Terminal Harvest Procedure of a Large Animal Model of Chronic Myocardial Ischemia with Hemodynamic Characterization and Perfusion Analysis

Summary

The following surgical protocol represents the second and final procedure to thoroughly and reliably characterize the cardiac function and myocardial perfusion after successful induction of chronic myocardial ischemia in a swine model in an effort to investigate novel therapies.

Abstract

Reproducibility and research integrity are foundational tenets to scientific discovery, which are produced utilizing well-established, proven principles and protocols. Furthermore, with the ever-increasing prevalence and burden cardiovascular disease (CVD) places on individuals and society at large, it deems essential to cultivate robust and validated model for investigation. Our group utilizes a two-surgery protocol in a swine model that has been progressively refined over the last twenty years, in which we first induce chronic myocardial ischemia by placement of an ameroid constrictor mimicking the pathophysiology of coronary artery disease (CAD) in humans. The second and terminal procedure is a comprehensive and thoughtful collection of cardiac hemodynamic parameters as well as regional coronary perfusion, followed by beating heart euthanasia and detailed sectioning of the heart. Our lab uses isotope-labeled microspheres to analyze myocardial blood flow to the ischemic area as a powerful method to quantify collateralization due to experimental therapeutic agents. Furthermore, and likely the most important measurable cardiovascular outcome, we utilize left ventricular pressure-volume (PV) loop catheterization to measure a broad range of hemodynamic parameters. We hope by detailing the comprehensive cardiac analysis of our swine model of chronic myocardial ischemia, other researchers may improve and cultivate their own protocols and therapy investigation for the elimination of cardiovascular disease.

Introduction

In the United States, nearly half of individuals have at least one component of CVD, comprising a large group of disease states of the heart and blood vessels¹. By the year 2060, it is projected that the prevalence of multiple diagnoses comprising CVD will substantially increase compared to 2025. For instance, myocardial infarction is projected to increase by 30%, ischemic heart disease (IHD) by 31.3%, and heart failure by 33%². These staggering statistics are further worrisome when taken with similar predictions of cardiovascular risk factors, in which the number of individuals with diabetes mellitus is estimated to rise by 39.3%, dyslipidemia by 27.5%, and hypertension by 27.2%². Altogether, this highlights the imminent need for reliable and reproducible research models to effectively investigate innovative therapies for CVD and its risk factors. We have demonstrated our physiologically analogous model of CAD in swine³. The use of swine helps fulfill the validity of our research model in accurately representing the human condition of CAD, in which pre-clinical studies have previously been criticized for translatability. Swine have similar coronary anatomy, cardiac physiology, and myocardial proteome and metabolism to humans⁴. Furthermore, a crucial component of our model is to then accurately account for subsequent physiological, myocardial, and molecular outcomes due to the investigational therapies.

Typically, clinical trials are used to implement novel medical or surgical therapies to investigate any beneficial cardiovascular outcomes, which is discovered solely via observation. This is evident in the landmark trials of several novel anti-diabetic drugs investigating their cardiovascular impact. For example, in the SUSTAIN-6 randomized control trial, patients with type 2 diabetes treated with semaglutide, a glucagon-like peptide-1 (GLP-1) agonist, were found to have lower rates of major cardiovascular adverse events compared to placebo⁵. Similar observations were seen in another RCT studying sotagliflozin, a dual sodium-glucose cotransporter 1 and 2 (SGLT-1 and SGLT-2) inhibitor, showing a reduction of major adverse cardiovascular events in patients with type 2 diabetes, chronic kidney disease, and additional cardiovascular risk factors⁶. These findings have been crucial in expanding the FDA approval of such therapies to reduce the incidence of cardiovascular risk factors and disease. However, a major limitation to these observational findings is a lack of understanding of the mechanisms by which these new therapies perpetuate such effects. This exemplifies the importance of our model in the scientific discovery of the underlying mechanistic determinants for new therapies for CVD.

