Intramural funding was provided for this project by the Division of Intramural Research, National Heart, Lung and Blood Institute at the National Institutes of Health. We would like to acknowledge Dr. James Hawkins, Dr. Robert Hoyt, The NIH Division of Veterinary Resources and the staff of the Cardiothoracic Surgery Research program at the NIH.

The procedures described here were approved by the National Heart, Lung, and Blood Institute Animal Care and Use Committee at the National Institutes of Health and conform to the policies outlined in The Public Health Service Policy on Humane Care and Use of Laboratory Animals, The Animal Welfare Act, and the Guide for the Care and Use of Laboratory Animals.
NOTE: The purpose of this surgical procedure is to produce an animal model of chronic myocardial ischemia that can be used to develop treatments that are clinically relevant for patients with severe CAD. This cannot be accomplished using an in vitro model.
1. Animals
<ol>
	<li>Use male Yorkshire swine weighing between 12 and 15 kg.</li>
</ol>
2. Pre-surgical Procedures
<ol>
	<li>For a prophylactic antibiotic, use amoxicillin and clavulanate potassium (15 mg/kg) orally twice a day beginning 24 h prior to surgery. 
	NOTE: This oral antibiotic is started before surgery to make sure that the pig will swallow it and to also make sure that they will not have an allergic reaction to the medication post-operatively.</li>
	<li>Withhold food and water from the pig 12 h prior to surgery.</li>
	<li>On the day of surgery, anesthetize the pig with a cocktail of ketamine (33 mg/kg), midazolam (0.5 - 0.75 mg/kg), and glycopyrrolate (0.01 mg/kg) given intramuscular (IM).</li>
	<li>Place a 20 G intravenous (IV) catheter in an ear vein and intubate the animal with a 7.0 Fr endotracheal tube. 
	NOTE: Adjust the size of the IV catheter and the endotracheal tube as appropriate for the size of the pig.</li>
	<li>Remove the hair (shave) on the left side of the pig from the shoulder to the umbilicus and from the abdominal midline to the vertebral column.</li>
	<li>Administer the prophylactic antibiotics (piperacillin and tazobactam: 100 mg/kg IV) and pain medication (buprenorphine sustained release: 0.2 mg/kg subcutaneous (SC) or a fentanyl transdermal patch 25-50 &#181;g/h). 
	NOTE: The IV antibiotic is given because the animals are fasted for 12 h prior to surgery and do not receive the previously mentioned oral antibiotic during this time.</li>
	<li>Transport the pig to the operating room and connect the endotracheal tube to the anesthesia machine equipped with a sidestream sensor to monitor the airway gases. Mechanically ventilate the pig using a tidal volume of 10 mL/kg as a starting point and do not exceed an airway pressure of 30 cm H2O. Set the respiratory rate between 10-20 breaths per min (bpm) and adjust as needed to keep the end-tidal CO2 (PetCO2) levels between 28 and 35 mm Hg. Maintain the anesthesia with either isoflurane (1-3%) or sevoflurane (2-5%).</li>
	<li>Position the pig right-side down on the operating table. Attach all necessary probes to monitor body temperature (BT), electrocardiogram (ECG), pulse oximetry (SpO2), etc.</li>
	<li>Prepare the surgical site via three alternating scrubs with 2% chlorohexidine and 70% alcohol (apply in a circular motion beginning with center and moving outward).</li>
	<li>Drape the surgical site using aseptic technique21.</li>
</ol>
3. Surgical Procedure
<ol>
	<li>Expose the heart through a left thoracotomy.
	<ol>
		<li>Make a skin incision parallel to the 4th and 5th intercostal space about 8 - 9 cm in length using a no. 10 scalpel blade.</li>
		<li>Cut through the latissimus dorsi and serratus ventralis muscles using a pair of curved Metzenbaum scissors and Brown Adson forceps. Use electrocautery to maintain hemostasis as necessary.</li>
		<li>Enter the pleural cavity through the intercostal muscles between the 4th and 5th ribs with the Metzenbaum scissors cutting along the anterior aspect of the 4th rib. To reduce the chance of accidentally damaging the lung, turn the ventilator off on the exhale just prior to entering the pleural cavity. Once the pleural cavity has been breached, turn the ventilator back on.</li>
		<li>Use a small Finochietto retractor to spread the ribs apart, exposing the heart.</li>
	</ol>
	</li>
	<li>Use Debakey forceps to grasp and lift the pericardium. Use Potts scissors to make a small hole in the pericardium, allowing air to enter the pericardial space. Continue the incision with the Potts scissors over the junction of the LAD and LCX arteries.</li>
	<li>Use Babcock forceps to retract the left atrial appendage. Using Debakey and small right angle forceps, dissect the LCX artery from the surrounding tissue prior to or proximal to the 1st&#160;obtuse marginal branch.</li>
	<li>Place two vessel loops under the dissected LCX artery, one at each end. Hold a size 3.0 mm ameroid constrictor with the small right angle forceps and lift the vessel loops to gently guide the LCX artery through the opening of the constrictor. Gently rotate the constrictor so the opening is facing up. Remove the vessel loops. See Figure 1. 
	NOTE: Adequate dissection of the artery is essential to prevent kinking. Select the appropriate sized constrictor for the individual animal. The appropriate sized constrictor will closely encircle the vessel without initially constricting the vessel. Domestic pigs weighing between 12 and 15 kg require ameroid constrictors sized 2.5 - 3.5 mm. The location of the ameroid constrictor on the vessel depends on the desired size of the ischemic area.</li>
	<li>Re-approximate the pericardium and close with 4-0 polypropylene sutures.</li>
	<li>Reestablish the negative thoracic pressure by inserting a 12 Fr chest tube or equivalent in the pleural space with the end exiting out between the layers of closure, generally two intercostal spaces posterior to thoracotomy site. 
	NOTE: Observe the lung for atelectasis before closing the ribs. If any signs of underinflation are observed, re-inflate the lungs manually up to a pressure of 30 mm Hg.</li>
	<li>Close the chest using 1 polypropylene sutures for the intercostal layer, 0 polypropylene sutures for the serratus and fascial layer, 2-0 polypropylene sutures for the subcutaneous layer, and 3-0 polypropylene sutures for the subcuticular layer. Staple or suture the skin.</li>
	<li>Attach a three-way stop cock to the end of the chest tube and evacuate the air in the chest with a 40-60 cc syringe until a negative seal is achieved. Roll the animal onto its sternum or opposite side to facilitate removing all the air. Once the chest maintains negative pressure, remove the chest tube and suture the exit site. If animals are housed singly post-operatively, the incision can be bandaged for several days.</li>
</ol>
4. Post-operative
<ol>
	<li>Following closure, infiltrate the incision along both sides of the thoracotomy site with 0.25% bupivacaine.</li>
	<li>Wean the animal off the ventilator and turn off the anesthesia. Remove the endotracheal tube once the animal is breathing on its own and swallowing.</li>
	<li>Monitor the animal until it is fully recovered (awake and sternal). There should be no abnormalities or arrhythmias on the ECG, the BT should be 38.7 - 39.8 &#176;C, the SpO2 should be 95 - 100%, and the respiratory rate should be 32 - 58 bpm.</li>
	<li>Administer amiodarone (100 mg, orally twice a day) and clopidogrel bisulfate (75 mg, orally once a day) to prevent thrombosis and arrhythmias. Continue with amoxicillin and clavulanate potassium (15 mg/kg) orally twice a day for 10 days post-operation. Post-operative pain is controlled with buprenorphine SR &#160;(sustained release) 0.2 mg/kg. SQ every three days. &#160;The buprenorphine SR may be supplemented with carprofen (4.4 mg/kg) once and &#160;then (2.2 mg/kg) PO or IM BID.</li>
	<li>Keep all bedding and foraging material out of the animal&#39;s housing area until the incision is fully healed.</li>
</ol>

