Precision Ligation of the Left Anterior Descending Coronary Artery Using Anatomical Landmarks in a Murine Myocardial Infarction Model

Summary

This protocol induces myocardial infarction in mice by permanent left anterior descending coronary artery ligation using the anterior interventricular sulcus as a reproducible surface landmark for precise suture placement. Infarction is validated by electrocardiography, echocardiography, histological staining, and serum biomarker analysis.

Abstract

Permanent ligation of the left anterior descending (LAD) coronary artery is the most widely used approach for establishing mouse models of myocardial infarction (MI) in preclinical cardiovascular research. In conventional protocols, however, the ligation site is typically placed 1–2 mm below the left atrial appendage, a landmark that is often difficult to identify reliably during surgery. Combined with considerable inter-individual variability in murine coronary anatomy, this frequently results in imprecise LAD localization, inconsistent ligation sites, and reduced reproducibility between operators. To address these limitations, this protocol introduces the anterior interventricular sulcus (AIS) as a guiding landmark for LAD ligation. The AIS is a shallow surface groove that is directly visible under the microscope, with the LAD running immediately beneath it. Unlike conventional approaches, it provides a stable anatomical reference that supports more consistent selection of the ligation site and is easier for inexperienced operators to identify. The protocol combines standardized isoflurane inhalation anesthesia with non-invasive tracheal intubation and validates MI using electrocardiography, echocardiography, enzyme-linked immunosorbent assay (ELISA), histopathological staining, and three-dimensional coronary artery imaging. This protocol provides a reliable and reproducible experimental method for inducing MI in mice to investigate the pathophysiological and immune changes following MI.

Introduction

Myocardial infarction (MI), one of the leading causes of cardiovascular morbidity and mortality worldwide, poses a substantial threat to human health¹. Characterized by an abrupt onset, a critical clinical course, and a high fatality rate, MI is primarily caused by thrombotic occlusion of the coronary arteries and may lead to profound myocardial ischemia, subsequent cardiomyocyte necrosis, and a cascade of complications, including arrhythmias, heart failure, and sudden cardiac death². Despite notable advances in clinical cardiology, limited access to human myocardial tissue continues to impede investigation of the underlying pathophysiological mechanisms of MI and the development of effective therapeutic strategies³. Consequently, reliable, reproducible, and clinically relevant in vivo animal models that faithfully recapitulate the clinical features and progression of MI remain essential for mechanistic studies and preclinical therapeutic evaluation.

Since Johns and Olson first established a murine MI model in 1954, methodologies for inducing MI in mice have undergone continuous refinement⁴. Permanent ligation of the left anterior descending (LAD) coronary artery under direct microscopic visualization has become the most widely adopted technique because it reproduces key pathological features of human MI, including localized myocardial ischemia, myocardial necrosis, and post-infarction ventricular remodeling⁵. However, conventional LAD ligation requires considerable microsurgical expertise that is typically acquired only through extensive training, limiting procedural consistency and reproducibility across operators and laboratories⁶. Moreover, intraoperative and postoperative complications—including hemorrhage, pulmonary injury or collapse, ligature misplacement, and mechanical tissue damage resulting from inadequate ventilation—contribute to relatively high perioperative mortality, further restricting the widespread application of this model⁷. In response to these limitations, Gao et al. introduced a modified technique involving LAD ligation through cardiac eversion, which simplifies the surgical procedure and lowers the technical barrier to model establishment⁸. Additional experimental approaches, including coronary cryoinjury and pharmacological induction of myocardial injury, have also been developed for specific research applications^9,10. Nevertheless, permanent LAD ligation under direct visualization remains the preferred method because it most closely reproduces the principal pathophysiological features of human MI while permitting controlled induction of infarct size and location.

Despite its widespread use, conventional LAD ligation remains limited by variable identification of the ligation site, procedural inconsistency, relatively high perioperative mortality, and inter-laboratory variability in reproducibility, all of which may affect the reliability and comparability of experimental outcomes¹¹. The overall goal of the present protocol is to establish a more standardized and reproducible method for permanent LAD ligation in mice by using the anterior interventricular sulcus (AIS) as an external anatomical landmark for vessel localization. Rather than estimating the ligation site at a fixed distance below the left atrial appendage, this protocol identifies the AIS—a shallow surface groove that is directly visible under microscopic visualization and overlies the LAD—as a stable anatomical reference for guiding ligation. Compared with conventional landmark-based estimation, this approach is intended to facilitate more consistent selection of the ligation site, reduce operator-dependent variability, and improve accessibility for less-experienced surgeons while maintaining compatibility with standard murine MI procedures. This protocol is therefore appropriate for investigators seeking a reproducible surgical method for generating permanent MI models for studies of myocardial injury, ventricular remodeling, inflammation, and related cardiovascular disease mechanisms.

Protocol

All experiments involving animals were performed with approval from the Ethics Committee of the Animal Experiment Center of Southwest Medical University (Approval No. SWMU20240146). Male C57BL/6J mice (specific pathogen-free [SPF]), aged 8–10 weeks and weighing 20.0 ± 3.0 g, were used in this study. Acclimate the mice for 1 week under standard laboratory conditions (22°C ± 2°C; 12 h light/12 h dark cycle) before beginning the experiments.

