle into the chest cavity. Remove the needle, leaving the tip of the plastic shield in the thoracic cavity. Close the ribcage using a 4-0 nylon suture.
7. Connect a syringe to the 22 G plastic shield and slowly remove excess air trapped in the thoracic cavity by gently pressing the chest to establish a negative air pressure. Remove the plastic shield.
8. Close the skin with a 4-0 nylon suture.
9. Switch off the isoflurane supply. At this stage, the mouse is on the ventilator supplying oxygen.

3. Post-operative care

1. Switch off the ventilator once spontaneous breathing starts.
   NOTE: The procedure takes about 30-35 min per animal from preparation of the mice up to this step.
2. Keep the mouse under a heat lamp and monitor it until it is awake. The animal should not be left unattended until it has recovered enough consciousness to maintain sternal recumbency.
3. After surgery, place the animal is in a separate cage and return it to the original cage with other animals only after it fully recovers.
4. Monitor the mouse daily for any sign of pain or discomfort.
5. Continue intraperitoneal injection of buprenorphine (0.1 mg/kg) every 6-8 hours for an additional 2 days following the surgery.

4. Echocardiographic Evaluation

NOTE: Echocardiography was performed to evaluate the parameters of heart failure on day 28 after MI.

1. After 28 days following the surgery, anesthetize the mice with 3% isoflurane and 1 L/min oxygen, apply sterile ophthalmic lubricant on the eyes, and remove chest hair using hair removal cream. Disinfect the chest area with povidone-iodine and 70% ethanol three times.
2. Secure the anesthetized mice atop the imaging platform (see Table of Materials) in the supine position and maintain a steady level of anesthesia throughout the procedure using a nose cone connected to the anesthetic system (1%-2% isoflurane and 1 L/min oxygen).
3. Tape the four paws to the ECG electrodes with electrode gel (see Table of Materials). Monitor the temperature of the animal by inserting a rectal probe (see Table of Materials).
4. Apply the scanning gel (see Table of Materials) to the chest, place the transducer vertically, lower it to the parasternal line (parallel to the thorax), and rotate 35° anticlockwise to obtain the parasternal long axis view of the left ventricle.
5. Tap the B-mode imaging button on the imaging software (see Table of Materials) to get a complete long axis view of the heart. Adjust the gate size and brightness and save the images using Save Clip or Save Frame for later measurements¹⁶.
6. Switch to M-mode (motion-mode) and place the M-mode axis at the level of the papillary muscle. Adjust the gate size and tap the M-mode Start button. Save the images using Save Clip or Save Frame^16,17.
7. As the 4D mode image acquisition process is automated, verify that the ECG and respiration signals are active (Figure 1) before acquiring the data.
8. Start acquiring the data in B-Mode. Open the 4D scan panel and initiate the 3D motor. Set the image parameters in the 4D scan panel and tap the Scan button to begin the scanning. After reviewing the images in the 2D view, load the images into 4D mode using the Load into 4D button.

Results

Figure 1 demonstrates the representative active ECG and respiration signals during the echocardiographic evaluation of sham (Figure 1A) and MI (Figure 1B) mice. Verification of active ECG and respiration signals are important before acquiring the echocardiographic data. Figure 2 shows echocardiographic measurement of cardiac functional parameters following 28 days after LCA ligation.

Discussion

The murine model of MI is gaining popularity in cardiovascular research laboratories, and this study describes a reproducible and clinically relevant MI model. This protocol improves the LCA ligation process in several ways. To begin with, the use of injectable pre-operative anesthetics such as xylazine/ketamine or sodium pentobarbital^14,15 is avoided. Only isoflurane anesthesia was used, which helps enhance animal survival rates (>80% survival 28 days after ...

Materials

Name	Company	Catalog Number	Comments
22 G catheter needle	Exel INT	26741	Thoracentesis
24 G catheter needle	Exel INT	26746	Endotracheal intubation
4-0 nylon suture	Covetrus	29263	Suturing of muscles and skin
8-0 nylon suture	S&T	3192	Ligation of LAD
Anesthetic Vaporizers	Vet equip	VE-6047	Anesthetic support
Animal physiology monitor	Fujifilm	VEVO 3100	Monitor heart rate,respiration rate and body temperature
Betadine solution	PBS animal health	11205	Antispetic
Buprenorphine	Covetrus	55175	Analgesic
Disecting microscope	OMANO	OM2300S-V7	Binocular
Electric razor	Wahl	79300-1001M	Shaving
Electrode gel	Parker Laboratories	W60698L	Electrically conductive gel
Ethanol	Decon Laboratories	22-032-601	Disinfectant
Forceps	FST	11065-07	Stainless Steel
Gauze	Curity	CAR-6339-PK	Sterile
Heat lamp	Satco	S4998	Post surgery care
Heating pad	Kent scientific	Surgi-M	Temperature control
Hot Bead sterilizer	Germinator 500	11503	Sterilization of surgical instrument
Isoflurane	Covetrus	29405	Anesthesia
Masson’s trichrome staining kit	Thermoscientific	87019	Measurement of cardiac Fibrosis
Micro Needle Holder	FST	12500-12	Stainless Steel
Micro scissors	FST	15000-02	Stainless Steel
Ophthalmic ointment	Dechra	Puralube Vet	Sterile occular lubricant
Scanning Gel	Parker Laboratories	Aquasonic 100	Aqueous ultrasound transmission gel
Scissors	FST	14060-11	Stainless Steel
Small Animal Laryngoscope	Penn-Century	Model LS-2-M	Illuminating the oropharynx
Small animal ventilator	Harvard apparatus	557058	Ventilator support
Surgical light	Cole parmer	41723	Illuminator Width (in): 7
Vevo 3100 preclinical imaging platform	Fujifilm	VEVO 3100	Echocardiography
VevoLAB software	Fujifilm	VevoLAB 3.2.6	Echocardiography data analysis