Mouse Model of Coronary Collateral Growth Through Repetitive Ischemia

This paper presents a method to study postnatal coronary collateral growth induced by repetitive ischemia in mice, including the surgical implantation of a pneumatic occluder on the left anterior descending artery, an automated inflation system for the repetitive ischemia protocol, and potential methods to evaluate collateral growth.

Coronary collaterals are a natural bypass in ischemic heart diseases (IHD), and so for many years, coronary collateral growth (CCG) has been a promising therapeutic target for IHD, particularly in patients with type 2 diabetes or metabolic syndrome in which CCG is impaired. However, this process is understudied, partly due to the lack of mouse models of CCG, even though other animal models, such as pigs, dogs, and rats, have been established. A mouse model can take advantage of the many genetic modifications available for the species, including lineage tracing and gene regulation (overexpression or knockout), to elucidate the process and mechanism of CCG, including the pathways and cell types involved.  We, therefore, set out to develop a mouse model of CCG induced by repetitive ischemia (RI) via transient, repetitive occlusion of the left anterior descending artery (LAD). This manuscript provides details of this mouse CCG model, including the RI surgery to implant a pneumatic occluder on the LAD, the automated pressure-based inflation system used for controlling the pressure and timing of inflation, and the sequence of the RI protocol. This method has already generated one publication to elucidate the process of CCG induced by RI, showing that sprouting angiogenesis gives rise to mature coronary arteries in CCG in adult mouse hearts.

Ischemic heart disease (IHD) is the leading cause of mortality in the United States, and more than 200,000 coronary artery bypass surgeries are performed annually in an effort to treat the disease¹. Coronary collaterals, anastomoses between branches of the coronary arterial tree, are a natural bypass that can resupply blood to ischemic tissue downstream of a blockage²; however, people exhibit a wide variation in the extent of their native collateral networks^3,4. Patients with IHD who have more extensive coronary collateralization have better outcomes during cardiac events, including reduced infarct size and mortality. Hence, coronary collateral growth (CCG) has been a therapeutic target for over a decade^5,6,7. It is of particular interest for the growing number of patients with metabolic syndrome⁸, who exhibit poorer coronary collateralization⁹. However, until the process and mechanism of CCG are better understood, attempting to induce CCG for the treatment of IHD is unlikely to be fruitful.

Coronary collaterals have been studied in large animal models, and brief, repetitive occlusions of main coronary arteries have been used to induce CCG in pigs¹⁰, dogs¹¹, and rats¹². A mouse model of CCG, however, would have more advantages in studying the molecular and cellular mechanisms of CCG because of the many genetically modified mouse lines readily available, including lineage tracing, gene-specific or cell-specific transgenic and knockout lines. Interestingly, unlike humans, mice are reported to have no native coronary collaterals^13,14, making them an attractive model to study coronary collateral formation. Indeed, a recent report showed that in patients with obstructive artery disease, nearly half (47%) had no collateralization (Rentrop grade 0)³; thus, a mouse model of CCG could be clinically relevant for patients with minimal native collateralization.

We, therefore, developed a mouse model of CCG induced by repetitive ischemia, with an inflatable balloon occluder over the left anterior descending artery (LAD) that uses a pressure-based inflation system automated with a timer. The repetitive ischemia protocol is able to stimulate collateral growth, as shown in a recent publication¹⁴. This mouse model of CCG will provide new insight into the process of CCG at cellular and molecular levels and can be used to validate potential targets to promote CCG.

The described animal experiments were conducted in accordance with the Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee of Northeast Ohio Medical University.

1. Surgical preparation

NOTE: For the RI protocol, use C57BL/6 mice of either sex weighing at least 25 g. Use aseptic technique throughout the surgery.