1. Pre-surgical preparation

1. Prepare and sterilize all surgical instruments on the day of surgery, including forceps, ophthalmic scissors, a needle holder, a micro-mosquito hemostat, a V-shaped eyelid retractor, 3/8-circle 7-0 nylon sutures with needle, 6-0 nylon sutures, and 5-0 nylon sutures. Autoclave all metal instruments at 121°C for 20 min before the procedure, and re-sterilize them between animals using a glass-bead sterilizer (250°C for 15 s), followed by cooling on a sterile pad. Use sutures from sealed sterile packaging.
2. Set up two separate working areas to maintain aseptic technique. Prepare one area with forceps, a murine tracheal intubation kit, and an anesthesia machine, and prepare a second sterile surgical area with sterilized instruments, cotton swabs, povidone–iodine, and sterile gauze.
3. Apply depilatory cream to the thoracic region for 1 min. Remove the cream and hair gently using sterile gauze moistened with sterile water or saline.
   1. Hair removal alone does not constitute surgical skin preparation. Perform definitive aseptic skin disinfection and sterile draping only after anesthetizing, intubating, and securing the mouse on the surgical platform, immediately before the skin incision (Step 2.6).
4. Administer cefazolin (300 mg/kg) by intraperitoneal injection 1 h before surgery. Reconstitute cefazolin sodium for injection in sterile 0.9% saline to a working concentration of 30 mg/mL, and administer approximately 0.2 mL per 20 g body weight to achieve the target dose. Continue administration once daily for 5 consecutive days after surgery.
   NOTE: Do not administer systemic or topical pharmacological hemostatic agents in this protocol. Achieve intraoperative hemostasis by applying gentle, continuous pressure with a sterile cotton swab for 10–30 s, or until bleeding stops.

2. Anesthesia Induction and Tracheal Intubation

1. Place the mouse in the induction chamber (16 × 10 × 11 cm) connected to the anesthesia machine. Set the carrier gas flow rate to 0.5 L/min and induce anesthesia with 3.5%–4.0% isoflurane.
   NOTE: Confirm adequate anesthesia by the absence of the pedal withdrawal reflex. Under these settings, the mouse generally reaches a surgical plane of anesthesia within 2–4 min.
2. Position the anesthetized mouse supine on the murine tracheal intubation platform. Suspend the mouse by hooking the upper incisors onto the platform’s rubber band so that the head and neck are extended at an angle of approximately 45°. Align the oral, pharyngeal, and laryngeal axes to bring the glottis into direct view.
3. Perform orotracheal intubation
   1. Perform orotracheal intubation under direct laryngoscopy without tracheotomy to establish controlled mechanical ventilation. Use a 20 G intravenous catheter (outer diameter, approximately 0.9 mm; length, approximately 15 mm) as the endotracheal tube.
   2. Open the mouth and gently displace the tongue to one side using forceps. Insert the blade of a small-animal laryngoscope along the dorsum of the tongue and depress the base of the tongue.
   3. Advance the laryngoscope until the epiglottis is visible. Lift the laryngoscope upward and forward along the axis of the handle to elevate the epiglottis and expose the V-shaped glottic opening and the pale vocal cords.
   4. Advance the catheter through the open glottis during inspiration while maintaining direct visualization of the vocal cords. Remove the laryngoscope after the catheter passes through the glottis.
   5. Advance the catheter to a depth of approximately 5–7 mm. Withdraw the stylet and connect the catheter to the ventilator.
      NOTE: Confirm correct endotracheal tube placement by observing symmetrical bilateral chest-wall movement synchronized with the ventilator. If chest-wall movement is minimal and rhythmic abdominal movement occurs instead, accompanied by progressive lightening of anesthesia, remove the catheter from the esophagus and repeat the intubation.
   6. If the glottis cannot be visualized, withdraw the laryngoscope and reposition the mouse on the intubation platform with the head and neck extended at approximately 45°. Re-align the oral, pharyngeal, and laryngeal axes, remove any oropharyngeal secretions with a sterile cotton swab, and repeat the laryngoscopy. If the mouse shows signs of emergence from anesthesia, return it to the induction chamber, deepen the anesthesia, and then repeat the intubation attempt.
   7. Abort the procedure if the mouse develops cyanosis, laryngeal bleeding, or persistent apnea. Discontinue isoflurane administration and place the mouse on a temperature-controlled heating pad (37°C) until it has fully recovered.
   8. Do not proceed with thoracotomy if tracheal intubation is unsuccessful. Reschedule the procedure for a later session or exclude the animal from the study and record it as a procedural failure.
      NOTE: Repeated intubation attempts may cause laryngeal edema, increasing the risk of postoperative respiratory distress. In most cases, successful intubation is achieved within the first two attempts after direct visualization of the glottis. Proficiency in orotracheal intubation requires practice and familiarity with the technique.
4. Deliver air as the carrier gas at a flow rate of 0.5 L/min and maintain anesthesia with 1.5%–2.0% isoflurane. Set the ventilator to a respiratory rate of 130 breaths/min, a tidal volume of 2 mL, and an inspiratory-to-expiratory ratio of 5:4.
   NOTE: The tidal volume set on the ventilator exceeds the theoretical value estimated from body weight because it compensates for the compressible volume of the ventilator circuit and the dead space of the endotracheal tube. Evaluate ventilation throughout the procedure by monitoring the animal for cyanosis and for signs of barotrauma, including pneumothorax and subcutaneous emphysema.
   1. Monitor the depth of anesthesia continuously throughout the procedure. Assess the respiratory pattern and pedal withdrawal reflex every 2 min. Monitor the heart rate continuously using the electrocardiogram (ECG), and adjust the isoflurane concentration to maintain a heart rate of 400–500 beats/min; an isoflurane concentration of 1.5%–2.0% is generally sufficient. Throughout the procedure, observe for spontaneous respiratory effort against the ventilator, whisker movement, and changes in mucous membrane color.
   2. Increase the isoflurane concentration in 0.5% increments, up to a maximum of 3.0%, if the pedal withdrawal reflex reappears, whisker movement is observed, or spontaneous respiratory effort against the ventilator occurs. Decrease the isoflurane concentration in 0.5% decrements, to a minimum of 1.0%, if the heart rate falls below 350 beats/min. Pause the surgical procedure whenever these signs are observed, continue monitoring the heart rate, respiratory pattern, mucous membrane color, and whisker movement, and resume the procedure only after the physiological parameters have stabilized.
5. Tilt the mouse toward the right side with the left side uppermost to expose the left thoracic wall. Secure the four limbs and tail to the platform using adhesive tape.
6. Disinfect the surgical site with three successive applications of povidone–iodine. For each application, use a fresh sterile cotton swab, beginning at the planned incision site and moving outward in a concentric spiral to approximately 1 cm beyond the intended incision in all directions. Discard the swab after each application, and do not return a used swab to the center of the surgical field. Allow the final application to air-dry completely, then apply a sterile drape, exposing only the surgical field before making the skin incision.

