The original study in which this protocol was used was supported by National Heart, Lung, and Blood Institute Grants HL-103642 and HL-088206

This investigation was conducted according to the National Institutes of Health&#39;s guidelines on animal care and use and was approved by the Boston Children&#39;s Hospital&#39;s Animal Care and Use Committee (Protocol 20-08-4247R). All the animals received humane care in compliance with the Guide for the Care and Use of Laboratory Animals.
1. Animal species, anesthetic, and analgesic agents
<ol>
	<li>Animal species: Use New Zealand White rabbits (wild type strain; female sex; sexually mature 15-20 weeks old; 3-4 kg body weight) for experimental studies.</li>
	<li>Anesthetic and analgesic agents:
	<ol>
		<li>Use atropine at a dose of 0.01 mg/kg intramuscular (IM)</li>
		<li>Use acepromazine at a dose of 0.5 mg/kg IM for initial sedation and 0.5 mg/kg intravenous (IV) for full anesthetization.</li>
		<li>Use butorphanol at a dose of 0.5 mg/kg IM.</li>
		<li>Use isoflurane via a precision vaporized system face mask at 3% for induction, followed by intubation at 1%-2%, oxygen (O2) at 100% at 2 L/min, and general anesthesia at 1% for maintenance.</li>
		<li>Use medetomidine at a dose of 0.25 mg/kg IM.</li>
		<li>Use ketamine at a dose of 10 mg/kg IV.</li>
		<li>Use a bupivacaine intercostal block at the thoracotomy site at a dose not exceeding 3 mg/kg IM.</li>
		<li>Use 1% lidocaine at a dose of 1-1.5 mL/kg IV.</li>
		<li>Use a 1-4 &#181;g/kg fentanyl transdermal patch for 72 h.</li>
	</ol>
	</li>
</ol>
2. Procedural steps (Figure 1)
<ol>
	<li>Sedate New Zealand White adult rabbits with a single combined IM injection of atropine, acepromazine, and butorphanol. Induce the animal with 3% isoflurane via a precision vaporized system face mask.</li>
	<li>Preparation prior to blinded endotracheal intubation (i.e., without visualization of the glottis)
	<ol>
		<li>Spray the larynx with 1% lidocaine to prevent laryngospasm.</li>
		<li>Pre-measure the endotracheal tube (ETT) length on the outside of the rabbit from the teeth to the predicted carina, and place the rabbit in a sternal recumbency position with the neck extended.</li>
	</ol>
	</li>
	<li>Intubate the animal with a cuffed pediatric size (3-0 or 3-5 inner diameter) ETT under continuous inhaled anesthetic at 1%-2% and O2 at 100% at 2 L/min.
	<ol>
		<li>Insert the ETT into the mouth, and direct it past the torus into the pharynx.</li>
		<li>Advance the ETT until either the tip of the tube contacts the glottis or the breath sounds are lost, indicating the tube tip has passed through the glottic opening.</li>
		<li>Slightly withdraw the tube until the breath sounds are regained, and then re-advance again, and secure the tube in place.</li>
	</ol>
	</li>
	<li>Ventilate the animal with mechanical support (tidal volume: 10 mL/kg, fraction of inspired O2: 40%, respiratory rate: 30-40 breaths/min, positive end-expiratory pressure: 5-10 cmH2O).
	<ol>
		<li>Adjust the FiO2 as tolerated to achieve an O2 saturation greater than 92% as measured by pulse oximetry to prevent hyperoxia, which can provoke a systemic inflammatory response.</li>
	</ol>
	</li>
	<li>Verify the proper placement of the ETT by a physical exam (i.e., auscultation), clinical signs (i.e., observation of condensation at the end of the endotracheal tube), and with objective measures (i.e., end-tidal carbon dioxide).</li>
	<li>After approximately 10 min, deliver an IM injection of medetomidine to the rabbit to provide simultaneous anesthetic and analgesic effects.</li>
	<li>Maintain the general anesthesia with 1% isoflurane for the duration of the surgical procedure.</li>
	<li>Insert a 22 G IV catheter into the marginal ear vein, and secure it with tape to obtain peripheral IV access. 
	NOTE: The femoral vein can be used as an alternative site of venous access.
	<ol>
		<li>Fully anesthetize the animal with acepromazine IV and ketamine IV.</li>
		<li>Prior to incision, inject 1,000 U/mL heparin at a dose of 3 mg/kg IV.
		<ol>
			<li>Administer 1,000 U/mL heparin at a dose of 3 mg/kg initially, and re-dose hourly until the end of the experiment to maintain an activated clotting time of &#62;400 s, in keeping with the current surgical protocol.</li>
		</ol>
		</li>
		<li>Administer 1% lidocaine IV and/or epicardial asynchronized defibrillation as needed if ventricular fibrillation occurs during the surgery. Ventricular fibrillation usually stops with one or two doses of lidocaine.</li>
		<li>Perfuse lactate Ringer&#39;s solution continuously at 10 mL/kg/h. 
		NOTE: Given the small volume of fluids administered and the short operative times, the animals in survival studies in this work did not require diuresis prior to extubation or during the recovery period. If the animal develops a worsening pulmonary status (i.e., increasing the ventilator settings, evidence of pulmonary edema on auscultation, etc.), diuresis is advised.</li>
	</ol>
	</li>
	<li>Perform a carotid cut down, and place a 4 or 5 French arterial line to facilitate the intra-operative monitoring of the arterial blood pressure (BP). 
	NOTE: The femoral artery can be used as an alternative site of arterial access.</li>
	<li>Monitor and record all the physiologic and mechanical variables by continuous real-time analysis.
	<ol>
		<li>Monitor the arterial BP with the carotid arterial line, and record the O2 saturation using pulse oximetry via a sensor placed on a shaved paw.</li>
		<li>Monitor with an electrocardiogram (ECG) with three limb leads: I, II, and III, and three computed augmented leads: aVL, aVR, and aVF.
		<ol>
			<li>Record the ECG tracings at the pre-ischemic baseline, during ischemia, during reperfusion, and serially during days 7-28 of recovery (if performing a survival study).</li>
		</ol>
		</li>
		<li>Monitor the level of sedation by continuous monitoring of the BP and heart rate (HR).</li>
		<li>Monitor the temperature with a rectal probe.</li>
		<li>Use 2D ECHO from the left parasternal and apical views to assess the myocardial function at desired time points in both the survival and non-survival cases.
		<ol>
			<li>Assess the myocardial function using the fractional shortening (FS) by measuring the left ventricle end-diastolic distance (LVEDD) and the left ventricle end-systolic distance (LVESD) and using the following formula: 
			FS = (LVEDD &#8722; LVESD)/LVEDD &#215; 100</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>During surgery, place the animal on a circulating hot water blanket to maintain a stable core body temperature.</li>
	<li>Prepare and drape the animal in a sterile fashion:
	<ol>
		<li>Shave the surgical site, and prep with betadine and 70% isopropyl alcohol, each applied in triplicate. Pat the area dry with sterile gauze pads, and drape the entire animal with sterile towels.</li>
	</ol>
	</li>
	<li>Left mini-thoracotomy (survival studies)
	<ol>
		<li>Perform an intercostal block at the predetermined thoracotomy site with bupivacaine IM.</li>
		<li>Administer 1% lidocaine IV via the auricular vein prior to incision.</li>
		<li>Perform a left mini-thoracotomy through the fourth intercostal space along the upper portion of the fifth rib to avoid the neuromuscular bundle, which is located parallel to the undersurface of each rib.
		<ol>
			<li>Perform an anterolateral thoracotomy for the best visualization of the anterolateral surface of the heart (i.e., the anatomic location of the LAD diagonal branches).</li>
			<li>Position the rabbit with the left side elevated approximately 30&#176; using a pillow or bean bag.</li>
			<li>Secure the rabbit&#39;s ipsilateral leg above its head to create room for both the operative field and between the rib spaces.</li>
			<li>Palpate and outline the bony landmarks, including the ribs, sternum, and scapula, with a felt-tipped marking pen. Incise the skin overlying the fifth rib using a #10 blade. Ensure that the incision remains parallel to the rib.</li>
			<li>Use electrocautery to divide the pectoralis major muscle and serratus anterior muscle. Divide the intercoastal muscles right above the fifth rib with electrocautery to preserve the neurovascular bundle.</li>
			<li>Cautiously enter the pleural space through the fourth intercostal space with sharp or blunt dissection. Extend the initial pleural incision parallel to the rib in both directions with sharp or blunt dissection until a rib spreader or sternal retractor can be inserted.</li>
		</ol>
		</li>
		<li>Place a rib spreader or sternal retractor within the rib space, and widen to provide adequate visualization of the heart and pericardial sac. Lift the pericardium with DeBakey forceps, and open the pericardium with Metzenbaum scissors.</li>
		<li>LAD artery isolation
		<ol>
			<li>Encircle the second or third diagonal branch of the LAD artery with a polypropylene suture (3-0) on a taper needle. Remove the needle, and thread both ends of the polypropylene filament through a small vinyl tube to form a snare.</li>
			<li>Place a pledget between the snare and the coronary artery to avoid damaging the coronary and/or causing vasospasm with ligation.
			<ol>
				<li>Using DeBakey forceps, pick up a rectangular PTFE felt pledget (approximately 7 mm x 3 mm). Place the pledget between the two polypropylene filaments so that it is sandwiched between the isolated LAD artery and the vinyl tube when the snare is tightened.</li>