By using a large animal model, we are able to invasively characterize the macroscopic changes in cardiac function and myocardial perfusion, as well as comprehensively understand the microscopic molecular shifts in response to experimental therapies. We employ the use of PV loop catheterization to determine variations in cardiac function. The use of PV loops is the gold standard for invasively determining ventricular function during systole and diastole and serves as a benchmark for comparing non-invasive imaging techniques, such as echocardiogram or cardiac MRI⁷. This technique allows us to take several measurements to determine each hemodynamic variable and re-position the catheter in real time to ensure reproducible data acquisition. That is to say, the utilization of this technique is limited in the inability to collect longitudinal data, such as baseline cardiac functional, cardiac function at ischemic onset, and during treatment. We feel this is offset by the ability to measure load-independent measures, gaining the intrinsic functional status of the heart. Furthermore, the use of isotope-labelled microsphere injections after mapping the most ischemic territory aids in identifying any ischemic benefit of a therapeutic agent. Microsphere injections have long since been proven to be an accurate representation of changes in regional myocardial blood flow⁸.

Our lab consistently uses microspheres with a diameter of 15 μm and at a concentration of 2.5 million microspheres/mL. This diameter ensures the most precise measurement of myocardial blood flow, as confirmed by a previous study showing that microspheres were properly distributed according to blood flow, and the smaller diameter exhibited the least variation in distribution among left ventricular layers⁹. Additionally, a minimum amount of uptake of microspheres is necessary to ensure reliable results, which is easily produced at such high concentrations used in our lab. Taken together, the techniques employed during the terminal harvest consistently and meticulously analyze key cardiac physiological changes after any number of experimental therapies. This characterization is further bolstered by the ability to collect myocardial tissue and thoroughly analyze regional changes with advanced technologies, such as multiomic studies, providing robust, quantitative characterization at each layer of the central dogma within the myocardium.

Protocol

The following surgical procedure was approved by the Institutional Animal Care and Use Committee of Rhode Island Hospital and Brown University (Protocol #23-08-0005).

1. Animal model

1. Choose Yorkshire swine that have previously undergone placement of an ameroid constrictor around the proximal left circumflex coronary artery (LCx), thereby inducing chronic myocardial ischemia, and subsequent treatment with an experimental therapeutic agent. Keep ~7 weeks between the first operation for ameroid placement and subsequent terminal harvest. Allow 2 weeks for ameroid closure and ensure the swine undergo an experimental therapy afterwards. Ensure that the swine are ~18 weeks old and weigh ~40 kg at the time of this terminal harvest and that the experimental and control groups are matched by sex.
   NOTE: A representative figure demonstrates the anatomy of the heart we are harvesting, which includes where the ameroid was previously placed and the expected area of ischemic myocardium (Figure 1).

2. Presurgical procedure

1. Withhold food on the morning of the procedure.
2. Anesthetize the swine using intramuscular injections of telazol (4.4 mg/kg) and xylazine (2.2 mg/kg) and intubate them with a cuffed endotracheal tube.
3. Initiate analgesia using 0.03 mg/kg of buprenorphine.
4. Place an intravenous (IV) catheter (20-22 G) in one of the large auricular veins and start infusing 0.9% saline at a rate of ~5-10 cc∙kg^-1∙h^-1.
   NOTE: The IV fluid is solely at a maintenance rate in the context of the swine's NPO status and length of the surgical case.
5. Shave the surgical site and other sites used for ECG monitoring.
6. Transport the swine to the operating room table, positioned supine with the legs strapped to the table, and continue anesthesia using isoflurane (0.75-3.0%).
7. Start ECG and blood pressure monitoring and apply an electrocautery grounding pad on the swine.
8. Cleanse the surgical site.
   NOTE: This is a terminal procedure; therefore, sterility does not need to be maintained. It is individual preference with respect to surgical site preparation and draping.