3. Thoracotomy and Exposure of the Heart

NOTE: The overall surgical workflow, including animal positioning, thoracotomy, exposure of the heart, LAD coronary artery ligation, thoracic closure, and postoperative recovery, is illustrated in Figure 1 and Figure 2.

Figure 1: Anatomical localization of the left anterior descending coronary artery ligation site using the anterior interventricular sulcus. (A) Ventral view of the exposed murine heart before ligation, showing the anterior interventricular sulcus (AIS). (B) View of the same region after ligation of the left anterior descending (LAD) coronary artery; the circled region indicates the ligation site. (C) Schematic showing the spatial relationship between the AIS and the coronary arteries. The LAD courses along the AIS, and the ligation point is located at the superior third of the sulcus. RCA, right coronary artery; LCA, left coronary artery; LCX, left circumflex coronary artery. Please click here to view a larger version of this figure.

Figure 2: Surgical procedure for anterior interventricular sulcus-guided ligation of the left anterior descending coronary artery in mice. (A) Position and secure the anesthetized mouse with the left thoracic region depilated. (B) Make a skin incision over the left thoracic wall. (C) Separate the pectoral muscles and identify the appropriate intercostal space for thoracotomy. (D,E) Insert and position the chest retractor to expand the intercostal space and expose the operative field. (F) Visualize the exposed heart and identify the anterior interventricular sulcus (AIS). (G) Pass a 7-0 nylon suture across the AIS and beneath the left anterior descending (LAD) coronary artery. (H) Secure the suture to produce permanent LAD ligation. (I) Remove the chest retractor and close the thoracic wall. (J,K) Close the muscle and skin layers sequentially. (L) Allow the mouse to recover under warming after surgery. Please click here to view a larger version of this figure.

1. Using ophthalmic scissors, make a 1 cm longitudinal skin incision on the left thoracic wall, parallel to the long axis of the sternum and approximately 2 mm lateral to the left sternal border. Expose the underlying pectoral muscles.
2. Using fine forceps, bluntly separate the pectoralis major and pectoralis minor muscles along the direction of their fibers. Retract the muscles laterally to expose the ribs and intercostal spaces, and avoid cutting the muscles to minimize bleeding.
3. Identify the appropriate intercostal space before opening the thoracic cavity. Locate the anterior intercostal artery, a branch of the internal thoracic artery that courses within an intercostal space parallel to the ribs, and identify the third intercostal space immediately cranial to the most consistently visible artery in the fourth intercostal space.
4. If the anterior intercostal artery is not clearly visible, palpate the first rib and count caudally to identify the space between the third and fourth ribs. Confirm that the selected intercostal space provides a direct, unobstructed view of the left atrial appendage and the anterior free wall of the left ventricle before proceeding.
   NOTE: Accurate identification of the third intercostal space is essential because an incorrect thoracotomy site may reduce visualization of the heart and compromise the procedure.
5. Perform a thoracotomy through the left third intercostal space. Gently spread the intercostal space using a V-shaped eyelid retractor until the left atrial appendage and the upper one-third of the left ventricle are clearly exposed within the surgical field.
   NOTE: At this stage, the heart remains covered by the pericardium; therefore, the AIS cannot be identified reliably. Identify the AIS only after opening the pericardium (Step 3.6), as described in Steps 4.2–4.3.
6. Grasp the pericardium gently with one pair of fine forceps and lift it away from the myocardium. Use a second pair of fine forceps to tear the pericardium open slowly and completely expose the heart.

4. Landmark-guided Ligation of the Left Anterior Descending Coronary Artery

1. Pass a 6-0 nylon suture through the chest wall and apply gentle traction to widen the operative field. Expose the AIS completely before identifying the ligation site (Figure 1A).
2. Identify the AIS as the shallow groove running obliquely from the cardiac base toward the apex and marking the surface boundary between the left and right ventricles (Figure 1C). Under the microscope, identify it as a quasi-circular depression located approximately 1–2 mm below the inferior margin of the left atrial appendage.
3. If the AIS is difficult to identify, gently swab the epicardial surface with a cotton tip moistened with warm saline. Re-examine the operative field under the microscope to identify the groove more clearly.
   NOTE: Under the dissecting microscope, the AIS is a readily identifiable shallow groove that anatomically demarcates the interventricular boundary, with the LAD coronary artery coursing immediately beneath it. If the AIS remains indistinct after swabbing, first confirm that the thoracotomy provides adequate exposure. Then identify the LAD directly according to previously reported methods: relative to its accompanying cardiac vein, the artery is pinker in color, smaller in caliber, and distinctly pulsatile¹².
   1. If the AIS or the LAD coronary artery cannot be identified, first verify that the thoracotomy was performed through the left third intercostal space. If necessary, reposition the thoracotomy as described in Steps 3.3–3.5, then repeat identification of the AIS and the LAD as described in Steps 4.1–4.3.
   2. Terminate the procedure if the AIS or the LAD cannot be identified after 10 min of careful inspection, if uncontrollable bleeding occurs, if the total open-chest procedure exceeds 20 min, or if the heart rate remains below 300 beats/min despite adjustment of the anesthetic depth. Deeply anesthetize the mouse with 5% isoflurane, euthanize it by cervical dislocation, and record the animal as a procedural failure for inclusion in the surgical outcome analysis.
4. Pass a 7-0 nylon suture across the groove at the superior third of the AIS (Figure 1B). Insert the needle perpendicular to the epicardial surface to a depth of approximately 0.5–1 mm, with the entry and exit points positioned on opposite sides of the groove and spanning approximately its maximal width.
5. Secure the ligature using a surgeon’s knot. Adjust the ligature until immediate blanching of the myocardium distal to the ligation site occurs after the first throw without appreciable local tissue indentation.
   NOTE: Tighten the suture only enough to occlude coronary blood flow. Excessive tension may injure the adjacent myocardium or cause the suture to cut through the vessel and surrounding tissue.
6. Confirm successful coronary occlusion within 1 min of ligation by observing patchy ischemic blanching of the myocardium distal to the ligation site¹². Confirm the occlusion further by identifying ST-segment elevation on the ECG.
7. If the ischemic area is excessively small or blanching is absent, release the ligature and repeat the ligation procedure.