			</ol>
			</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Midline sternotomy (non-survival studies) 
	NOTE: The midline sternotomy approach is ideal for non-survival cases, for which more invasive monitoring with LVDP and sonomicrometry can be utilized.
	<ol>
		<li>Perform a midline sternotomy using curved Mayo scissors. Place a sternal retractor, and widen it to provide adequate visualization of the heart and pericardial sac.</li>
		<li>Lift the pericardium with DeBakey forceps, and open the pericardium with Metzenbaum scissors.</li>
		<li>Placing the three piezoelectric sonomicrometry crystals:
		<ol>
			<li>Make three small 1 mm cuts on the epicardium of the LV, forming the corners of a triangle. Place the piezoelectric sonomicrometry crystals inside the epicardium cuts.</li>
			<li>Secure the wires to the cardiac surface with a 5-0 polypropylene U-stitch. When recording using sonomicrometry, pause the mechanical ventilation to allow for an accurate recording over two to three heartbeats. 
			NOTE: If the heart fibrillates, 1% lidocaine is not effective, and epicardial defibrillation is needed, turn off the sonomicrometer, and disconnect it from the data acquisition system to protect both from electrical input.</li>
		</ol>
		</li>
		<li>LAD artery isolation:
		<ol>
			<li>Encircle the second or third diagonal branch of the LAD artery with a polypropylene suture (3-0) on a taper needle.</li>
			<li>Remove the needle, and thread both ends of the polypropylene filament through a small vinyl tube to form a snare.</li>
			<li>Place a pledget between the snare and the coronary artery to avoid damaging the coronary artery and/or causing vasospasm with ligation.</li>
			<li>Using DeBakey forceps, pick up a rectangular PTFE felt pledget (approximately 7 mm x 3 mm). Place the pledget between the two polypropylene filaments so that it is sandwiched between the isolated LAD artery and the vinyl tube when the snare is tightened.</li>
		</ol>
		</li>
		<li>Measurement of the LVDP:
		<ol>
			<li>Place a 5-0 polypropylene U-stitch in the apex of the LV. Make a small 1 mm incision with an 11 blade into the LV apex.</li>
			<li>Insert a 3 French balloon catheter into the LV lumen. Secure the catheter to the LV by tying it to the 5-0 polypropylene U-stitch suture.</li>
			<li>Connect the catheter to the transducer connected to the monitor to record the LVDP. Record the LVDP using the data acquisition system (described below). Zero the catheter to record the hemodynamic variables by opening the three-way stopcock to the air and zeroing on the monitor.</li>
		</ol>
		</li>
		<li>Data acquisition system
		<ol>
			<li>Initiate the data acquisition system (see the Table of Materials) on the computer/laptop being used. Connect the wire from the monitor to the computer/laptop.</li>
			<li>Select Channel 1 on the data acquisition system, and name it LVDP. Zero the transducer using the monitor. 
			NOTE: If connecting the BP and HR to the data acquisition system, follow the same process: connect the wire to the laptop, select Channel, and zero if measuring the BP.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Occlude the coronary artery by tightening the snare by pressing down on the vinyl tube while pulling up on the polypropylene suture filaments. Maintain the desired tightness with a mosquito clamp by directly clamping the tube and fixing it in place.</li>
	<li>Confirm myocardial ischemia visually by the regional cyanosis of the epicardium. Regional ischemia can also be confirmed on ECG with the presence of an ST segment and T wave changes.</li>
	<li>After visual confirmation, induce regional ischemia for 30 min under anesthesia.
	<ol>
		<li>At 0 min, 10 min, 20 min, and 30 min during the regional ischemia, assess the FS by 2D ECHO for both the survival and non-survival cases.</li>
		<li>Assess the LVDP and sonomicrometry continuously during the pre-ischemia time, myocardial ischemia time, and post-ischemic time for the non-survival cases.</li>
		<li>If needed, delineate the area at risk by ligating the artery again with the polypropylene suture stitch left in place. Cross-clamp the aorta, and inject Monastral Blue pigment 98% (diluted 1:5 in PBS) through the aorta using a cardioplegia needle. The perfused areas of the myocardium will stain blue, and the area at risk will remain unstained.</li>
		<li>Continuously monitor and record the HR, BP, and O2 saturation.</li>
		<li>Allow the animal to recover for 2 h (non-survival) or 28 days (survival). 
		NOTE: ECG can be used to confirm reperfusion. Although not seen in the experiment conducted in this study, hypokalemia can often occur during reperfusion and can be corrected with potassium control or an appropriate infusion.</li>
	</ol>
	</li>
	<li>Conclusion of the procedure
	<ol>
		<li>Survival cases
		<ol>
			<li>In survival cases, trim the 3-0 polypropylene thread used for the snare, loosely tie the ends together, and leave it in place. Identify the area at risk and the infarct zone by the 3-0 polypropylene thread.</li>
			<li>After the procedure is completed, close the incision in three layers.
			<ol>
				<li>Close the first layer by tying two 2-0 polyglactin 910 figure-of-eight stitches around the ribs.</li>
				<li>Close the muscular and subcutaneous layers with a 3-0 polydioxanone suture in a running fashion.</li>
				<li>Close the skin in a subcuticular fashion using a 5-0 monofilament suture. Use a buried running suture to minimize the irritation felt by the animal.</li>
			</ol>
			</li>
			<li>Evacuate the pleural air by performing a needle thoracentesis.</li>
			<li>Apply a fentanyl transdermal patch for 72 h to facilitate the postoperative pain management.</li>
			<li>Perform transthoracic echocardiography at the 1 week and 2 week time points postoperatively to assess the trends in the FS.</li>
			<li>Following the predetermined recovery period, sedate, intubate, and anesthetize the animal as above. Perform a median sternotomy. Expose and open the pericardial sac. Euthanize the rabbit under deep anesthesia by removing the heart en bloc,&#160;allowing the animal to expire by exsanguination.</li>
		</ol>
		</li>
		<li>Non-survival cases
		<ol>
			<li>After the experiment and having assured deep anesthesia, completely expose the heart, and remove it en bloc for biochemical and tissue analysis. The animal expires by exsanguination.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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Heart and Circulatory Physiology. 304 (7), H966-H982 (2013).</li><li>Pombo, J. F., Troy, B. L., Russell, R. O. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Left+ventricular+volumes+and+ejection+fraction+by+echocardiography.">Left ventricular volumes and ejection fraction by echocardiography.</a> Circulation. 43 (4), 480-490 (1971).</li><li>McCully, J. D., Wakiyama, H., Hsieh, Y. J., Jones, M., Levitsky, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+contribution+of+necrosis+and+apoptosis+in+myocardial+ischemia-reperfusion+injury.">Differential contribution of necrosis and apoptosis in myocardial ischemia-reperfusion injury.</a> American Journal of Physiology. Heart and Circulatory Physiology. 286 (5), H1923-H1935 (2004).</li><li>Masuzawa, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transplantation+of+autologously+derived+mitochondria+protects+the+heart+from+ischemia-reperfusion+injury.">Transplantation of autologously derived mitochondria protects the heart from ischemia-reperfusion injury.</a> American Journal of Physiology. Heart and Circulatory Physiology. 304 (7), 966-982 (2013).</li><li>Abarbanell, A. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+myocardial+and+vascular+injury.">Animal models of myocardial and vascular injury.</a> Journal of Surgical Research. 162 (2), 239-249 (2010).</li><li>Pirat, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+feature-tracking+echocardiographic+method+for+the+quantitation+of+regional+myocardial+function:+Validation+in+an+animal+model+of+ischemia-reperfusion.">A novel feature-tracking echocardiographic method for the quantitation of regional myocardial function: Validation in an animal model of ischemia-reperfusion.</a> Journal of the American College of Cardiology. 51 (6), 651-659 (2008).</li><li>Verdouw, P. D., vanden Doel, M. A., de Zeeuw, S., Duncker, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+in+the+study+of+myocardial+ischaemia+and+ischaemic+syndromes.">Animal models in the study of myocardial ischaemia and ischaemic syndromes.</a> Cardiovascular Research. 39 (1), 121-135 (1998).</li><li>Bolukoglu, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+animal+model+of+chronic+coronary+stenosis+resulting+in+hibernating+myocardium.">An animal model of chronic coronary stenosis resulting in hibernating myocardium.</a> The American Journal of Physiology. 263, H20-H29 (1992).</li><li>Heyndrickx, G. R., Millard, R. W., McRitchie, R. J., Maroko, P. R., Vatner, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regional+myocardial+functional+and+electrophysiological+alterations+after+brief+coronary+artery+occlusion+in+conscious+dogs.">Regional myocardial functional and electrophysiological alterations after brief coronary artery occlusion in conscious dogs.</a> Journal of Clinical Investigation. 56 (4), 978-985 (1975).</li></ol>