3. Surgical procedure

1. Femoral catheterization
   1. Palpate for the femoral pulse.
   2. Inject 2% lidocaine subcutaneously to prepare for a femoral groin cut down in either the left or right groin, roughly perpendicular to the inguinal crease, moving craniocaudally.
   3. Make a 4-5 cm incision using a #10 scalpel, again staying perpendicular to the inguinal crease, starting caudally where the femoral pulse was palpated and moving cranially. (Figure 2A)
   4. Using electrocautery, dissect through the subcutaneous tissue down to the level of the femoral artery. Use a Weitlaner retractor to spread the muscle further exposing the femoral artery.
      NOTE: This may require a small amount of splitting the semimembranosus and vastus medialis muscle.
   5. Bluntly dissect using a right-angle clamp to circumferentially expose the femoral artery from surrounding tissue. Use Metzenbaum scissors to sharply divide any adhesions that cannot be taken down bluntly.
   6. Isolate the femoral artery by encircling the artery with a vessel loop.
   7. Place a second vessel loop to ensure proximal and distal control of the artery.
   8. Systemically heparinize the swine before cannulation at a dose of 80 IU/kg.
      NOTE: Make sure the median sternotomy and cardiac exposure are complete before heparinizing and achieving femoral access. We make sure cardiac exposure is complete before heparinization because any lysis of adhesions can create bleeding. The overall purpose of heparinization is for thrombus prevention when accessing both the femoral artery and the left ventricle.
   9. Using the Seldinger technique, puncture the femoral artery with an 18 G access needle, avoiding piercing the posterior arterial wall. When blood returns, advance a guidewire through the needle into the artery. Remove the needle and guide the 7F arterial sheath into the artery. Remove the guidewire and vascular dilator, leaving only the arterial sheath in place.
      NOTE: The femoral artery can be difficult to access as it can easily dissect. Given this is a terminal procedure, it is recommended to sharply cut the femoral artery with Metzenbaum scissors and directly insert the guidewire into the lumen if this occurs. The arterial sheath can then easily be passed into the vessel over the guidewire. Make sure to clamp off the distal portion of the femoral artery using a small hemostat.
   10. Ensure blood flow via the sheath. Connect to the Harvard Apparatus withdrawal pump (Figure 3).
2. Median sternotomy
   1. Depending on the number of co-surgeons, this portion can be completed simultaneously with the groin cut down.
   2. Inject 2% lidocaine subcutaneously to prepare for midline sternotomy.
   3. Make a midline incision from the sternal notch to the xiphoid process using a #10 scalpel. (Figure 2B)
   4. Using electrocautery, divide the subcutaneous tissue until the sternum is reached and maintain hemostasis.
   5. Excise the xiphoid process by freeing any adhesions laterally and inferiorly with electrocautery. Use a Kelley clamp to lift the xiphoid up and transect at its base horizontally using electrocautery.
   6. Ensure the sternal notch is dissected free superiorly to ease dividing this portion of the sternum. This typically requires dividing some superficial strap muscles.
   7. Bluntly dissect underneath the sternum using a finger.
   8. Using a Lebsche knife, sharply divide the sternum, ensuring to stay midline. Change the angle of the knife as necessary and continue slowly once the latter one-third of the sternum is reached.
      NOTE: The use of the knife is preferred over a sternal saw, as in humans, due to the steep angulation and thickness of the cranial portion of the sternum, ideally reducing the risk of damage to underlying great vessels. It can be useful to place gauze underneath the cranial portion of the sternum to protect the underlying great vessels.
   9. Once the sternotomy is complete, bluntly and sharply lyse any adhesions, if present, between the chest wall and pericardium or pleura.
   10. Place a sternal retractor to spread the ribs and expose the heart.
   11. Open the pericardium using Metzenbaum scissors. Extend towards the right atrium and continue caudally towards the left ventricular apex.
   12. Continue to lyse adhesions either bluntly or sharply to expose the left and right atrial appendages. The use of peanut sponge on an Allis clamp can be helpful for this dissection.
   13. If it is difficult to dissect and identify the atrial appendages, a thymectomy may be helpful. To perform this, use electrocautery to dissect out thymic tissue.
   14. Once the atrial appendages are exposed, apply pacing clamps to both appendages, which are connected to a single-chamber external pacing box. Test pacing to 150 bpm, ensuring that each beat is captured on ECG.
   15. Ensure exposure of the left ventricular apex, again using blunt and sharp dissection.
   16. Lastly, enter the left pleural space and encircle the inferior vena cava (IVC) with a large Satinsky clamp. Isolate with a vessel loop.
   17. Once this is complete, it is an appropriate time to heparinize the swine and obtain femoral access.
3. Microsphere injection-perfusion analysis