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The authors have no conflict of interest to disclose.

Ameroid constrictors have been used extensively to create animal models of chronic myocardial ischemia, chronic myocardial infarction, and heart failure5,6,7,9,10,11,12,13,15,16,17,18,19,22. Although these models can be created through the use of stents, hydraulic occluders, or fixed stenosis occluders, the prevalence of ameroid use in scientific literature allows researchers to more accurately compare the results of their work with previously published studies5,6,7,9,10,11,12,13,15,16,17,18,19,22. Another advantage of using the ameroid constrictor is that the procedure is relatively straightforward and can be performed successfully by anyone with reasonable surgical skills, and requires no specialized instruments. The other aforementioned methods require greater technical skills and, in the case of the hydraulic occluder, requires intense post-operative monitoring due to the externalized device6,13.
The main limitation of using the ameroid constrictor is the variability in the rate of occlusion9,13. In general, most studies using the ameroid constrictor have found the rate of stenosis to be highest in the first two weeks and then gradually taper off until complete occlusion occurs23,24. Ex vivo studies directly measuring the closure rate of the ameroid lumen have confirmed that the greatest reduction in lumen diameter occurs in the first two weeks and slows thereafter25,26. However, these same studies have also showed that surrounding glucose and protein concentrations can affect the rate and completeness of the ameroid closure, thereby suggesting that in vivo conditions are most likely responsible for the variability in ameroid occlusion rates25,26. Other investigators have suggested that mechanical trauma, inflammation, fibrosis, and thrombus formation caused by the ameroid placement procedure itself may contribute to this variability5,6,25. The latter scenario is most likely the reason for the premature deaths and heart failures observed in this study. It cannot be stressed enough the importance of doing delicate, yet adequate, artery dissection on either side of the ameroid to prevent kinking of the vessel.
Proper placement of the ameroid constrictor is critical to develop an ischemic area of the heart that is large enough to treat but not so large that it results in death. Placing the ameroid constrictor on the LCX before the first obtuse marginal branch appears to work best in Yorkshire pigs. As reported elsewhere, this resulted in an ischemic area of around 25-30% of the left ventricle6. Placing the ameroid any lower resulted in an ischemic area that was inadequate for the specific treatment that was being investigated in this study.
Arrhythmias occurring during the surgical procedure are rare. Administering oral amiodarone post-operation through the end of the study, may prevent arrhythmias during the 2 to 3 weeks post-operation, when the ameroid is closing off. Additionally, caretaking and medical staff should be especially vigilant in noting any coughing during this time, which could be an early sign of pulmonary edema/heart failure.
Over the years, the design of the ameroid constrictor has been improved upon. Ameroids are currently available in numerous sizes allowing the surgeon to select an ameroid that matches the diameter of the vessel to be occluded. The outer ring of the ameroid is available in stainless steel, titanium, or plastic. This is especially important when evaluating the affected hearts using MRI. All metal will cause an artifact on the MRI images. Titanium ameroids cause less of an artifact than stainless steel, while plastic leaves none at all. MRI images showing the difference between a titanium ameroid and a plastic ameroid are shown in Figure 2 and Figure 3, respectively. The artifact on the right side of the heart in Figure 2 is caused by the titanium ring encasing the ameroid. Until recently, the containment ring on the plastic ameroid was too bulky to place around a coronary artery without kinking the vessel. The ameroid constrictor used in this study (see Table of Materials) is a much more streamlined version, which has the same outer diameter as the metal ameroids.