No conflicts of interest, financial or otherwise, are declared by the authors.

Our protocol demonstrates a reliable methodology for performing acute regional myocardial ischemia in the rabbit. The left mini-thoracotomy approach is ideal for survival cases, for which the incision and associated pain must be minimized. Importantly, diuretic therapy was not necessary prior to extubation, and there was no mortality intraoperatively in the non-survival group or at 4 weeks postoperatively in the survival group. When the design of the protocol requires a non-survival case, or when more detailed monitoring of the global and regional myocardial function is needed, a midline sternotomy can be used (Figure 1).
The most critical steps of the protocol are to carefully encircle the LAD with a taper needle without damaging the artery or creating venous bleeding and to occlude the LAD in order to create a consistent area at risk.
Some complications that can be experienced when performing the described surgery are lung overdistension during mechanical ventilation due to a high tidal volume, bleeding from damage to the LAD, bleeding secondary to intercostal vessel injury, which usually occurs upon entry or from retractor manipulation, and/or cardiac arrhythmia (intraoperative ventricular fibrillation) with LAD ligation. Other postoperative complications may also occur, such as surgical site infection, poor animal mobilization due to pain, and/or residual myocardial regional hypokinesis. Despite the incidence of these complications being very low, the investigator should be able to readily and effectively address them.
Rabbits present as an excellent animal model for myocardial studies. Their heart rate is similar to the human heart rate, and their size is sufficiently small but allows for histological analysis under an optical microscope.
A limitation in this study should be acknowledged; specifically, the rabbit heart is smaller and clinically less relevant for comparisons to the human heart than the hearts of other large animal models such as the pig.
Due to the incidence and prevalence of cardiovascular disease, having an animal model that simulates regional myocardial ischemia is of paramount importance. This methodology could have multiple applications and has been proven to be useful in models of vascular injury, chronic myocardial ischemia, and short periods of myocardial stunning27,28,29,30,31.