   1. Access the left atrial appendage with a fine-gauge butterfly needle attached to IV tubing and a three-way stopcock. Use a small hemostat to guide the needle and hold in place.
   2. Connect a syringe to the stopcock with sterile heparinized saline and withdraw back to ensure the correct positioning. If blood is easily withdrawn, the position is correct.
   3. Connect a syringe with 5 mL of lutetium-labeled microspheres (concentration: 2.5 million microspheres/mL) to the three-way stopcock.
   4. Assign an operating room assistant to time for 90 s.
   5. Begin femoral artery blood withdrawal via the Harvard Apparatus pump for a total of 10 mL of blood at a rate of 6.67 mL/min and begin timing.
   6. Over the first 30 s of blood withdrawal, inject the lutetium microspheres into the left atrial appendage (Figure 4). Halfway through, confirm positioning by withdrawing on the syringe containing microspheres.
   7. Once this is complete, flush the tubing with a syringe of heparinized saline and again ensure correct positioning.
   8. After 90 s, disconnect the tubing from the femoral access to the withdrawal pump and reconnect the new tubing to obtain analysis during pacing.
   9. Begin pacing to 150 bpm.
   10. Repeat steps 3.3.5, 3.3.6, and 3.3.7 using 5 mL of samarium-labelled microspheres (concentration: 2.5 × 10⁶ microspheres/mL).
   11. Stop pacing and disconnect the femoral arterial sheath from the Harvard Apparatus withdrawal pump.
   12. Remove the butterfly needle and apply pressure to ensure hemostasis of the left atrial appendage.
4. Pressure-volume loop catheterization
   1. Place a purse string suture at the left ventricular apex with a 4-0 polypropylene.
   2. In the middle of the area encompassed by the suture, access and place another arterial sheath in the left ventricle using the Seldinger technique described above.
   3. Insert a calibrated micromanometer-tipped pressure-volume transduction catheter into the left ventricle via the sheath. Pull the arterial sheath out of the left ventricle over the catheter and begin transducing the pressure waveform using the Pressure-Volume Measurement System. This will inform the location of the catheter, for example, if the catheter is in the aorta, valve complex, or the left ventricle. Draw back on the catheter as necessary to ensure it is in the left ventricular cavity and reposition it to ensure the pressure-volume loop morphology appears appropriate (Figure 5).
      NOTE: Removing the sheath from the left ventricle allows for better transduction of the pressure-volume tracing from the catheter and allows more ease in changing the catheter's position to obtain the most appropriate morphology of each PV loop.
   4. Place another calibrated PV catheter in the femoral access to obtain systolic and diastolic blood pressure and mean arterial pressure.
   5. Once satisfied with the morphology of the PV loops, perform three breath holds at rest to reduce respiratory variation on the pressure-volume loops. This will measure several load-dependent variables, such as cardiac output and stroke volume.
   6. Following that, occlude the IVC by gently retracting the vessel loop and perform a simultaneous breath hold to obtain load-independent variables, such as end-systolic pressure volume relationship (ESPVR) and end-diastolic pressure volume relationship (EDPVR). Repeat two more times.
   7. Begin pacing to 150 bpm and obtain three breath holds to characterize cardiac performance under stress conditions.
   8. This completes the profiling of cardiac function. Remove both catheters and tie down the purse string suture on the left ventricle.
5. Beating euthanasia
   1. Remove the pacing leads and all instruments from the chest.
   2. Obtain suction for the co-surgeon.
   3. Bluntly dissect any lateral adhesions of the heart to the chest wall and ensure the left ventricular apex can be easily lifted superiorly.
   4. Ask anesthesia to deepen sedation.
   5. Place a large Kelley clamp on the superior vena cava (SVC), if this is easily accessible, and a large Satinsky clamp around the IVC. Clamp at the same time.
   6. Immediately take a #10 scalpel to remove the heart from the chest. This includes separating the heart from the IVC, SVC, aorta, pulmonary arteries, and pulmonary veins.
      NOTE: The co-surgeon should provide suction to keep the surgical field as clear as possible during this time.
   7. Deliver the heart from the chest and place on a separate table.
   8. Cut in two halves: apical and basal (Figure 6A). This will separate the ventricular tissue from the atria and great vessels.
   9. Separate the left ventricle from all other myocardial tissue.
   10. Each half should be circumferentially sectioned into twelve sections based on proximity to the left anterior descending coronary artery (LAD) and the posterior descending coronary artery, as depicted in Figure 6B.
   11. Collect tissue samples from all other chambers, valves, great vessels, and peripheral tissue, such as the pancreas, liver, lung, kidney, etc.
   12. A small sample from each left ventricular section should be allocated to BioPAL for microsphere quantification. Certain sections are fixed in formalin for histological analysis, while a majority of tissue is snap frozen in liquid nitrogen for future molecular studies.