Coronary artery disease (CAD), which leads to myocardial ischemia, is the leading cause of disability, death, and health care costs around the world4 and attributable for approximately 1 out of every 3 deaths in the U.S.1,2,3,4,5,6. Although there have been many advances in both percutaneous and surgical treatments, as many as one third of patients suffering from CAD are ineligible for these treatments due to age, poor health, or suboptimal anatomy4,5,6,7. In order to evaluate new methods of diagnostic imaging or treatment, the development of an appropriate animal model is critical.
When developing an animal model for a disease, the induced disorder should closely emulate the anatomical and physiological characteristics of the disorder in humans8,9. One of the most widely used large animals for cardiovascular studies is the pig. The pig heart is most closely analogous to the human heart in terms of size, anatomy, and physiology3,6. Similar to the human heart, the myocardium of the pig&#39;s heart does not possess an extensive collateral circulation6. Because of this, the pig heart does not tolerate acute coronary occlusion well, but it can tolerate a gradual coronary occlusion. If a coronary artery is slowly occluded, it can be used as a model for chronic myocardial ischemia, chronic myocardial infarction, and heart failure5,6,9,10,11,12,13. Chronic myocardial ischemia can be induced through stent implantation or the placement of a hydraulic occluder, a fixed stenosis occluder, or an ameroid constrictor. There are advantages and disadvantages for all these methods which are outlined in detail in various publications6,9,13 but the most commonly used method is the ameroid placement5,6,10,11.
The ameroid constrictor consists of casein material encased within a stainless steel, plastic, or titanium ring. Once placed around an artery (usually the left anterior descending coronary artery (LAD) or the left circumflex coronary artery (LCX)), the casein material absorbs the surrounding fluid causing the inner lumen to narrow gradually, mimicking the slow stenosis of an artery and resulting finally in full occlusion9,13,14. This procedure used by itself or in conjunction with other methods results in a region of chronic myocardial ischemia and/or infarction of the left ventricle of the heart that has been useful for developing and evaluating new imaging techniques8,10,15, therapeutic treatments7,16,17,18 and surgical procedures19,20.