Cardiovascular diseases are the leading cause of death in the world and contribute to over 18 million deaths each year1,2,3. Acute myocardial infarction (MI) is a common medical emergency that develops when a blood clot or a piece of atheromatous plaque blocks the blood flow of a coronary artery. This causes regional myocardial ischemia in the territory that the artery perfuses.
The present study describes a protocol that utilizes a simple and reliable methodology to create in situ acute regional myocardial ischemia in a rabbit model for non-survival and survival experiments. The initial goal of this method was to evaluate the effects of mitochondrial transplantation on modulating myocardial necrosis and increasing the post-ischemic heart function following an ischemic event. Previous research has demonstrated the occurrence of mitochondrial alterations and a rapid decline in high-energy phosphate levels following the onset of ischemia and a reduction in the oxygen supply, resulting in a drastic decrease in the cardiac energy stores4. Investigators have attempted to improve post-ischemic function and lessen myocardial tissue necrosis using pharmacological interventions and/or procedural techniques, but these techniques provide limited cardioprotection and have minimal impact on mitochondrial damage and dysfunction5,6,7. Our team and others have previously shown that mitochondrial damage primarily occurs during ischemia and that contractile recovery can be enhanced and the myocardial infarct size decreased with the preservation of mitochondrial respiratory function during reperfusion8,9,10. Thus, we hypothesized that mitochondrial transplantation from tissues unaffected by ischemia to the area of ischemia prior to reperfusion would provide an alternative approach to reduce myocardial necrosis and enhance myocardial function. Herein, we detail the protocol used to test this theory and the representative results obtained from our initial study analysis.
Furthermore, several investigators have focused on other topics integral to defining the impact of myocardial ischemia-reperfusion injury and establishing appropriate therapeutic interventions. One such area of research is that of preconditioning. Myocardial ischemic preconditioning is a cardioprotective mechanism activated by brief ischemic stress that results in a reduction in the rate of cardiac cell necrosis during subsequent episodes of prolonged ischemia. These mechanisms can be activated by either hypoxia or coronary occlusion. Mandel et al. demonstrated that hypoxic-hyperoxic preconditioning helped to maintain the balance of nitric oxide metabolites, reduced endothelin-1 hyperproduction, and supported organ protection11. Moreover, the concept of remote ischemic preconditioning, a phenomenon whereby single-organ preconditioning provides systemic protection, has been explored. Ali et al. found that, in patients undergoing elective open abdominal aortic aneurysm repair, remote preconditioning, performed by intermittently cross-clamping the common iliac artery to serve as a stimulus, reduced the incidence of postoperative myocardial injury, myocardial infarction, and renal impairment12.
Rabbit models offer potential advantages over models with other species and have been used in multiple different scenarios for decades, including the induction of arrhythmias, global and regional ischemic models, and cardiac contraction research, amongst others13,14,15. Although the rabbit heart is smaller than that of a dog or pig, it is large enough to easily perform surgical procedures at a much lower cost13. The rabbit heart is often used as it closely parallels the human heart; indeed, it has a similar metabolic rate, expresses &#946;-myosin heavy chain, andlacks significant myocardial xanthine oxidase16. The technique herein described to induce regional myocardial ischemia is simple, repeatable, and cost-effective. This method allows for both non-survival and survival cases, as only regional ischemia is induced rather than global ischemia, and the materials needed are non-specialized. Two different surgical approaches (i.e., sternotomy and mini-thoracotomy) can be utilized, thus providing the operator and experimental protocols more freedom in terms of the study design. Additionally, the procedure does not require the use of a cardiopulmonary bypass. In this context, minimally invasive approaches to coronary artery bypass grafting have become valuable alternatives for patients in need of multi-vessel revascularizaiton17,18. This model could be used to study the differences between these approaches and provide an animal-based learning tool for surgical trainees. Additionally, performing cardiac catheterization utilizing this model may be useful for physiological research and/or surgical training.
Our model provides a methodology for applications in which inducing regional myocardial ischemia and subsequently measuring the infarct size, myocardial function, and cellular changes are of importance. With this protocol, we have been able to evaluate several markers of cellular function and adaptation to ischemia and the proposed therapeutic intervention (i.e., mitochondrial transplantation) by examining the internalization of organelles, oxygen consumption, high-energy phosphate synthesis, and the induction of cytokine mediators and proteomic pathways. These outcomes are important in preserving the myocardial energetics, cell viability, and cardiac function and allow for the objective evaluation of cardioprotective techniques following ischemia-reperfusion injury. This model could be used to study similar biologic pathways and alternatives in the field of post-ischemic myocardial pathology and recovery.
The goal of this protocol is to provide a highly reproducible methodology to induce in situ acute regional myocardial ischemia in the rabbit for non-survival and survival experiments. This model provides a methodology with high survival, low intraoperative mortality, and minimal morbidity19. Other models for acute regional myocardial ischemia have been described using radiolabeled materials, contrast agents, magnetic resonance imaging, or computer simulations20,21,22. Our protocol provides a reliable and simple methodology that is cost-effective, consistently reproducible, and has a low technical demand and, thus, can be performed by investigators without surgical expertise. This protocol accommodates either a survival project using a left mini-thoracotomy or a non-survival model using a midline sternotomy.