5. Closure and Postoperative Recovery

1. Carefully remove the V-shaped eyelid retractor. Close the thoracotomy with a single interrupted 5-0 nylon suture by passing the needle around the adjacent third and fourth ribs to re-approximate the third intercostal space. Keep the needle close to the rib surface to avoid injuring the intercostal neurovascular bundle. Before tying the suture, gently compress the thorax at end-expiration to evacuate residual intrathoracic air, then maintain compression while securing the suture with a surgeon’s knot.
2. Close the skin incision in layers using 5-0 nylon sutures. Ensure complete apposition of the wound edges before proceeding.
3. Place the mouse on a temperature-controlled heating pad (37°C), discontinue isoflurane administration, and continue mechanical ventilation until spontaneous respiration resumes. Observe the animal for recovery from anesthesia, including movement of the tail or whiskers and restoration of a regular respiratory pattern of approximately 150 breaths/min.
4. Confirm readiness for extubation by verifying regular spontaneous respiration, visible chest-wall movement synchronized with the ventilator cycle, and purposeful movement of the limbs or whiskers. Withdraw the endotracheal tube slowly during the expiratory phase, then continue monitoring the mouse on the heating pad for 3–5 min to confirm the absence of respiratory distress.
   NOTE: Respiratory distress after extubation is most commonly caused by pneumothorax resulting from incomplete closure of the intercostal space. If respiratory distress occurs, temporarily re-anesthetize and ventilate the mouse, reopen the skin sutures, inspect the intercostal closure, and place additional sutures at any site of air leakage before repeating the extubation procedure.
5. Return the mouse to its home cage after recovery from anesthesia. Administer buprenorphine (0.1 mg/kg, subcutaneously) immediately after surgery and every 4–6 h for the first 48 h.
6. Monitor each mouse at least twice daily for the first 3 postoperative days and once daily thereafter. Assess body weight, posture, activity, respiratory pattern, and the surgical wound, and euthanize any animal meeting the predefined humane endpoints, including persistent labored breathing, inability to access food or water, or a hunched, immobile posture that is unresponsive to stimulation.

6. Electrocardiographic (ECG) Assessment

1. Record electrocardiographic signals using a multi-channel physiological recording system equipped with an ECG amplifier. Insert three subcutaneous limb-lead electrodes into the mouse, connecting the positive input (VIN+) to the left hindlimb, the negative input (VIN−) to the right forelimb, and the ground (GND) to the right hindlimb.
2. Connect the ECG amplifier module to the data acquisition unit. Set the notch filter to 50 Hz (or 60 Hz, according to the local mains frequency), then open the acquisition software and add a new ECG amplifier module under the analog channel settings.
   1. Configure the amplifier with the following parameters: gain = 2000, mode = normal, low-pass filter = 150 Hz, and high-pass filter = 0.05 Hz. Open the acquisition settings, set the sampling rate to 1 kHz, and close the setup window.
   2. After placing the electrodes, click Start to begin recording. Set the display time base to 0.5 s per major division and the vertical scale to 0.25 mV per major division, then record the ECG continuously for at least 30 s. Select a stable, artifact-free segment for analysis and representative presentation.
3. Record the ECG immediately after ligation while maintaining the mouse under 1.5% isoflurane anesthesia and normothermia on a warming pad. Record ECGs from both the MI and normal control (NC) groups at the same time point under comparable anesthetic conditions.

7. Echocardiography

1. Remove the hair from the anterior thoracic region using depilatory cream 1 day before imaging. Anesthetize the mouse with 1.5% isoflurane throughout the examination.
2. Place the mouse supine on a heated platform, secure the four limbs to the ECG electrodes, and apply ultrasound gel to the precordium. Adjust the isoflurane concentration as needed to maintain a heart rate greater than 400 beats/min.
3. Position the ultrasound probe at the third-to-fourth intercostal space along the left sternal border. Direct the probe marker toward the right forelimb at an angle of approximately 30° and align the probe with the long axis of the heart.
4. Perform all imaging using a linear-array transducer with a center transmit frequency of 30 MHz, a bandwidth of 20–46 MHz, and an axial resolution of 50 μm. Set the transmit power to 100%, the image depth to 13 mm, the image width to 11 mm, the dynamic range to 60 dB, and the two-dimensional gain to 40 dB. Acquire B-mode cine loops at a frame rate of 210 frames/s.
5. Save these acquisition parameters as a system preset. Use the same transducer, preset, transmit power, dynamic range, image depth, and image width for all animals and all imaging sessions, and do not modify the image depth between the NC and MI groups.
6. Within the same long-axis view, position the M-mode sampling line at the mid-ventricular level, perpendicular to the interventricular septum and the left ventricular posterior wall, with the sampling gate spanning the full anteroposterior diameter of the left ventricle. Record the M-mode tracing for subsequent analysis.
7. Acquire B-mode images in which the left ventricle, left atrium, and aortic root are clearly visualized, and record cine loops for subsequent analysis. Acquire corresponding M-mode images at the same imaging plane.
8. Measure the left ventricular end-diastolic internal diameter (D_d), left ventricular end-systolic internal diameter (Ds), and interventricular septal thickness from the acquired images. Acquire five consecutive cardiac cycles and report the mean values.
   NOTE: Calculate left ventricular end-diastolic volume (LVEDV) and left ventricular end-systolic volume (LVESV) automatically using the imaging software. Calculate fractional shortening (FS) as FS = (D_d − D_s) / D_d × 100% and ejection fraction (EF) as EF = (LVEDV − LVESV) / LVEDV × 100%.