<table border="0" cellpadding="0" cellspacing="0" width="965">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:323px;">Anesthesia: ketamine</td>
			<td style="width:195px;">Putney, INC</td>
			<td style="width:175px;"></td>
			<td style="width:273px;">Dose: 25mg/kg, IM</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Anesthesia: midazolam</td>
			<td>App Pharmaceuticals</td>
			<td></td>
			<td>Dose: 0.75mg/kg, IM</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Anesthesia: isoflurane</td>
			<td>Baxter</td>
			<td></td>
			<td>Dose: 1-3%, INH</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Antibiotics: amoxicillin and clavulanate potassium</td>
			<td>WG Critical Care</td>
			<td></td>
			<td>Dose: 15mg/kg, IV</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Antibiotics: piperacillin and tazobactam</td>
			<td>Fresenius Kabl</td>
			<td></td>
			<td>Dose: 100mg/kg, PO</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Analgesia: buprenorphine (sustained release)</td>
			<td>Zoo Pharm</td>
			<td></td>
			<td>10mg/ml bottle, Dose: 0.2mg/kg SC</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Analgesia: fentanyl patch</td>
			<td>Mylan</td>
			<td></td>
			<td>Dose: 25-50mcg/hr, TD</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Analgesia: bupivicaine 0.25%</td>
			<td>Hospira</td>
			<td></td>
			<td>Give SC at incision site for analgesia</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Alcohol 70%</td>
			<td>Humco</td>
			<td>NDC 0395-4202-28</td>
			<td>for scrubbing surgical site</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Chlorhexidine scrub 2%</td>
			<td>Vet One</td>
			<td>510083</td>
			<td>for scrubbing surgical site</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Datex-Ohmeda Aestiva /5 anesthesia machine</td>
			<td>GE Healthcare Madison WI.</td>
			<td style="width:175px;"></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Pediatric anesthesia circuit</td>
			<td>Westmed</td>
			<td>7-8901</td>
			<td>www.westmedinc.com</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Water blanket</td>
			<td>Cincinnati Sub Zero</td>
			<td>Blanketrol 2</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">EKG</td>
			<td>Medtronics- Physiocontrol</td>
			<td>LifePak 20</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Oxygen saturation monitor</td>
			<td>GE Healthcare Madison WI.</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Temperature probe</td>
			<td>GE Healthcare Madison WI.</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Electrical cautery unit</td>
			<td>Valley Labs</td>
			<td>Force 2</td>
			<td>Cut-30 Coag-40</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Endotracheal tube</td>
			<td>Hudson RCI</td>
			<td>HUD510312</td>
			<td>Hudson brand - appropriate size for animal</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Intravenous catheter, size 20 gauge</td>
			<td>Santa Cruz Biotechnology, Inc</td>
			<td>SC-360097</td>
			<td>Fluid and drug administration</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Lactated Ringers Solution</td>
			<td>Hospira</td>
			<td>NDC 0409-7953-03</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Macro IV drip set 15 drops/ml</td>
			<td>Hospira</td>
			<td>12672-28</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Surgical gowns</td>
			<td>Kimberly Clark</td>
			<td>90142</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Utility drapes</td>
			<td>Kimberly Clark</td>
			<td>89731</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Adult laparotomy drape</td>
			<td>Medline</td>
			<td>DYNJP3004</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sterile surgical gloves</td>
			<td>Cardinal Health (Allegiance)</td>
			<td>22537-570</td>
			<td>Size 7 - use appropriate size for surgeon</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Needle mat - sterile</td>
			<td>Medline</td>
			<td>NC30MBR</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cautery pencil</td>
			<td>Medline</td>
			<td>ESPB 2000</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Suction tubing</td>
			<td>Medline</td>
			<td>DYND50251</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Suction tip; Yankauer</td>
			<td>Medline</td>
			<td>DYND50130</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Suction cannister</td>
			<td>Cardinal Health (Allegiance)</td>
			<td>65651-220</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Ameroid constrictors sizes 2.5mm -3.5mm</td>
			<td>Research Instruments SW, Inc.</td>
			<td>760-764-9411</td>
			<td>Types: Stainless steel, titanium, or plastic</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">3-way stopcock</td>
			<td>Cardinal Health (Allegiance)</td>
			<td>455991</td>
			<td>For chest tube</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Stomach tube - 18 Fr</td>
			<td>Cardinal Health (Allegiance)</td>
			<td>DC4418</td>
			<td>For chest tube</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">60cc syringe</td>
			<td>Cardinal Health (Allegiance)</td>
			<td>SY35060LL</td>
			<td>For chest tube</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">3cc syringe</td>
			<td>Cardinal Health (Allegiance)</td>
			<td>SY35003LL</td>
			<td>For post-op analgesia injection</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Skin stapler - 35W</td>
			<td>Ethicon&#160; Endo-Surgery</td>
			<td>PMW 35</td>
			<td>skin closure</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Scalpel blade - size #10</td>
			<td>Cardinal Health (Allegiance)</td>
			<td>32295-010</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mini silicone vessel loops&#160; 1 Pak</td>
			<td>Aspen Surgical</td>
			<td>011001PBX</td>
			<td>for elevating circumflex artery to apply ameroid</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">1 (polypropylene) taper</td>
			<td>Ethicon</td>
			<td>BB #8425H</td>
			<td>close ribs</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">2-0 (polypropylene) taper</td>
			<td>Ethicon</td>
			<td>SH #8523</td>
			<td>close muscle layers</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">4-0 (polypropylene) taper</td>
			<td>Ethicon</td>
			<td>BB #8581H</td>
			<td>close pericardium</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">20 gauge needle</td>
			<td>Cardinal Health (Allegiance)</td>
			<td>81-200219</td>
			<td>post-op analgesia injection</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">antimicrobial drape</td>
			<td>3M</td>
			<td>6640EZ</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Finochietto rib retractor - infant size</td>
			<td>Codman</td>
			<td>50-8057</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Potts-Smith scissors</td>
			<td>Roboz</td>
			<td>RS-6043</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Babcock forceps</td>
			<td>Roboz</td>
			<td>RS-8022</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Pean hemostatic forceps curved</td>
			<td>Codman</td>
			<td>30-4565</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">scalpel handle #7</td>
			<td>Codman</td>
			<td>11-5534</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Brown Adson forceps</td>
			<td>Roboz</td>
			<td>RS-5231</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Debakey forceps</td>
			<td>Roboz</td>
			<td>RS-7562</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mayo scissors</td>
			<td>Roboz</td>
			<td>RS-6870SC</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Metzenbaum dissecting scissors</td>
			<td>Codman</td>
			<td>36-5011</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Metzenbaum dissecting scissors</td>
			<td>Codman</td>
			<td>36-5016</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mayo-Hegar needle holder</td>
			<td>Codman</td>
			<td>36-2017</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Ryder needle holder</td>
			<td>Codman</td>
			<td>36-3012</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Kelly hemostatic forceps</td>
			<td>Roboz</td>
			<td>RS-7130</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Reynolds hemostatic forcep</td>
			<td>Roboz</td>
			<td>RS-7211</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Weitlander retractor</td>
			<td>Roboz</td>
			<td>RS-8612</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Yorkshire domestic pigs</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