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			<td>#10 blade</td>
			<td>Bard Parker</td>
			<td>371210</td>
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			<td>#11 blade</td>
			<td>Fisher Scientific</td>
			<td>B3L</td>
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			<td>22 G PIV needle</td>
			<td>BD Insyte</td>
			<td>381423</td>
			<td></td>
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			<td>Acepromazine</td>
			<td>VETONE</td>
			<td>NDC 13985-587-50</td>
			<td>0.5 mg/kg IM and IV</td>
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			<td>Aline pressure bag</td>
			<td>Infu-Stat</td>
			<td>2139</td>
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			<td>Angiocath</td>
			<td>Becton Dickinson</td>
			<td>382512</td>
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			<td>Arterial Catheter</td>
			<td>Teleflex</td>
			<td>MC-004912</td>
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			<td>Atropine</td>
			<td>Hikma Pharmaceuticals</td>
			<td>NDC 0641-6006-01</td>
			<td>&#160;0.01 mg/kg IM</td>
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			<td>Betadine and 70% isopropyl alcohol</td>
			<td>McKesson</td>
			<td>NDC 68599-2302-6</td>
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			<td>Blood gas machine</td>
			<td>Siemens</td>
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			<td>Valleylab</td>
			<td>E6008</td>
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			<td>Bulldog clamps</td>
			<td>World Precision Instruments</td>
			<td>14119</td>
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			<td>Auromedics</td>
			<td>NDC 55150-249-50</td>
			<td>&#160;3 mg/kg IM</td>
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			<td>Butorphanol</td>
			<td>Roxane</td>
			<td>NDC 2054-3090-36</td>
			<td>0.5 mg/kg IM</td>
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			<td>Clear acetate sheet</td>
			<td>Oxford Instruments</td>
			<td>ID 51-1625-0213</td>
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			<td>Clipers</td>
			<td>Andis</td>
			<td>AGC2</td>
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			<td>DeBakey forceps</td>
			<td>Integra</td>
			<td>P6280</td>
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			<td>Echocardiography machine</td>
			<td>Philips</td>
			<td>IE33 F1</td>
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			<td>Electrocardiography machine</td>
			<td>Meditech</td>
			<td>MD908B</td>
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			<td>Endotracheal tube</td>
			<td>Medline</td>
			<td>#922774</td>
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			<td>Fentanyl</td>
			<td>West-Ward</td>
			<td>NDC 0641-6030-01</td>
			<td>1&#8211;4 &#181;g/kg transdermal patch</td>
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			<td>Formaldehyde solution 10%</td>
			<td>Epredia</td>
			<td>94001</td>
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			<td>Glass plates</td>
			<td>&#160;United Scientific</td>
			<td>B01MUHX6MR</td>
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			<td>Heparin Sodium</td>
			<td>Sagent</td>
			<td>NDC 69-0058-02</td>
			<td>1000U in 1 mL 3 mg/kg</td>
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			<td>Hot water blanket</td>
			<td>3M</td>
			<td>55577</td>
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			<td>Isoflurane</td>
			<td>Penn Veterinary Supply, INC</td>
			<td>NDC 50989-606-15</td>
			<td>1%&#8211;3%</td>
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		<tr>
			<td>Ketamine</td>
			<td>Dechra</td>
			<td>NDC 42023-138-10</td>
			<td>10 mg/kg IV</td>
		</tr>
		<tr>
			<td>Lab Chart 7 Acquisition Software</td>
			<td>Adinstruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Lactated Ringer&#39;s solution</td>
			<td>ICUmedical</td>
			<td>NDC 0990-7953-09</td>
			<td>10 mL/kg/h</td>
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		<tr>
			<td>Laryngoscope</td>
			<td>Welch Allyn</td>
			<td>68044</td>
			<td></td>
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		<tr>
			<td>Left ventricule lumen catheter&#160;3Fr</td>
			<td>McKesson</td>
			<td>385764-EA</td>
			<td></td>
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			<td>Lidocaine (1%)</td>
			<td>Pfizer</td>
			<td>4276-01</td>
			<td>1&#8211;1.5 mL/kg IV</td>
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		<tr>
			<td>LVDP transducer</td>
			<td>Edward</td>
			<td>PDP-ED</td>
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			<td>Marking pen</td>
			<td>Viscot</td>
			<td>1451SR-100 Unsterile</td>
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			<td>Mayo scissors</td>
			<td>Mayo</td>
			<td>S7-1098</td>
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		<tr>
			<td>Medetomidine</td>
			<td>Entireoly Pets Pharmacy</td>
			<td>NDC 015914-005-01</td>
			<td>0.25 mg/kg IM</td>
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			<td>Metzenbaum scissors</td>
			<td>Cole-Parmer</td>
			<td>UX-10821-05</td>
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		<tr>
			<td>Monastra. Blue pigment 98%</td>
			<td>Chemsavers</td>
			<td>MBTR1100G</td>
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		<tr>
			<td>Monocryl 5-0</td>
			<td>Ethicon</td>
			<td>Y463G</td>
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		<tr>
			<td>Mosquito clamp</td>
			<td>Shioda</td>
			<td>802N</td>
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		<tr>
			<td>PDS 3-0</td>
			<td>Ethicon</td>
			<td>42312201</td>
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			<td>Piezoelectric sonomicrometry crystals</td>
			<td>Sonometrics</td>
			<td>Small 2mm round</td>
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			<td>Plegets</td>
			<td>DeRoyal</td>
			<td>32-363</td>
			<td></td>
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		<tr>
			<td>Povuine Iodine Prep Solutions</td>
			<td>Medline</td>
			<td>MDS093940</td>
			<td></td>
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		<tr>
			<td>Precision vaporized system face mask</td>
			<td>Yuwell</td>
			<td>B07PNH69BF</td>
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			<td>Prolene 3-0</td>
			<td>Ethicon</td>
			<td>8665G</td>
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		<tr>
			<td>Proline 5-0</td>
			<td>Ethicon</td>
			<td>8661G</td>
			<td></td>
		</tr>
		<tr>
			<td>Pulse oximetry probe</td>
			<td>Masimo</td>
			<td>9216-U</td>
			<td></td>
		</tr>
		<tr>
			<td>Rib spreader</td>
			<td>Medline</td>
			<td>MDS5621025</td>
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		<tr>
			<td>S12 Pediatric Sector Probe</td>
			<td>Phillips</td>
			<td>21380A</td>
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			<td>Sonomicrometer</td>
			<td>Sonometrics</td>
			<td>BZ10123724</td>
			<td></td>
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		<tr>
			<td>Sterile gauze</td>
			<td>Medline</td>
			<td>3.00802E+13</td>
			<td></td>
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			<td>Sterile towels</td>
			<td>McKesson</td>
			<td>MON 277860EA</td>
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			<td>Sternal retractor</td>
			<td>Medline</td>
			<td>MDS5610321</td>
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		<tr>
			<td>Sutures for closure</td>
			<td>J&#38;J Dental</td>
			<td>8698G</td>
			<td></td>
		</tr>
		<tr>
			<td>Telemetriy monitor</td>
			<td>Meditech</td>
			<td>MD908B</td>
			<td></td>
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		<tr>
			<td>Temperature probe</td>
			<td>Omega</td>
			<td>KHSS-116G-RSC-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Triphenyl tetrazolium chloride (1%)</td>
			<td>Millipore</td>
			<td>MFCD00011963</td>
			<td></td>
		</tr>
		<tr>
			<td>Ventilator</td>
			<td>MedGroup</td>
			<td>MSLGA 11</td>
			<td></td>
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		<tr>
			<td>Vicryl 2-0</td>
			<td>Ethicon</td>
			<td>V635H</td>
			<td></td>
		</tr>
		<tr>
			<td>Vinyl tubing</td>
			<td>ABE</td>
			<td>DISW 3001</td>
			<td></td>
		</tr>
	</tbody>
</table>