8. Tissue Harvesting, Histological Staining, and ELISA

1. Tissue harvesting
   1. On day 7 after MI induction, perform echocardiography on each mouse. Anesthetize the mouse with 4% isoflurane, collect approximately 1.5 mL of blood by retro-orbital bleeding, euthanize the mouse by cervical dislocation, and excise the heart.
   2. Allow the blood samples to clot overnight at 4°C. Centrifuge the samples at 2000 × g for 10 min at 4°C, collect the serum, and store it at −80°C until analysis.
2. Enzyme-linked immunosorbent assay (ELISA)
   1. Quantify serum B-type natriuretic peptide (BNP), interleukin-1β (IL-1β), tumor necrosis factor-α (TNF-α), and nuclear factor-κB p65 (NF-κB p65) using enzyme-linked immunosorbent assay (ELISA) kits.
      1. Bring all reagents and serum samples to room temperature and prepare the working solutions according to the kit instructions. Dilute the serum samples 1:5 in phosphate-buffered saline (PBS), that is, one volume of serum plus four volumes of PBS.
      2. Add 50 μL of standards, PBS, and diluted serum samples to the standard, zero, and sample wells, respectively, leaving the blank well empty. Add 100 μL of horseradish peroxidase (HRP)-conjugated detection antibody to all wells except the blank, seal the plate, and incubate at 37°C for 60 min in the dark.
      3. Discard the contents of each well and wash the plate five times with wash buffer, allowing the buffer to remain in each well for 20 s during each wash cycle. Blot the plate dry on lint-free paper.
      4. Add 100 μL of freshly prepared substrate (Solutions A and B mixed at a 1:1 volume ratio) to each well. Incubate the sealed plate at 37°C for 15 min in the dark, add 50 μL of stop solution, and measure the optical density at 450 nm.
   2. Subtract the blank optical density from each measurement and generate a standard curve using a four-parameter logistic (4PL) regression model. Calculate the concentration of each sample from the fitted curve, multiply the value by the dilution factor of 5, and report the mean value from duplicate wells.
3. Tissue processing, embedding, and sectioning
   1. Rinse each excised heart briefly in ice-cold phosphate-buffered saline to remove residual blood. Fix the tissue in 4% paraformaldehyde for at least 24 h at room temperature.
   2. Trim each heart to expose a flat short-axis surface and place it into a labeled tissue cassette. Dehydrate the tissue through a graded ethanol series (75% ethanol for 4 h, 85% ethanol for 2 h, 90% ethanol for 2 h, 95% ethanol for 1 h, and two changes of absolute ethanol for 30 min each). Transfer the tissue through an ethanol–xylene mixture for 5–10 min, clear it in two changes of xylene for 10 min each, and infiltrate it with three changes of molten paraffin for 1 h each.
   3. Embed each heart with the short axis facing the block surface. Cut serial 3 μm sections at the mid-papillary level, float the sections on a 40°C water bath, mount them on glass slides, and dry them at 60°C.
   4. Deparaffinize the sections in two changes of xylene (15 min each), followed by three changes of absolute ethanol (5 min each), 75% ethanol (5 min), and reverse osmosis (RO) water (5 min).
      CAUTION: Paraformaldehyde is a probable carcinogen and a respiratory and skin irritant, and xylene is flammable and neurotoxic. Handle these reagents in a chemical fume hood while wearing aseptic medical gloves, a laboratory coat, and eye protection, and dispose of waste according to institutional hazardous-waste procedures. Ethanol and n-butanol are flammable; keep them away from ignition sources.
4. Histological staining
   1. Hematoxylin and eosin staining
      1. Immerse the deparaffinized and rehydrated sections in hematoxylin for 3–5 min. Rinse the sections in water.
      2. Differentiate the sections in 1% acid alcohol, and rinse them in water. Blue the sections in 0.5% aqueous ammonia, and rinse them again.
      3. Transfer the sections sequentially through 85% ethanol, 95% ethanol, and absolute ethanol for 5 min each. Counterstain the sections in eosin for 3–5 min.
      4. Dehydrate the sections through three changes of absolute ethanol for 5 min each and then in n-butanol for 5 min. Clear the sections through three changes of xylene for 5 min each, and mount them with neutral resin.
         NOTE: Nuclei appear blue, whereas the cytoplasm and extracellular matrix appear red.
   2. Masson’s trichrome staining
      1. Mordant the deparaffinized and rehydrated sections in 2.5% potassium dichromate solution overnight for approximately 15 h at room temperature. Rinse the sections under running water for approximately 30 s until the yellow tint disappears.
      2. Stain the sections in Ponceau–acid fuchsin solution for 6 min until the tissue appears bright red. Extend the staining time if the color remains faint, and rinse the sections under running water until the runoff is colorless.
      3. Drain excess water without allowing the sections to dry. Differentiate the sections in aqueous phosphomolybdic acid for 1–2 min until the collagen appears pale and the muscle fibers remain red.
      4. Transfer the sections directly into aniline blue solution without rinsing. Incubate the sections for 6–30 s.
      5. Differentiate the sections through three to five changes of 1% acetic acid for approximately 8 s per change. Dehydrate them rapidly through three changes of absolute ethanol for approximately 3, 5, and 5 s, respectively.
      6. Clear the sections in n-butanol for 5 min and then in xylene for 5 min. Mount the sections with neutral resin.
         NOTE: Collagen fibers appear blue, whereas muscle fibers, fibrin, and erythrocytes appear red.
   3. Sirius Red staining
      1. Incubate the deparaffinized and rehydrated sections in Sirius Red solution A in a 65°C oven for 30 min. Wash the sections under running water until the yellow tint disappears.
      2. Blot off excess liquid and immerse the sections in Sirius Red solution B for 2 min. Wash the sections briefly and blot off excess liquid.
      3. Immerse the sections in Sirius Red solution C for 30 min. Rinse them briefly for 1–2 s.
      4. Dehydrate the sections through three changes of absolute ethanol for approximately 3 s each. Clear them through three changes of xylene for 5 min each, and mount them with neutral resin.
         NOTE: Under brightfield illumination, collagen fibers appear red against a yellow background.
5. Image acquisition and quantification
   1. Acquire all histological images using a digital slide scanner. Use identical illumination, white-balance, exposure, and magnification settings for all sections from all groups, and include a calibration scale bar in every image.
   2. Calibrate the spatial scale before image analysis. Load a reference image containing the scale bar acquired at the same resolution and pixel dimensions as the study images. Under Measure > Calibration > Set System, create a new spatial calibration, set the unit to μm, define the pixel-to-micrometer conversion by aligning the calibration points with the scale bar, save the calibration, and apply it to all images.
   3. Load a Masson’s trichrome-stained section and outline the left ventricle using the irregular area-of-interest (AOI) tool. Exclude the ventricular cavity, the right ventricle, and tissue-processing artifacts, then record the total left ventricular area.
   4. Open Measure > Count/Size and select Select Colors to open the segmentation dialog. Choose Color Cube Based segmentation and set the parameters as follows: sensitivity = 4, expand selection by 1 color index, minimum color size = 1 pixel, and kernel size = 3 × 3. Apply the same segmentation settings to all sections.
   5. Select blue collagen-positive pixels using the eyedropper tool as seed points. Sample only clearly collagen-positive regions, generate a preview image, and confirm that the mask includes the collagen-positive areas while excluding the red-stained myocardium, ventricular cavity, and background. Adjust the seed points if necessary before applying the mask.
      NOTE: Re-select seed pixels for each section to account for minor staining-intensity differences between sections. Maintain identical segmentation parameters (sensitivity, expansion degree, minimum color size, and kernel size) for all images, and accept the segmentation only when the preview demonstrates complete coverage of collagen-positive areas with exclusion of non-collagen structures.
   6. Apply the segmentation mask and close the dialog. Under Measure > Select Measurements, select Area, click Count, open View > Statistics, and record the summed collagen-positive area.
   7. Calculate the collagen volume fraction (CVF) by dividing the collagen-positive area by the total left ventricular area and express the result as a percentage.