a chronic cardiac ischemia model in swine using an ameroid constrictor

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but regardless of the approach, an in vivo model is needed to test each treatment. The pig is one of the most used large animal models for cardiovascular disease. Its heart is very similar in anatomy and function to that of a human. The ameroid placement technique creates an ischemic area of the heart, which has many useful applications in studying myocardial infarction. This model has been used for surgical research, pharmaceutical studies, imaging techniques, and cell therapies.
There are several ways of inducing an ischemic area in the heart. Each has its advantages and disadvantages, but the placement of an ameroid constrictor remains the most widely used technique. The main advantages to using the ameroid are its prevalence in existing research, its availability in various sizes to accommodate the anatomy and size of the vessel to be constricted, the surgery is a relatively simple procedure, and the post-operative monitoring is minimal, since there are no external devices to maintain. This paper provides a detailed overview of the proper technique for the placement of the ameroid constrictor.

After analyzing the data obtained from ameroid placement surgeries performed in our facility over a two-year period, we found the survival rate to be 80%. The procedure was performed on 25 Yorkshire pigs weighing between 12 - 15 kg. Of the 25 pigs, 20 survived to the follow-up procedure, 2 fibrillated and died shortly after closing, 2 were euthanized for severe heart failure and pulmonary edema, and 1 had an anesthetic death during a follow-up radiograph. Necropsy revealed an infarcted area of the left ventricle in the animals that died within 24 hours of the procedure. It is suspected but could not be confirmed that the artery kinked due to the presence of the ameroid since the lumen of the ameroid was still open. The ameroids obtained from the animals that were euthanized due to heart failure were examined. There was total closure of the lumen at 18 days.
The surviving animals were imaged by Magnetic Resonance Imaging (MRI) at 28 days post-ameroid placement to measure the heart function and size of the ischemic area. Figure 2 and Figure 3 show the MRI images obtained from a pig with a titanium encased ameroid and a pig with a plastic encased ameroid, respectively. After imaging, a second thoracotomy was performed for either a cell injection or sham procedure. Animals were followed out as long as 16 weeks post-ameroid placement.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56190/56190fig1.jpg" /> 
Figure 1: Images of the pig heart during the ameroid placement procedure. (A) Image of the pig heart showing the LAD, LCX, and obtuse marginal artery (OM). (B) Image of the dissected LCX artery before the ameroid placement. (C) Image of the proper placement of the ameroid around the LCX with the opening of the ameroid facing up or away from the heart. <a href="https://cloudflare.jove.com/files/ftp_upload/56190/56190fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56190/56190fig2.jpg" /> 
Figure 2: MRI image of pig heart with a titanium encased ameroid. MRI image of a pig heart taken four weeks after the placement of a titanium encased ameroid constrictor on the LCX artery. The arrow points to the artifact created by the titanium. <a href="https://cloudflare.jove.com/files/ftp_upload/56190/56190fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56190/56190fig3.jpg" /> 
Figure 3: MRI image of a pig heart with a plastic encased ameroid. MRI image of a pig heart taken four weeks after the placement of a plastic encased ameroid constrictor on the LCX artery. The arrow points to the ameroid constrictor. No artifact is observed. <a href="https://cloudflare.jove.com/files/ftp_upload/56190/56190fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Chronic Cardiac Ischemia Model in Swine Using an Ameroid Constrictor at JoVE.com

A Chronic Cardiac Ischemia Model in Swine Using an Ameroid Constrictor

The purpose of this protocol is to demonstrate the placement of a delayed constricting device (an ameroid constrictor) around a coronary artery in a swine model. This device creates an ischemic area of the heart that is useful for studying new diagnostic imaging techniques and new methods of treatment.

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but ...

a-chronic-cardiac-ischemia-model-in-swine-using-an-ameroid-constrictor

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

10.5K Views.  National Institutes of Health. The purpose of this protocol is to demonstrate the placement of a delayed constricting device (an ameroid constrictor) around a coronary artery in a swine model. This device creates an ischemic area of the heart that is useful for studying new diagnostic imaging techniques and new methods of treatment. 

Video: A Chronic Cardiac Ischemia Model in Swine Using an Ameroid Constrictor

Normothermic Cardiac Arrest and Cardiopulmonary Resuscitation: A Mouse Model of Ischemia-Reperfusion Injury

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused by whole-body hypoperfusion.5,6 Successful reproduction of whole-body hypoperfusion in rodent models has been fraught with 
difficulty.7-9,9,10 Models which employ focal ischemia have repeatedly demonstrated results which do not translate to the clinical setting, and larger 
animal models which allow for whole body hypoperfusion lack access to the full toolset of genetic manipulation possible in the mouse.11,12 However, in recent 
years a mouse model of cardiac arrest and cardiopulmonary resuscitation has emerged which can be adapted to model AKI.13 This model reliably reproduces 
physiologic, functional, anatomic, and histologic outcomes seen in clinical AKI, is rapidly repeatable, and offers all of the significant advantages of a murine surgical 
model, including access to genetic manipulative techniques, low cost relative to large animals, and ease of use. Our group has developed extensive experience 
with use of this model to assess a number of organ-specific outcomes in AKI.14,15

A powerful model for perioperative and critical care related acute kidney injury is presented. Using whole body hypoperfusion induced by cardiac arrest it is possible to nearly replicate the histologic and functional changes of clinical AKI.

