A Modified Simple Method for Induction of Myocardial Infarction in Mice

Summary

Under adequate anesthesia, the mouse heart was externalized through the intercostal space, and myocardial infarction was successfully induced by ligating the left anterior descending artery (LAD) using materials readily available in most laboratories.

Abstract

Myocardial infarction (MI) represents one of the leading causes of death. MI models are widely used for investigating the pathomechanisms of post-MI remodeling and evaluation of novel therapeutics. Different methods (e.g., isoproterenol treatment, cryoinjury, coronary artery ligation, etc.) have been used to induce MI. Compared with isoproterenol treatment and cryoinjury, coronary artery ligation may better reflect the ischemic response and chronic remodeling after MI. However, traditional methods for coronary ligation in mice are technically challenging. The current study describes a simple and efficient process for induction of MI in mice with readily available materials. The mouse chest skin was cut open under stable anesthesia. The heart was immediately externalized through the intercostal space after blunt separation of the pectoralis major and pectoralis minor. The left anterior descending branch (LAD) was ligated with a 6-0 suture 3 mm from its origin. Following LAD ligation, staining with 2,3,5-Triphenyltetrazolium chloride (TTC) indicated successful induction of MI and temporal changes of post-MI scar size. Meanwhile, survival analysis results showed overt mortality within 7 days after MI, mainly due to cardiac rupture. Moreover, post-MI echocardiographic assessment demonstrated successful induction of contractile dysfunction and ventricular remodeling. Once mastered, an MI model can be established in mice within 2-3 min with readily available materials.

Introduction

Myocardial infarction (MI) represents one of the significant causes of death and disability worldwide^1,2,3,4,5. Despite timely reperfusion, there is currently a lack of effective therapies to treat post-MI cardiac remodeling. Correspondingly, considerable efforts have been made to mechanistic exploration and therapy exploitation for MI^6,7,8. Of note, the establishment of MI models is a prerequisite to meet these ends.

Several methods (e.g., isoproterenol treatment, cryoinjury, coronary artery ligation, etc.) have been proposed to induce MI models in small animals. Isoproterenol treatment is a simple method for MI induction, but it cannot induce infarction of the targeted area⁹. Cryoinjury leads to myocardial necrosis via the generation of ice crystals and disruption of the cell membrane rather than direct ischemia¹⁰. By contrast, coronary artery ligation permits precise control of occlusion site and extent of infarct area and faithfully recapitulates remodeling response following infarction^11,12. Coronary artery ligation is typically performed following intubation, mechanical ventilation, and thoracotomy, which is technically challenging^13,14. Several modified protocols for coronary artery ligation (e.g., ventilation free) were reported and potentiated the induction of MI, but detailed visual demonstrations are lacking^15,16,17. These issues pose a significant financial and technical barrier for groups wishing to engage in research using MI models. This report presents an approach for induction of MI in mice. The current method is easy, timesaving, and uses surgical tools and equipment found readily in most laboratories.

Protocol

The experiments involving animal work are performed with all necessary approvals from the Laboratory Animal Welfare Ethics Committee of Renji Hospital, Shanghai Jiao Tong University, School of Medicine (R52021-0506). Female and male C57BL/6J mice aged between 8-10 weeks were used in the study.

1. Preparation of the simplified anesthesia equipment (OPTIONAL)

NOTE: This is an optional pre-operative set up and can be replaced with titratable anesthesia as mentioned in section 2. The institutional animal ethics committee and veterinarian(s) should be consulted prior to adapting this set up in animal procedures.

1. Take a 15 mL centrifuge tube, and make a cut perpendicular to the long axis of the tube about 3 cm from the opening.
   NOTE: Ensure that the cut is greater than half of the circular circumference of the tube lumen so that the valve can be successfully inserted.
2. Drill holes (diameter, 2 mm) at the centrifuge tube wall between the cut and the tube opening.
3. Cut a suitably sized piece of the valve from a plastic sheet and insert the valve into the cut on the tube wall.
   NOTE: The valve can be used to control the release rate of isoflurane by changing the depth of the insertion.
4. Inside a fume hood, cut open the bottom of the tube and connect it to the oxygen supply. Place a cotton ball near the bottom end of the tube, add load 0.5 mL isoflurane (as obtained, see Table of Materials) onto the cotton ball, and close the valve.
5. Test the anesthesia efficacy by masking the mice with tubes prepared as described above. Monitor the breathing rate and anesthesia depth by toe pinch response.
   ​NOTE: A breathing rate less than 10 times/10 s suggests excessive anesthesia, and the insertion depth of the valve should be adjusted. For all the procedures involving anesthesia, a gas filter filled with activated charcoal sheets must be used (Figure 1A-i), and surgery should be performed within a hood.

2. Operative preparation and anesthesia

1. Prepare and sterilize all the required instruments on the day of surgery, including a pair of forceps, a micro-mosquito hemostat, a pair of surgical scissors, two pairs of needle holders, 4-0 silk surgical suture, 6-0 silk surgical suture, a gas filter, and a light source (see Table of Materials) (Figure 1A).
2. Put on a surgical mask and sterile gloves.
3. Apply the depilatory cream to the mouse chest and wait for 1 min. Gently wipe off the depilatory cream and hair with wet gauze.
4. Hold the mouse with the dominant hand after depilation. Induce anesthesia via inhalation of vaporized isoflurane (4%) with oxygen supply (1L/min) and maintain at 2-3% isoflurane.
5. Confirm adequate anesthesia by the lack of toe pinch response.
6. Apply sterile eye cream to both eyes to prevent corneal dryness.
7. Secure the mice on a surgery platform in the supine position. Apply povidone-iodine swabs (see Table of Materials) to the chest three times and cover the disinfected chest with a sterile drape.