9. Tissue Clearing and Three-Dimensional Coronary Artery Imaging

1. Fluorescence Labeling of the Coronary Vasculature
   1. Inject 200 µL of 2% Evans Blue solution into the lateral tail vein to label the coronary vasculature. Return the mouse to its cage for 60 min to allow systemic circulation of the dye.
   2. Place the mouse in the induction chamber (16 × 10 × 11 cm) connected to the anesthesia machine and induce anesthesia with 4% isoflurane. Euthanize the mouse by cervical dislocation, open the thoracic cavity, excise the intact heart, rinse it in PBS, and fix it in 4% paraformaldehyde (PFA) for 24 h at 4°C.
      CAUTION: PFA is a probable carcinogen and a respiratory and skin irritant. Handle PFA in a chemical fume hood while wearing aseptic medical gloves, a laboratory coat, and eye protection, and dispose of waste according to institutional hazardous-waste procedures.
   3. Prepare a 3% agarose solution in RO water and embed the fixed heart with the ligation site appropriately oriented for imaging. Inject 1 µL of DAPI solution into each of eight sites surrounding the ligation area using a microliter syringe, and incubate the sample at room temperature for 2 days.
2. Tissue clearing
   1. Perform tissue clearing using an FDISCO (three-dimensional imaging of solvent-cleared organs with superior fluorescence-preserving capability) tissue-clearing kit according to the manufacturer's instructions. The kit comprises six reagents (A–F): Reagents A, B, C, D, and E contain 50%, 70%, 80%, 100%, and 100% tetrahydrofuran (THF) in water, respectively, and Reagent F is dibenzyl ether.
      CAUTION: THF is highly flammable, forms explosive peroxides during storage, and is a respiratory and skin irritant. Dibenzyl ether is a skin and eye irritant. Handle all reagents in a chemical fume hood while wearing appropriate personal protective equipment, including aseptic medical gloves, a laboratory coat, and eye protection. Dispose of waste according to institutional hazardous-waste procedures.
   2. Incubate each heart sequentially in Reagents A, B, C, D, and E for 6 h per reagent. Use 10 mL of each reagent per heart, corresponding to a minimum sample-to-reagent volume ratio of 1:5.
   3. Transfer the sample to Reagent F and continue incubation until the tissue becomes optically transparent, typically for approximately 1 h.
   4. Perform all clearing steps at 6°C–8°C with gentle agitation. Place the sample in a glass container wrapped in aluminum foil and protect it from light throughout the procedure to preserve fluorescence.
3. Light-sheet fluorescence imaging
   1. Mount the cleared sample on the sample holder. Use a light-sheet fluorescence microscope equipped with a 4× objective lens (numerical aperture = 0.28; working distance = 20.0 mm).
   2. Immerse the mounted sample in dibenzyl ether within the imaging chamber. Illuminate the specimen from both sides and sequentially excite DAPI at 405 nm and Evans Blue at 647 nm.
   3. Set the acquisition parameters as follows: for the 405 nm channel, use a laser intensity of 3,000 arbitrary units and an exposure time of 30 ms; for the 647 nm channel, use a laser intensity of 4,000 arbitrary units and an exposure time of 30 ms. Acquire all images using a z-step size of 6 μm and an image resolution of 1,024 × 1,024 pixels.
   4. Set the camera acquisition mode to XYCZT, in which lateral (XY) images are acquired first, followed by sequential channel acquisition (C), axial stepping (Z), and time-point acquisition (T), as configured by the imaging software.
4. Image data processing and three-dimensional reconstruction
   1. Stitch the raw TIFF image files using the image-processing software supplied with the microscope. Convert the stitched image files to the IMS format by selecting File > Convert to Imaris.
   2. Import the converted IMS files into the three-dimensional analysis software. Generate volume-rendered reconstructions using the default rendering settings, and export representative images using the Snapshot module.
      NOTE: Three-dimensional reconstructions are generated using the software's default rendering pipeline without modification. No image segmentation, thresholding, denoising, background subtraction, or other preprocessing is applied. The reconstructed images are used for qualitative visualization of the coronary vasculature and are not intended for quantitative analysis.