1. Setup
   1. Sterilize all tools in an autoclave or bead sterilizer. Sterilize delicate materials and implants, such as occluder, tether, and PE tubing, with ethylene oxide (EtO).
   2. Clean the surgical area by wiping all areas with 70% ethanol. Prepare the area by laying out all tools and supplies on sterile drapes. See Table of Materials for a complete list of supplies.
2. Mouse intubation
   1. Briefly anesthetize the mouse with 3% isoflurane with oxygen (1 L/min flow rate) until the righting reflex is lost. Shave the chest area, the center of the back, and behind the right ear; completely remove loose hair. Inject glycopyrrolate intramuscularly at 0.01-0.02 mg/kg.
   2. Anesthetize again with 3%-4% isoflurane for 5 min. Place the mouse supine on an incline, restrained from its upper incisors, and use blunted forceps to move the tongue aside. Quickly intubate the mouse with a 20G angiocath using a fiber optic light and a magnifying laryngoscope.
   3. Place the mouse supine on a warming surgical pad and connect the intubation tube to a small animal ventilator with 3% isoflurane. Confirm intubation by checking for rhythmic bilateral chest rise. Confirm adequate depth of anesthesia by lack of toe pinch response.
3. Preparation of the surgical field
   1. Apply ophthalmic ointment to the eyes to prevent drying.
   2. Clean the shaved areas with betadine and then 70% ethanol, wiping once in a unidirectional manner.
   3. Apply electrode cream to the electrocardiogram (EKG) contacts on the surgical pad and tape the mouse's limbs to them. Drape sterile gauze over the lower half of the mouse.
   4. Monitor breathing rate, temperature, and depth of anesthesia over the course of the surgery using the surgical pad's software interface.
4. Left thoracotomy and occluder implantation
   1. Decrease isoflurane to 2%. Make a 2-3 cm midline incision in the skin on the chest using scissors, then use curved forceps to gently separate the skin and muscle layers on the left side of the chest (Figure 1A).
   2. Using blunted forceps, gently make an opening through the thoracic wall to expose the heart, typically between the 3^rd and 5^th intercostals.
   3. Use a retractor to gently spread the ribs and visualize the heart. Using blunted forceps, tear the pericardium to expose the heart. Locate the left auricle (Figure 1B) and visualize the basal part of the left anterior descending artery (LAD), if possible.
   4. Pass an 8-0 polypropylene suture through the myocardium underneath the LAD.
      NOTE: The distance between the entrance and exit should be about equal to the width of the occluder and perpendicular to the LAD.
   5. Remove retractors. Using pointed forceps, exteriorize the end of the occluder tubing through the chest wall in the second inferior intercostal space and pull the occluder tubing out through the chest wall until the occluder is located over the heart in the chest cavity.
   6. Tie the occluder securely to the heart using a surgeon's knot (Figure 1C) so that the occluder lies on the heart but does not press into it.
   7. Briefly inflate the occluder (~10 s) at 10 psi using the push-button inflation device (Figure 2). Check the EKG to confirm ST elevation during inflation and, if possible, visually confirm blanching of the heart apex. Adjust the position or tension of the suture as necessary to confirm ischemia.
5. Closing
   1. Close the flanking ribs with 6-0 polypropylene suture, inserting the chest tube. Evacuate air and blood from the chest, then remove the chest tube. Apply 2% lidocaine to the closed incision in the chest wall.
   2. Using pointed forceps, tunnel the occluder tubing under the skin of the right shoulder to exteriorize the tubing through the skin behind the right ear. Replace chest muscles and close the skin with a 6-0 polyglactin suture (Figure 1D).
   3. Flip the mouse to the prone position. Administer ketoprofen subcutaneously at a dosage of 3 mg/kg (using a 1 mg/mL dilution in 0.9% sterile saline) at least 30 min before the end of surgery.
6. Tether implantation
   1. Using small scissors, make a small (1 cm) incision on the skin at the center of the back and gently separate the skin from the subcutaneous fat and muscle tissue.
   2. Tunnel the occluder tubing to the incision at the center of the back, exteriorize it, and thread it through the tether. Use a 6-0 suture to close the small hole near the ear.
   3. Use 6-0 polypropylene sutures to fix the tether to the back muscles (Figure 1E). Close the skin over the tether button with 6-0 sutures.
7. Recovery
   1. Remove the intubation tube once the mouse can breathe independently; place the mouse in a post-surgical RI cage (for a single animal; Figure 3) once the foot pinch response has returned. Monitor the mouse continuously until it has regained sufficient consciousness to maintain sternal recumbency. Place the cage on a heat pad until the mouse regains full mobility.
   2. The next day, administer a second dose of ketoprofen subcutaneously at 3 mg/kg.
   3. Allow the mouse to rest for 5-7 days before beginning the RI protocol. Monitor mice daily for integrity of instrumentation and change the cages as needed. Provide appropriate enrichment as needed, as mice are single housed for the duration of the RI protocol.

2. Repetitive ischemia

1. Check the placement of the occluder by echocardiography on Day 0 of the RI protocol, as previously described¹⁵. Observe the decrease in cardiac function during inflation of the occluder.
2. Connect the mouse's occluder tubing to the RI inflation system (Figure 4). The system will inflate the occluder to 10 psi for 6 min, 4x a day, with a 3 h break between each inflation (Figure 5).
3. After 17 days of RI, re-check cardiac function as in step 2.1.

3. Polymer perfusion and tissue harvest

1. At the time of sacrifice, anesthetize the mouse with 3%-4% isoflurane and inject heparin (500 U/kg) intraperitoneally. Use a nose cone to continue isoflurane administration at 2% for at least 5 min, then confirm the adequate depth of anesthesia by lack of toe pinch response.
2. Open the chest cavity to expose the heart and thoracic aorta. Cannulate the descending thoracic aorta with PE20 tubing and bisect the inferior vena cava (IVC) to allow outflow; perfuse the heart retrogradely with 1x PBS until the fluid exiting the IVC is clear, followed by 3 mL of 1% lidocaine, then 3 mL of 4% paraformaldehyde (PFA) in PBS. Make a permanent ligation of the LAD at the exact position of the occluder.
3. Retrogradely perfuse the heart with a radiopaque reagent until the arterial circuit is filled. Clamp the PE20 tubing with hemostats and allow the polymer to cure for 90 min. Image the heart under a dissection scope.

Out of 136 C57BL/6 mice, including both males and females, the survival rate of the RI surgery was 93.4%, with 80.9% of mice surviving through the entire 17-day RI protocol.

The mouse RI protocol was optimized based on previous animal RI models^12,16, which have short episodes of ischemia without permanent injury to the myocardium. During the surgery, functional assessment of the occluder can be done by observing visible blanching of th...

Coronary collaterals are a natural bypass for IHD patients. After the failed clinical trials targeting angiogenesis¹⁷, promoting coronary collateral development might be a better therapeutic approach for these patients. Different from capillaries derived from angiogenesis, which have only a single layer of endothelial cells, collaterals are mature arteries with the coverage of smooth muscle cells. Collaterals resupply blood flow to the regions of myocardial ischemia caused by obstructive arteries....

The authors have nothing to disclose.

The authors thank Weiguo Wan, Cody Juguilon, Iyanuoluwa Ogunmiluyi, and Devan Richardson for their contributions to the methods discussed here. This work was supported by 1R15HL115540-01 and 1 R01 HL137008-01A1.