3. Induction of myocardial infarction

1. Change the contaminated gloves to ensure sterility.
2. Make a 0.5 cm skin cut along the line connecting the xiphoid and armpit after local block with lidocaine.
3. Bluntly separate the pectoral major and pectoral minor muscles using forceps and a micro-mosquito hemostat to expose the fourth intercostal space.
4. Open the fourth intercostal space using a micro-mosquito hemostat.
5. Externalize the heart by pushing the heart toward the fourth intercostal space with the index finger of the left hand.
6. Secure the heart with the left hand, and ligate the left anterior descending branch with a 6-0 suture 3 mm from its origin.
7. Place the heart back into the thoracic cavity quickly.
   NOTE: It is safe to externalize the heart for less than 30 s.
8. Evacuate the air out of the thoracic cavity by a gentle press of the chest cavity manually.
9. Close the muscle layer over the ribs with a 6-0 silk suture.
10. Close the skin with a 4-0 silk suture.
11. Place the mice on a pad (37 °C) immediately after the operation.
12. Inject buprenorphine (0.05-0.1mg/kg) subcutaneously every 4-6 hours to reduce post-operative pain for up to 72 hours.
13. Return the operated mice to cages when fully recovered.
    NOTE: The mice will be fully recovered within 3-5 min after surgery.
14. Carefully monitor the mice and provide wet food for up to 7 days.

4. Harvesting the tissues

1. Sacrifice the mice at different time points after MI establishment by cervical dislocation.
2. Secure the sacrificed mice on the surgery platform in the supine position.
3. Make a ventral incision (~3-4 cm) in the upper abdomen. Cut off the ribs from both sides of the thorax cavity, and remove the diaphragm.
4. Perfuse the heart with 10 mL cold phosphate-buffered saline (1x PBS, 4 °C) through intraventricular injection.
5. Collect the heart by cutting off the aortic root and immediately store the heart at -80 °C.
   NOTE: According to the authors' experience, it is feasible to perform TTC staining within two weeks of storage.
6. Stain the heart with 2,3,5-Triphenyltetrazolium chloride (TTC).
   1. Slice the frozen heart into 1 mm thick sections on ice using razor blades.
   2. Incubate the prepared heart slices in 1% TTC solution (dissolved in 1x PBS) at 37 °C for 10-15 min.
      NOTE: After 15 min incubation, discard the TTC solution and immerse the stained heart slices into 1x PBS.
7. Photograph the slices using a digital camera.

Results

The experimental protocol and some of the critical steps are shown in Figure 1. The simplified anesthesia equipment induced anesthesia. As shown in Figure 2A, the induced anesthesia was stable, as reflected by the regular breathing rates (varied from 90-107 breaths/min in the tested mice). Following coronary artery ligation, TTC staining analysis indicated successful induction of myocardial infarction and temporal changes of post-MI scar size (

Discussion

The present report demonstrated a easy protocol for MI induction in mice with readily available materials, which was modified from a method reported by Gao¹⁶. Murine MI models are indispensable for mechanistic exploration and drug screen for post-MI dysfunction and remodeling¹². Among the existing techniques for MI induction, coronary artery ligation represents the most commonly practiced one. Coronary artery ligation faithfully recapitulates the ischemia nature of myo...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
2,3,5-Triphenyltetrazolium chloride	SIGMA	T8877-25G	TTC staining
4-0 silk suture	YUANKANG	4-0	Surgical instrument
Autoclave	HIRAYAMA	HVE-50	Sterilization for the solid
Buprenorphine	Qinghai Pharmaceutical FACTORY Co., Ltd.	H10940181	reduce post-operative pain
Centrifugation tube	Biological Hope	1850-K	15ML
Depilatory cream	ZIKER BIOTECHNOLOGY	ZK-L2701	Depilation agent for laboratory animals
Forcep	RWD	F12028	Surgical instrument
Gas filter	ZHAOXIN	SA-493	Operator protection
Isoflurane	RWD	20071302	Used for anesthesia
Light source	Beijing PDV	LG-150B	Operating lamp
Micro-mosquito hemostat	FST	13011-12	Surgical instrument
Needle	BINXIONG	42180104	Surgical instrument
Needle and the 6-0 silk suture	JIAHE	SC086	Surgical instrument
Needle holder	ShangHaiJZ	J32030	Surgical instrument
Needle holder	ShangHaiJZ	J32010	Surgical instrument
Povidone-iodine swabs	SingleLady	GB26368-2010	Skin disinfection
Scissors	CNSTRONG	JYJ1030	Surgical instrument
Sterile eye cream	Shenyang Xingqi Pharmaceutical Co., Ltd.	H10940177	prevent corneal dryness
Ultra-high resolution ultrasound imaging system for small animals	VisualSonics	Vevo 2100	Echocardiographic analysis