10. Statistical Analysis

1. Express all data as the mean ± standard error of the mean (SEM). Perform statistical analyses using GraphPad Prism software (version 5.0).
2. Define the experimental unit as the individual animal. Analyze n = 8 mice per group for all functional, biomarker, and histomorphometric endpoints.
3. Perform data collection and analysis using coded sample identifiers while maintaining investigator blinding to group allocation. Reveal the group assignments only after completion of all quantitative analyses.
4. Assess data normality using the Shapiro–Wilk test and homogeneity of variance using Levene's test in GraphPad Prism (version 5.0) before selecting the appropriate statistical test.
5. Compare two groups using a two-tailed unpaired Student's t-test when the assumptions of normality and equal variance are satisfied. Otherwise, perform a Mann–Whitney U test.
6. Consider P < 0.05 statistically significant. Indicate statistical significance as P < 0.05, P < 0.01, and P < 0.001.

Results

The AIS served as the anatomical landmark for localization of the LAD coronary artery and identification of the ligation site (Figure 1A–C). After identification of the AIS, the sequential surgical workflow for MI induction, including thoracotomy, exposure of the heart, LAD ligation, thoracic closure, and postoperative recovery, was performed as illustrated in Figure 2A–L. The simplified protocol enabled rapid and reproducible i...