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused ...

A Swine Model of Neonatal Asphyxia

Annually more than 1 million neonates die worldwide as related to asphyxia. Asphyxiated neonates commonly have multi-organ failure including hypotension, perfusion deficit, hypoxic-ischemic encephalopathy, pulmonary hypertension, vasculopathic enterocolitis, renal failure and thrombo-embolic complications. Animal models are developed to help us understand the patho-physiology and pharmacology of neonatal asphyxia. In comparison to rodents and newborn lambs, the newborn piglet has been proven to be a valuable model. The newborn piglet has several advantages including similar development as that of 36-38 weeks human fetus with comparable body systems, large body size (&tilde;1.5-2 kg at birth) that allows the instrumentation and monitoring of the animal and controls the confounding variables of hypoxia and hemodynamic derangements.
We here describe an experimental protocol to simulate neonatal asphyxia and allow us to examine the systemic and regional hemodynamic changes during the asphyxiating and reoxygenation process as well as the respective effects of interventions. Further, the model has the advantage of studying multi-organ failure or dysfunction simultaneously and the interaction with various body systems. The experimental model is a non-survival procedure that involves the surgical instrumentation of newborn piglets (1-3 day-old and 1.5-2.5 kg weight, mixed breed) to allow the establishment of mechanical ventilation, vascular (arterial and central venous) access and the placement of catheters and flow probes (Transonic Inc.) for the continuously monitoring of intra-vascular pressure and blood flow across different arteries including main pulmonary, common carotid, superior mesenteric and left renal arteries. Using these surgically instrumented piglets, after stabilization for 30-60 minutes as defined by Z&lt;10% variation in hemodynamic parameters and normal blood gases, we commence an experimental protocol of severe hypoxemia which is induced via normocapnic alveolar hypoxia. The piglet is ventilated with 10-15% oxygen by increasing the inhaled concentration of nitrogen gas for 2h, aiming for arterial oxygen saturations of 30-40%. This degree of hypoxemia will produce clinical asphyxia with severe metabolic acidosis, systemic hypotension and cardiogenic shock with hypoperfusion to vital organs. The hypoxia is followed by reoxygenation with 100% oxygen for 0.5h and then 21% oxygen for 3.5h. Pharmacologic interventions can be introduced in due course and their effects investigated in a blinded, block-randomized fashion.

Large animal models have good translational values in the examination of physiology and pharmacology of neonatal asphyxia. Using newborn piglets, we develop an experimental protocol to simulate neonatal asphyxia which has advantages of studying the systemic and regional hemodynamics, oxygen transport with biochemical and pathologic pathways and correlations.

Annually more than 1 million neonates die worldwide as related to asphyxia. Asphyxiated neonates commonly have multi-organ failure including hypotension, ...

Using Quantitative Real-time PCR to Determine Donor Cell Engraftment in a Competitive Murine Bone Marrow Transplantation Model

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules that regulate HSCs. In these transplant model systems, the function of HSCs is determined by the ability of these cells to engraft and reconstitute lethally irradiated recipient mice. Commonly, the donor cell contribution/engraftment is measured by antibodies to donor- specific cell surface proteins using flow cytometry. However, this method heavily depends on the specificity and the ability of the cell surface marker to differentiate donor-derived cells from recipient-originated cells, which may not be available for all mouse strains. Considering the various backgrounds of genetically modified mouse strains in the market, this cell surface/ flow cytometry-based method has significant limitations especially in mouse strains that lack well-defined surface markers to separate donor cells from congenic recipient cells. Here, we reported a PCR-based technique to determine donor cell engraftment/contribution in transplant recipient mice. We transplanted male donor bone marrow HSCs to lethally irradiated congenic female mice. Peripheral blood samples were collected at different time points post transplantation. Bone marrow samples were obtained at the end of the experiments. Genomic DNA was isolated and the Y chromosome specific gene, Zfy1, was amplified using quantitative Real time PCR. The engraftment of male donor-derived cells in the female recipient mice was calculated against standard curve with known percentage of male vs. female DNAs. Bcl2 was used as a reference gene to normalize the total DNA amount. Our data suggested that this approach reliably determines donor cell engraftment and provides a useful, yet simple method in measuring hematopoietic cell reconstitution in murine bone marrow transplantation models. Our method can be routinely performed in most laboratories because no costly equipment such as flow cytometry is required.

Determining donor cell engraftment presents a challenge in mouse bone marrow transplant models that lack well-defined phenotypical markers. We described a methodology to quantify male donor cell engraftment in female transplant recipient mice. This method can be used in all mouse strains for the study of HSC functions.

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules ...