model of ischemia and reperfusion injury in rabbits

The protocol here provides a simple, highly replicable methodology to induce in situ acute regional myocardial ischemia in the rabbit for non-survival and survival experiments. New Zealand White adult rabbit is sedated with atropine, acepromazine, butorphanol, and isoflurane. The animal is intubated and placed on mechanical ventilation. An intravenous catheter is inserted into the marginal ear vein for the infusion of medications. The animal is pre-medicated with heparin, lidocaine, and lactated Ringer&#39;s solution. A carotid cut-down is performed to obtain arterial line access for blood pressure monitoring. Select physiologic and mechanical parameters are monitored and recorded by continuous real-time analysis.
With the animal sedated and fully anesthetized, either a fourth intercostal space small left thoracotomy (survival) or midline sternotomy (non-survival) is performed. The pericardium is opened, and the left anterior descending (LAD) artery is located.
A polypropylene suture is passed around the second or third diagonal branch of the LAD artery, and the polypropylene filament is threaded through a small vinyl tube, forming a snare. The animal is subjected to 30 min of regional ischemia, achieved by occluding the LAD by tightening the snare. Myocardial ischemia is confirmed visually by regional cyanosis of the epicardium. Following regional ischemia, the ligature is loosened, and the heart is allowed to re-perfuse.
For both survival and non-survival experiments, the myocardial function can be assessed via an echocardiography (ECHO) measurement of the fractional shortening. For non-survival studies, data from sonomicrometry collected using three digital piezoelectric ultrasonic probes implanted within the ischemic area and the left ventricle developed pressure (LVDP) using an apically inserted left ventricle (LV) catheter can be continuously acquired for evaluating the regional and global myocardial function, respectively.
For survival studies, the incision is closed, a left needle thoracentesis is performed for pleural air evacuation, and postoperative pain control is achieved.

Following the protocol (Figure 1), myocardial ischemia was confirmed immediately by the direct visualization of cyanosis of the epicardium.
Standard ECGs (three limb leads: I, II, and III, and three computed augmented leads: aVL, aVR, and aVF) were recorded continuously pre-ischemia, during ischemia, and at reperfusion (Figure 2). The ECGs demonstrate tachycardia, arrhythmias (i.e., ventricular fibrillation), conduction system defects (i.e., bundle branch block), the development of infarct-related Q waves, and ST segment deviation23.
During regional ischemia, regional hypokinesia was directly observed with the naked eye in the mid cavity of the anterior wall in all the hearts, consistent with the LAD artery&#39;s perfusion area that was made ischemic by the restriction of flow with the temporary snaring of the LAD. In both the survival and non-survival cases, 2D ECHO readings were obtained during pre-ischemia, just before inducing regional ischemia, and at different time points during the experiment: 5 min, 10 min, 15 min, 30 min, 60 min, and 120 min. The left ventricular end-diastolic (LVEDD) and left ventricular end-systolic dimensions (LVESD) were measured with a 2D-guided M-mode ECHO at the maximal and minimal LV circumferences, respectively. The regional LV wall contractility in the myocardial ischemic zone was assessed from short-axis views of the LV utilizing M-mode, with the curser line overlying the area at risk. The fractional shortening (FS) was calculated with the following formula: FS = (LVEDD &#8722; LVESD)/LVEDD &#215; 10024. The results showed that the fractional shortening decreased during the ischemic time and post-ischemic time compared to the pre-ischemic time (Figure 3)
To quantify the extent of myocardial injury, the infarct size can be measured biochemically with triphenyl tetrazolium chloride (TTC) (Sigma Chemical Co., St. Louis, MO) staining. In this work, the area at risk was delineated by the re-ligation of the involved artery by tying the Prolene stitch left in place. The aorta was cross-clamped, and Monastral Blue pigment (diluted 1:5 in PBS) was delivered through the aorta using a cardioplegia needle. The areas of perfused myocardium were stained blue, and the area at risk remained unstained owing to the ligation of the artery.
The heart was sliced across the long axis of the left ventricle, from apex to base, into 1 cm thick transverse sections, placed between glass plates, and compressed with bulldog clamps. The area at risk for each side of each section was traced on a clear acetate sheet. The heart sections were incubated in a dark container with 1% TTC in phosphate buffer (pH 7.4) at 38&#160;&#176;C for 20 min. The heart sections were then stored in a 10% formaldehyde solution for 24 h before the final measurements to enhance the visualization of the infarct zone. The sections were placed between glass plates and compressed with bulldog clamps. Myocardial necrosis was evidenced by a white area on the myocardial tissue, and the brick-red areas showed the viable tissue. (Figure 4) The infarcted regions (white) within the areas at risk for each side of each section were traced on the clear acetate sheet. Planimetry was used to measure the area at risk and the infarct zone. The volumes of the area at risk and infarcted zone were calculated by multiplying the planimetered areas by the slice thickness. The infarct volume was expressed as a percentage of the total LV volume for each heart25. The ratio of the area at risk to the LV weight was calculated, and the infarct size was expressed as a percentage of the area at risk. Our previous work demonstrated that, after 2 h and 28 days of recovery, the areas at risk (i.e., as a percentage of the LV mass) were approximately 29% and 27%, respectively, for both the mitochondrial and control groups However, after 2 h and 28 days of recovery, the infarct size (i.e., infarct size/area at risk) in the mitochondrial hearts was 9.8% and 7.9%, respectively, compared to 37% and 34% in the control hearts26. Additionally, in our previous experiments, the fractional shortening and LVDP were decreased in the control group to 50%-60% and 70%-80%, respectively, compared to baseline.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64752/64752fig01.jpg" /> 
Figure 1: Protocol diagram. The protocol can be adjusted based on the needs of the experiment either for survival or non-survival cases. Non-survival cases can be performed with a more invasive surgical approach using a midline sternotomy, thus allowing for the use of sonomicrometry crystals, epicardial echocardiography (ECHO), and an LV catheter for the measurement of the fractional shortening and LVDP. For survival cases, for which incision healing and pain management must be considered, a left mini-thoracotomy can be performed, and the myocardial function can be assessed at different time points during a longer study period using 2D ECHO. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64752/64752fig01large.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/64752/64752fig02.jpg" /> 
Figure 2: Representative electrocardiogram (limb II and computed augmented lead aVL) before the regional ischemia induction, during the ischemic time, and during the reperfusion. Millivolt and millisecond scales are shown on the left. The time points and the moment of the left anterior descending artery snaring are shown at the bottom. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64752/64752fig02large.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/64752/64752fig03.jpg" /> 
Figure 3: Echocardiographic assessment of the heart by measuring the fractional shortening (FS). The fractional shortening was measured by obtaining the left ventricular end-diastolic distance and the left ventricular end-systolic distance with 2D-guided M-mode at the maximal and minimal LV circumferences, respectively. The fractional shortening was assessed at (A) baseline/pre-ischemia, (B) during the temporary snaring of the left anterior descending (LAD) artery with the cursor line overlying the area at risk, and (C) during reperfusion after releasing the snare on the LAD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64752/64752fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64752/64752fig04.jpg" /> 
Figure 4: Representative images of the infarct size for a heart stained with 1% triphenyl tetrazolium chloride after 30 min of induced myocardial regional ischemia. The viable tissue is seen as red, while the infarct is seen as white areas. Scale bar = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64752/64752fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Model of Ischemia and Reperfusion Injury in Rabbits at JoVE.com