Results

First, by injecting isotope-labelled microspheres into the left atrial appendage, regional myocardial perfusion is measured. To calculate the myocardial blood flow, when a constant rate of blood withdrawal of 6.67 mL/min was utilized, the following equation can be applied: myocardial blood flow = (reference blood withdrawal [6.67 mL/min]/weight of tissue) (quantified tissue microsphere count/quantified blood reference microsphere count). Simply put, the ratio of measured microspheres in a...

Discussion

This surgical procedure has four critical steps: exposure of the heart, femoral catheterization, microsphere injection for perfusional analysis, and pressure-volume loop acquisition. In regard to exposing the heart, an important consideration should be the possibility of encountering significant adhesions due to the first surgical procedure. When performing the median sternotomy, the pericardium may be adhered to the chest wall, which should be bluntly swept before starting the sternotomy. When approaching the latter one...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
#10 Scalpel	Medline	MDS15310
0.9% Saline	Dechra/Vetivex	17033-492-50	Rate: 5-10 mL/kg during case
18G Needles	Patterson Veterinary	78032793	For drawing medication/microspheres
2% Lidocaine	Phoenix	57319-533-05	Dosing: 2-5mg/kg
4-0 Polypropylene Suture	Covidien	D9H0436Y	For purse string
All Surgical Instruments	Steris and Medline	Requires Sales Representative	All Commonly Found in Hospital Operating Rooms
Analgesia: Buprenorphine	Hospira	Brand Name: Bupredine	Dosing: 0.03 mg/kg
Anesthesia: Telazol	Zoetis	(tiletamine and zolazepam)	Dosing: 4.4 mg/kg
Anesthesia: xylazine (Rompun)	Dechra	N/A	Dosing: 2.2 mg/kg
Arterial Sheath	Cordis	401-711M	Either 6F or 7F; requires two
Butterfly needle infusion set	Surflo	SV-25BLK	For microsphere injection; typically 23-25G
Chlorhexidine 4%	BD E-Z Scrub	371073	Surgical Prep, if preferred
Cuffed Endotracheal Tube	RUSCH	7-0 typically	May need different sizes depending on swine size and weight
ECG Monitoring System	MidMark	N/A	Multiparameter Monitor
Electrocautery	Bovie Specialist Pro	1250S	Typically set up 2 if possible
Isoflurane, USP	Dechra		Dosing: 0.75%-3.0%
Laptop	HP Microsoft	N/A	Can use any laptop preferred by individual
Lebsche Knife	Integra	300-425	N/A
Lutetium STERIspheres	BioPAL, Inc.	C-15E20	Amount: 5cc per surgery
Mini White Vessel Loops (X-ray Detectable)	DeRoyal	30-711	To isolate the femoral artery and IVC
Multiple 5cc Syringes	Terumo	SS-05L	For flushes and microsphere injections
Pacemaker	Osypka Medical	2428001	A temporary pacer
PV Loop Catheters	Transonic	Model: FDH-5018B-E3450 and S/N: 50-679	For PV Loop Acquisition
PV Loop Computer Program	Transonic Scisense	ADV500 Software	Program on your laptop
PV Loop Data Acquisition System	Transonic and ADInstruments	Models: FY097B and PowerLab 8/35	Connects the catheters and the computer
Samarium STERIspheres	BioPAL, Inc.	C-15A20	Amount: 5cc per surgery
Small Mallet	Integra	410-335	N/A
Sterile Gloves	Encore	820955	Would still recommend sterile gloves and gown
Sterile Saline	BAXTER	G171328	Dilute to make heparanized saline flushes
Suction System	GOMCO	4042	Better if Wall Suction of Own Facility
Suction tip: Yankauer	Medline	DYND50130	Important to keep the field clear when resecting the heart
Systemic Heparin	Covetrus	82496	Dosage: 80 IU/kg
Three-way stopcock	Jorgensen Laboratories, Inc.	200820	For microsphere injections
Warm Water Bath	Fisher		To warm the catheters before use
Withdrawal Pump	Harvard Apparatus	70-2002	To withdraw peripheral blood at constant rate
Yorkshire Swine	CBSET, Inc.	N/A	Swine will have already undergone the initial surgical procedure