Discussion

In recent years, clinical and experimental models of MI have become increasingly important in cardiovascular research, yet many questions regarding the pathogenesis of MI and strategies for its treatment remain unresolved. This article describes a refined method for establishing a murine MI model, in which several critical procedural steps warrant particular attention. The first critical step is tracheal intubation. Some previous protocols establish the airway by tracheotomy, which requires incising the cervical skin and...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.9% sterile saline	Sichuan Kelun Pharmaceutical Co., Ltd.	-	Cefazolin reconstitution
3% agarose	Sigma-Aldrich	A0169	Sample embedding
4% paraformaldehyde	Sigma-Aldrich	158127	Tissue fixation
Absolute ethanol	Chengdu Kelong Chemical Co., Ltd.	05.001.0170A	Tissue dehydration; flammable
Acid fuchsin	Aladdin	A104916-100g	Preparation of Ponceau–acid fuchsin solution for Masson's trichrome staining
AcqKnowledge software	BIOPAC Systems	AcqKnowledge 4.0	Electrocardiogram acquisition and analysis
Ammonia solution, 0.5% aqueous	Chengdu Kelong Chemical Co., Ltd.	05.001.2528A	Bluing during hematoxylin and eosin staining
Aniline blue, acid-soluble	Aladdin	A111200-100g	Preparation of aniline blue solution for Masson's trichrome staining
Aseptic medical gloves	Nanchang Feixiang Latex Products Co., Ltd.	-	Chemical and biological protection
Buprenorphine	Qinghai Pharmaceutical Factory Co., Ltd.	H10940181	Postoperative analgesia
Cefazolin sodium for injection	CSPC Zhongnuo Pharmaceutical (Shijiazhuang) Co., Ltd.	-	Perioperative antibiotic prophylaxis
Cotton swabs	Shijiazhuang Kangertai Medical Equipment Co., Ltd.	-	Epicardial swabbing and aseptic preparation
DAPI staining solution	Thermo Fisher	D1306	Nuclear labeling
Depilatory cream	Veet	-	Thoracic hair removal
Digital slide scanner	3DHISTECH	Pannoramic MIDI II	Whole-slide imaging
ECG amplifier	BIOPAC Systems	ECG100C	Electrocardiogram amplification
Electrocardiogram recording system	BIOPAC Systems	MP160	Electrocardiogram recording
ELISA kit, mouse B-type natriuretic peptide	Quanzhou Ruixin Biological Technology Co.	RXW202726M	Serum BNP quantification
ELISA kit, mouse interleukin-1β	Quanzhou Ruixin Biological Technology Co.	RX203063M	Serum IL-1β quantification
ELISA kit, mouse nuclear factor-κB p65	Quanzhou Ruixin Biological Technology Co.	RX201879M	Serum NF-κB p65 quantification
ELISA kit, mouse tumor necrosis factor-α	Quanzhou Ruixin Biological Technology Co.	RX202412M	Serum TNF-α quantification
Embedding cassettes	Jiangsu Shitai	22022	Tissue embedding
Endotracheal catheter, 20 G	Shenzhen Bailide Biotechnology Co., Ltd.	B11103	Oral intubation; outer diameter approximately 0.9–1.0 mm; trimmed to approximately 15 mm before use
Eosin Y, alcohol-soluble	Hefei BASF Biotechnology Co., Ltd.	Y0310-100G	Cytoplasmic counterstaining during hematoxylin and eosin staining
Evans Blue	Jarvisbio	A312001	Coronary vasculature labeling
Fine forceps	RWD Life Science	F12011-10	Tissue handling and pericardial opening
Fine forceps, microsurgical	RWD Life Science	FC12002T-08	Microsurgical tissue handling
Fine scissors	RWD Life Science	S12000-09	Ophthalmic scissors for skin incision and dissection
Glacial acetic acid	Chengdu Kelong Chemical Co., Ltd.	-	Preparation of 1% acetic acid for Masson's trichrome differentiation
Glass slides	Jiangsu Shitai	80312/80302	Section mounting
Glass-bead sterilizer	RWD Life Science	RS3002	Inter-animal instrument sterilization
GraphPad Prism software	GraphPad Software	Version 5.0	Statistical analysis
Heating pad	RWD Life Science	69020	Postoperative thermoregulation
Hematoxylin staining solution	Wuhan Servicebio Technology Co., Ltd.	G1004	Nuclear staining during hematoxylin and eosin staining
Horseradish peroxidase-conjugated detection antibody	Quanzhou Ruixin Biological Technology Co.	Included with ELISA kits	ELISA detection
Hydrochloric acid	Chengdu Kelong Chemical Co., Ltd.	05.001.1337A	Preparation of acid alcohol for hematoxylin differentiation
Image-Pro Plus software	Media Cybernetics	Version 6.0	Collagen volume fraction quantification
Imaris software	Bitplane	Version 7.2.3	Three-dimensional reconstruction and analysis
Isoflurane	RWD Life Science	R510-22-10	Inhalation anesthetic
Isoflurane anesthesia system	RWD Life Science	TAIJI-IE	Isoflurane delivery
Laryngoscope, small animal	Shanghai Yuyan Scientific Instrument Co., Ltd.	SR310-MR	Direct visualization during intubation
Light-sheet fluorescence microscope	Light Innovation Technology	LiToneXL	Three-dimensional fluorescence imaging
LitScan software	Light Innovation Technology	Version 3.31	Image stitching and conversion
Mechanical ventilator	Chengdu Taimeng Software Co., Ltd.	HX-101E	Mechanical ventilation; 130 breaths/min; tidal volume 2 mL
Micro-mosquito hemostat	RWD Life Science	FC21001R-12	Needle holding during ligation and closure
Microliter syringe	Shanghai Gaoge Industry and Trade Co., Ltd.	-	Localized DAPI injection
Microplate reader	Molecular Devices (Shanghai) Co., Ltd.	SpectraMax iD5	Optical density measurement at 450 nm
Microscope, dissecting	Maishidi (Dongguan) Technology Co., Ltd.	MSD204	Intraoperative visualization
Microscope, upright light	Leica	DM500	Histological imaging
Microtome blades	Epredia	MX35 ULTRA	Paraffin sectioning
Modified Sirius Red staining kit	Wuhan Servicebio Technology Co., Ltd.	G1078	Collagen staining; includes Solutions A, B, and C
n-Butanol	Chengdu Kelong Chemical Co., Ltd.	05.001.0886A	Histological dehydration and clearing; flammable
Neutral mounting resin	Shanghai Yiyang	YSQN41-91	Slide mounting
Nylon suture, 3/8-circle 7-0 with needle	Ningbo Medical Needle Co., Ltd.	-	Left anterior descending coronary artery ligation
Nylon suture, 5-0	Ningbo Medical Needle Co., Ltd.	-	Thoracic and skin closure
Nylon suture, 6-0	Ningbo Medical Needle Co., Ltd.	-	Chest-wall exposure and traction
Paraffin embedding station	Wuhan Junjie Electronics	JB-L5	Tissue embedding
Paraffin wax	NA	NA	Tissue infiltration and embedding
Phosphate-buffered saline	Sigma-Aldrich	P3813	Tissue washing and reagent dilution
Phosphomolybdic acid	Tianjin Aopusheng Chemical Co., Ltd.	-	Masson's trichrome differentiation
Ponceau 2R	Aladdin	P104989-25g	Preparation of Ponceau–acid fuchsin solution for Masson's trichrome staining
Potassium dichromate	Zhengzhou Paini Chemical Reagent Factory	7778-50-9	Masson's trichrome mordant
Povidone–iodine	SingleLady	GB26368-2010	Skin disinfection
Reverse osmosis water	Laboratory-purified	Not applicable	Solution preparation
Rotary microtome	Leica Instruments (Shanghai)	RM2016	Paraffin sectioning
Sample-holder adapter	Light Innovation Technology	Supplied with LiToneXL	Mounting cleared samples for light-sheet imaging
Skin adhesive tape	Qingdao Schultz Biotechnology Co., Ltd.	-	Securing limbs and tail
Slide coverslips	Jiangsu Shitai	80340-1630	Coverslipping
Slide dryer	Shaoxing Luyue	101-3B	Slide drying
Stop solution	Quanzhou Ruixin Biological Technology Co.	Included with ELISA kits	ELISA reaction termination
Substrate solutions A and B	Quanzhou Ruixin Biological Technology Co.	Included with ELISA kits	ELISA color development
Tissue flotation bath	Wuhan Junjie Electronics	JK-5/6	Section flotation
Tissue processor	Wuhan Junjie Electronics	JT-12S	Automated tissue processing
Tissue-clearing kit, Reagents A–F	Jarvisbio	JA11012	Solvent-based tissue clearing
Tracheal intubation platform	RWD Life Science	Re20-M	Oral intubation positioning
Ultrasound analysis software	Fujifilm VisualSonics	VevoLAB 3.2.6	Echocardiography data analysis
Ultrasound gel	Tianjin Jinya Electronics Co., Ltd.	TM-100	Echocardiographic coupling
Ultrasound imaging system	Fujifilm VisualSonics	Vevo 3100	Echocardiography
V-shaped eyelid retractor	Suqian Shengshi Medical Equipment Co., Ltd.	SS5009G	Intercostal and rib retraction
Wash buffer	Quanzhou Ruixin Biological Technology Co.	Included with ELISA kits	ELISA plate washing
Water purification system	Sichuan ULUPURE Ultrapure Technology Co., Ltd.	UPT-II-20T	Production of reverse osmosis water
Xylene	Chengdu Kelong Chemical Co., Ltd.	05.001.0586A	Tissue clearing and deparaffinization; flammable and neurotoxic