Model of Ischemia and Reperfusion Injury in Rabbits

The present study demonstrates a highly reproducible animal model of acute regional myocardial ischemia and reperfusion injury in rabbits using a left mini-thoracotomy for survival cases or a midline sternotomy for non-survival cases.

The protocol here provides a simple, highly replicable methodology to induce in situ acute regional myocardial ischemia in the rabbit for non-survival and ...

model-of-ischemia-and-reperfusion-injury-in-rabbits

Nanyang Technological University

Research

JoVE Journal

Medicine

1.0K Views.  Harvard Medical School. The present study demonstrates a highly reproducible animal model of acute regional myocardial ischemia and reperfusion injury in rabbits using a left mini-thoracotomy for survival cases or a midline sternotomy for non-survival cases. 

Video: Model of Ischemia and Reperfusion Injury in Rabbits

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 ...

Ischemia-reperfusion Model of Acute Kidney Injury and Post Injury Fibrosis in Mice

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often unreported mortality rates that may confound analyses. Bilateral renal pedicle clamping is commonly used to induce IR-AKI, but differences between effective clamp pressures and/or renal responses to ischemia between kidneys often lead to more variable results. In addition, shorter clamp times are known to induce more variable tubular injury, and while mice undergoing bilateral injury with longer clamp times develop more consistent tubular injury, they often die within the first 3 days after injury due to severe renal insufficiency. To improve post-injury survival and obtain more consistent and predictable results, we have developed two models of unilateral ischemia-reperfusion injury followed by contralateral nephrectomy. Both surgeries are performed using a dorsal approach, reducing surgical stress resulting from ventral laparotomy, commonly used for mouse IR-AKI surgeries. For induction of moderate injury BALB/c mice undergo unilateral clamping of the renal pedicle for 26 min and also undergo simultaneous contralateral nephrectomy. Using this approach, 50-60% of mice develop moderate AKI 24 hr after injury but 90-100% of mice survive. To induce more severe AKI, BALB/c mice undergo renal pedicle clamping for 30 min followed by contralateral nephrectomy 8 days after injury. This allows functional assessment of renal recovery after injury with 90-100% survival. Early post-injury tubular damage as well as post injury fibrosis are highly consistent using this model.

We describe models of moderate and severe ischemia-reperfusion-induced kidney injury in which the mice undergo unilateral renal pedicle clamping followed by simultaneous or delayed contralateral nephrectomy, respectively. These models consistently give rise to renal dysfunction and post-injury fibrosis, but injury severity and survival are dependent on mouse background, age and surgical equipment.

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often ...

Murine Model of Intestinal Ischemia-reperfusion Injury

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, hypotension, necrotizing enterocolitis, bowel transplantation, trauma and chronic inflammation. Intestinal ischemia-reperfusion (IR) injury is a consequence of acute mesenteric ischemia, caused by inadequate blood flow through the mesenteric vessels, resulting in intestinal damage. Reperfusion following ischemia can further exacerbate damage of the intestine. The mechanisms of IR injury are complex and poorly understood. Therefore, experimental small animal models are critical for understanding the pathophysiology of IR injury and the development of novel therapies.
Here we describe a mouse model of acute intestinal IR injury that provides reproducible injury of the small intestine without mortality. This is achieved by inducing ischemia in the region of the distal ileum by temporally occluding the peripheral and terminal collateral branches of the superior mesenteric artery for 60 min using microvascular clips. Reperfusion for 1 hr, or 2 hr after injury results in reproducible injury of the intestine examined by histological analysis. Proper position of the microvascular clips is critical for the procedure. Therefore the video clip provides a detailed visual step-by-step description of this technique. This model of intestinal IR injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Here we describe the detailed procedure of intestinal ischemia-reperfusion in mice which results in reproducible injury without mortality to encourage the standardization of this technique across the field. This model of intestinal ischemia-reperfusion injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, ...

Confirmation of Myocardial Ischemia and Reperfusion Injury in Mice Using Surface Pad Electrocardiography

Many animal models have been established for the study of myocardial remodeling and heart failure due to its status as the number one cause of mortality worldwide. In humans, a pathologic occlusion forms in a coronary artery and reperfusion of that occluded artery is considered essential to maintain viability of the myocardium at risk. Although essential for myocardial recovery, reperfusion of the ischemic myocardium creates its own tissue injury. The physiologic response and healing of an ischemia/reperfusion injury is different from a chronic occlusion injury. Myocardial ischemia/reperfusion injury is gaining recognition as a clinically relevant model for myocardial infarction studies. For this reason, parallel animal models of ischemia/reperfusion are vital in advancing the knowledge base regarding myocardial injury. Typically, ischemia of the mouse heart after left anterior descending (LAD) coronary artery occlusion is confirmed by visible pallor of the myocardium below the occlusion (ligature). However, this offers only a subjective way of confirming correct or consistent ligature placement, as there are multiple major arteries that could cause pallor in different myocardial regions. A method of recording electrocardiographic changes to assess correct ligature placement and resultant ischemia as well as reperfusion, to supplement observed myocardial pallor, would help yield consistent infarct sizes in mouse models. In turn, this would help decrease the number of mice used. Additionally, electrocardiographic changes can continue to be recorded non-invasively in a time-dependent fashion after the surgery. This article will demonstrate a method of electrocardiographically confirming myocardial ischemia and reperfusion in real time.

During murine myocardial ischemia/reperfusion surgery, correct placement of the occluding ligature is typically confirmed by visible observation of myocardial pallor. Herein, a method of electrocardiographically confirming ischemia and reperfusion, to supplement observed myocardial pallor, is demonstrated in male C57Bl/6 mice.