Urinary Bladder Distention Evoked Visceromotor Responses as a Model for Bladder Pain in Mice

Approximately 3-8 million people in the United States suffer from interstitial cystitis/bladder pain syndrome (IC/BPS), a debilitating condition characterized by increased urgency and frequency of urination, as well as nocturia and general pelvic pain, especially upon bladder filling or voiding. Despite years of research, the cause of IC/BPS remains elusive and treatment strategies are unable to provide complete relief to patients. In order to study nervous system contributions to the condition, many animal models have been developed to mimic the pain and symptoms associated with IC/BPS. One such murine model is urinary bladder distension (UBD). In this model, compressed air of a specific pressure is delivered to the bladder of a lightly anesthetized animal over a set period of time. Throughout the procedure, wires in the superior oblique abdominal muscles record electrical activity from the muscle. This activity is known as the visceromotor response (VMR) and is a reliable and reproducible measure of nociception. Here, we describe the steps necessary to perform this technique in mice including surgical manipulations, physiological recording, and data analysis. With the use of this model, the coordination between primary sensory neurons, spinal cord secondary afferents, and higher central nervous system areas involved in bladder pain can be unraveled. This basic science knowledge can then be clinically translated to treat patients suffering from IC/BPS.

Approximately 3-8 million people in the United States suffer from interstitial cystitis/bladder pain syndrome (IC/BPS), a debilitating condition characterized in part by pelvic pain. In order to study nervous system contributions to the condition, a physiological model of urinary bladder pain is used in mice and rats.

Approximately 3-8 million people in the United States suffer from interstitial cystitis/bladder pain syndrome (IC/BPS), a debilitating condition ...

Tachycardia-Induced Cardiomyopathy As a Chronic Heart Failure Model in Swine

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment methods. Here, we present such a model by tachycardia-induced cardiomyopathy, which can be produced by rapid cardiac pacing in swine.
A single pacing lead is introduced transvenously into fully anaesthetized healthy swine, to the apex of the right ventricle, and fixated. Its other end is then tunneled dorsally to the paravertebral region. There, it is connected to an in-house modified heart pacemaker unit that is then implanted in a subcutaneous pocket. 
After 4 - 8 weeks of rapid ventricular pacing at rates of 200 - 240 beats/min, physical examination revealed signs of severe heart failure - tachypnea, spontaneous sinus tachycardia, and fatigue. Echocardiography and X-ray showed dilation of all heart chambers, effusions, and severe systolic dysfunction. These findings correspond well to decompensated dilated cardiomyopathy and are also preserved after the cessation of pacing.
This model of tachycardia-induced cardiomyopathy can be used for studying the pathophysiology of progressive chronic heart failure, especially hemodynamic changes caused by new treatment modalities like mechanical circulatory supports. This methodology is easy to perform and the results are robust and reproducible.

Here, we present a protocol to produce tachycardia-induced cardiomyopathy in swine. This model represents a potent way to study the hemodynamics of progressive chronic heart failure and the effects of applied treatment.

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment ...

Surgical Swine Model of Chronic Cardiac Ischemia Treated by Off-Pump Coronary Artery Bypass Graft Surgery

Chronic cardiac ischemia that impairs cardiac function, but does not result in infarct, is termed hibernating myocardium (HM). A large clinical subset of coronary artery disease (CAD) patients have HM, which in addition to causing impaired function, puts them at higher risk for arrhythmia and future cardiac events. The standard treatment for this condition is revascularization, but this has been shown to be an imperfect therapy. The majority of pre-clinical cardiac research focuses on infarct models of cardiac ischemia, leaving this subset of chronic ischemia patients largely underserved. To address this gap in research, we have developed a well-characterized and highly reproducible model of hibernating myocardium in swine, as swine are ideal translational models for human heart disease. In addition to creating this unique disease model, we have optimized a clinically relevant treatment model of coronary artery bypass surgery in swine. This allows us to accurately study the effects of bypass surgery on heart disease, as well as investigate additional or alternate therapies. This model surgically induces single vessel stenosis by implanting a constrictor on the left anterior descending (LAD) artery in a young pig. As the pig grows, the constrictor creates a gradual stenosis, resulting in chronic ischemia with impaired regional function, but preserving tissue viability. Following the establishment of the hibernating myocardium phenotype, we perform off-pump coronary artery bypass graft surgery to revascularize the ischemic region, mimicking the gold-standard treatment for patients in the clinic.

This protocol presents a surgical large animal model of chronic, single vessel ischemia that results in regional abnormalities but does not create infarct, known as hibernating myocardium. Following establishment of chronic ischemia, animals are treated with off-pump LIMA-LAD coronary artery bypass graft surgery to revascularize the ischemic tissue.

Chronic cardiac ischemia that impairs cardiac function, but does not result in infarct, is termed hibernating myocardium (HM). A large clinical subset of ...

Standardized Model of Ventricular Fibrillation and Advanced Cardiac Life Support in Swine

Cardiopulmonary resuscitation after cardiac arrest, independent of its origin, is a regularly encountered medical emergency in hospitals as well as preclinical settings. Prospective randomized trials in human subjects are difficult to design and ethically ambiguous, which results in a lack of evidence-based therapies. The model presented in this report represents one of the most common causes of cardiac arrests, ventricular fibrillation, in a standardized setting in a large animal model. This allows for reproducible observations and various therapeutic interventions under clinically accurate conditions, hence facilitating the generation of better evidence and eventually the potential for improved medical treatment.

Cardiopulmonary resuscitation and defibrillation are the only effective therapeutic options during cardiac arrest caused by ventricular fibrillation. This model presents a standardized regimen to induce, assess, and treat this physiological state in a porcine model, thus providing a clinical approach with various opportunities for data collection and analysis.

Cardiopulmonary resuscitation after cardiac arrest, independent of its origin, is a regularly encountered medical emergency in hospitals as well as ...

An Intact Pericardium Ischemic Rodent Model

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to induce myocardial injury). The majority of pre-clinical myocardial infarction models require the disruption of pericardial integrity with loss of the homeostatic cellular milieu. However, recently a methodology has been developed by us to induce myocardial infarction, which minimizes pericardial damage and retains the heart&#39;s resident immune cell population. An improved cardiac functional recovery in mice with an intact pericardial space following coronary ligation has been observed. This method provides an opportunity to study inflammatory responses in the pericardial space following myocardial infarction. Further development of the labeling techniques can be combined with this model to understand the fate and function of pericardial immune cells in regulating the inflammatory mechanisms that drive remodeling in the heart, including fibrosis.

This protocol outlines the steps for inducing myocardial infarction in mice while preserving the pericardium and its contents.

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to ...

Translational Rabbit Model of Chronic Cardiac Pacing

Animal models of cardiac pacing are beneficial for testing novel devices, studying the pathophysiology of artificially paced heart rhythms, and studying arrhythmia-induced cardiomyopathies and subsequent heart failure. Currently, only a few such models are available, and they mostly require extensive resources. We report a new experimental cardiac pacing model in small mammals with the potential to study arrhythmia-induced heart failure.
In six New Zealand white rabbits (mean weight: 3.5 kg) under general inhalational anesthesia the jugular region was dissected and a single pacing lead was inserted via the right external jugular vein. Using fluoroscopic guidance, the lead was further advanced to the right ventricular apex, where it was stabilized using passive fixation. A cardiac pacemaker was then connected and buried in a subcutaneous pocket.
The pacemaker implantation was successful with good healing; the rabbit anatomy is favorable for the lead placement. During 6 months of follow-up with intermittent pacing, the mean sensed myocardial potential was 6.3 mV (min: 2.8 mV, max: 12 mV), and the mean lead impedance measured was 744 &#937; (min: 370 &#937;, max: 1014 &#937;). The pacing threshold was initially 0.8 V &#177; 0.2 V and stayed stable during the follow-up.
This present study is the first to present successful transvenous cardiac pacing in a small-mammal model. Despite the size and tissue fragility, human-size instrumentation with adjustments can safely be used for chronic cardiac pacing, and thus, this innovative model is suitable for studying the development of arrhythmia-induced cardiomyopathy and consequent heart failure pathophysiology.

We present a minimally invasive leporine model of long-term cardiac pacing that can be utilized for artificial pacing and heart failure development in preclinical studies.

Animal models of cardiac pacing are beneficial for testing novel devices, studying the pathophysiology of artificially paced heart rhythms, and studying ...

Chronic Ovine Model of Right Ventricular Failure and Functional Tricuspid Regurgitation

The pathophysiology of severe functional tricuspid regurgitation (FTR) associated with right ventricular dysfunction is poorly understood, leading to suboptimal clinical results. We set out to establish a chronic ovine model of FTR and right heart failure to investigate the mechanisms of FTR. Twenty adult male sheep (6-12 months old, 62 &#177; 7 kg) underwent a left thoracotomy and baseline echocardiography. A pulmonary artery band (PAB) was placed and cinched around the main pulmonary artery (PA) to at least double the systolic pulmonary artery pressure (SPAP), inducing right ventricular (RV) pressure overload and signs of RV dilatation. PAB acutely increased the SPAP from 21 &#177; 2 mmHg to 62 &#177; 2 mmHg. The animals were followed for 8 weeks, symptoms of heart failure were treated with diuretics, and surveillance echocardiography was used to assess for pleural and abdominal fluid collection. Three animals died during the follow-up period due to stroke, hemorrhage, and acute heart failure. After 2 months, a median sternotomy and epicardial echocardiography were performed. Of the surviving 17 animals, 3 developed mild tricuspid regurgitation, 3 developed moderate tricuspid regurgitation, and 11 developed severe tricuspid regurgitation. Eight weeks of pulmonary artery banding resulted in a stable chronic ovine model of right ventricular dysfunction and significant FTR. This large animal platform can be used to further investigate the structural and molecular basis of RV failure and functional tricuspid regurgitation.

Right ventricular failure and functional tricuspid regurgitation are associated with left-sided heart disease and pulmonary hypertension, which contribute significantly to morbidity and mortality in patients. Establishing a chronic ovine model to study right ventricular failure and functional tricuspid regurgitation will help in understanding their mechanisms, progression, and possible treatments.

The pathophysiology of severe functional tricuspid regurgitation (FTR) associated with right ventricular dysfunction is poorly understood, leading to ...