Many animal models have been established for the study of myocardial remodeling and heart failure due to its status as the number one cause of mortality ...

Preclinical Model of Hind Limb Ischemia in Diabetic Rabbits

Peripheral vascular disease is a widespread clinical problem that affects millions of patients worldwide. A major consequence of peripheral vascular disease is the development of ischemia. In severe cases, patients can develop critical limb ischemia in which they experience constant pain and an increased risk of limb amputation. Current therapies for peripheral ischemia include bypass surgery or percutaneous interventions such as angioplasty with stenting or atherectomy to restore blood flow. However, these treatments often fail to the continued progression of vascular disease or restenosis or are contraindicated due to the overall poor health of the patient. A promising potential approach to treat peripheral ischemia involves the induction of therapeutic neovascularization to allow the patient to develop collateral vasculature. This newly formed network alleviates peripheral ischemia by restoring perfusion to the affected area. The most frequently employed preclinical model for peripheral ischemia utilizes the creation of hind limb ischemia in healthy rabbits through femoral artery ligation. In the past, however, there has been a strong disconnect between the success of preclinical studies and the failure of clinical trials regarding treatments for peripheral ischemia. Healthy animals typically have robust vascular regeneration in response to surgically induced ischemia, in contrast to the reduced vascularity and regeneration in patients with chronic peripheral ischemia. Here, we describe an optimized animal model for peripheral ischemia in rabbits that includes hyperlipidemia and diabetes. This model has reduced collateral formation and blood pressure recovery in comparison to a model with a higher cholesterol diet. Thus, the model may provide better correlation with human patients with compromised angiogenesis from the common co-morbidities that accompany peripheral vascular disease.

We describe a surgical procedure used to induce peripheral ischemia in rabbits with hyperlipidemia and diabetes. This surgery acts as a preclinical model for conditions experienced in peripheral artery disease in patients. Angiography is also described as a means to measure the extent of introduced ischemia and recovery of perfusion.

Peripheral vascular disease is a widespread clinical problem that affects millions of patients worldwide. A major consequence of peripheral vascular ...

Evaluation of Coronary Flow Reserve After Myocardial Ischemia Reperfusion in Rats

Coronary artery disease is the leading cause of death worldwide. After an acute myocardial infarction, early and successful myocardial intervention via recanalization of the coronary artery is the most effective strategy for reducing the size of ischemic myocardium. The coronary microvasculature cannot be visualized and imaged&#160;in vivo, but there are several invasive and noninvasive techniques that can be used to assess parameters which depend directly on coronary microvascular function. The endothelial function after ischemia reperfusion can be assessed also at the level of the coronary circulation via the coronary flow reserve (CFR).&#160;In this study, peak velocity of left anterior descending (LAD) coronary arteries was measured in rats in vivo via Transthoracic Doppler Echocardiography during resting and stress challenge (induced by Dobutamine). A normal heart can increase its coronary blood flow up to four times above the resting values during stress induction. Following ischemia reperfusion, we found a significantly diminished CFR, which can be used as a marker of coronary microvascular dysfunction. CFR has opened a window on the importance of microvascular dysfunction and has been shown to predict cardiovascular risk independent of whether the severe obstructive disease is present.

Coronary flow reserve (CFR), is defined as the ratio of maximal coronary blood flow to the resting coronary blood flow. We present a protocol for evaluating CFR in rats via ultrasound, which offers the opportunity to predict cardiovascular risk factors in the absence of obstructive coronary disease.

Coronary artery disease is the leading cause of death worldwide. After an acute myocardial infarction, early and successful myocardial intervention via ...

A Pre-clinical Rat Model for the Study of Ischemia-reperfusion Injury in Reconstructive Microsurgery

Ischemia-reperfusion injury is the main cause of flap failure in reconstructive microsurgery. The rat is the preferred preclinical animal model in many areas of biomedical research due to its cost-effectiveness and its translation to humans. This protocol describes a method to create a preclinical free skin flap model in rats with ischemia-reperfusion injury. The described 3 cm x 6 cm rat free skin flap model is easily obtained after the placement of several vascular ligatures and the section of the vascular pedicle. Then, 8 h after the ischemic insult and completion of the microsurgical anastomosis, the free skin flap develops the tissue damage. These ischemia-reperfusion injury-related damages can be studied in this model, making it a suitable model for evaluating therapeutic agents to address this pathophysiological process. Furthermore, two main monitoring techniques are described in the protocol for the assessment of this animal model: transit-time ultrasound technology and laser speckle contrast analysis.

Here, we describe a pre-clinical animal model for studying the pathophysiology of ischemia-reperfusion injury in reconstructive microsurgery. This free skin flap model based on the superficial caudal epigastric vessels in the rat may also allow for the evaluation of different therapies and compounds to counteract ischemia-reperfusion injury-related damage.

Ischemia-reperfusion injury is the main cause of flap failure in reconstructive microsurgery. The rat is the preferred preclinical animal model in many ...

Delayed Intramyocardial Delivery of Stem Cells after Ischemia Reperfusion Injury in a Murine Model

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, cardiac stem cell therapy is studied by delivering SCs concurrently with the induction of myocardial injury. However, this approach presents two significant limitations: the early hostile pro-inflammatory ischemic environment may affect the survival of transplanted SCs, and it does not represent the subacute infarction scenario where SCs will likely be used. Here we describe a two-part series of surgical procedures for the induction of ischemia-reperfusion injury and delivery of mesenchymal stem cells (MSCs). This method of stem cell administration may allow for the longer viability and retention around damaged tissue by circumventing the initial immune response. A model of ischemia reperfusion injury was induced in mice accompanied by the delivery of mesenchymal stem cells (3.0 x 105), stably expressing the reporter gene firefly luciferase under the constitutively expressed CMV promoter, intramyocardially 7 days later. The animals were imaged via ultrasound and bioluminescent imaging for confirmation of injury and injection of cells, respectively. Importantly, there was no added complication rate when performing this two-procedure approach for SC delivery. This method of stem cell administration, collectively with the utilization of state-of-the-art reporter genes, may allow for the in vivo study of viability and retention of transplanted SCs in a situation of chronic ischemia commonly seen clinically, while also circumventing the initial pro-inflammatory response. In summary, we established a protocol for the delayed delivery of stem cells into the myocardium, which can be used as a potential new approach in promoting regeneration of the damaged tissue.

Stems cells are continuously investigated as potential treatments for individuals with myocardial damage, however, their decreased viability and retention within injured tissue can impact their long-term efficacy. In this manuscript we describe an alternative method for stem cell delivery in a murine model of ischemia reperfusion injury.

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, ...

Organ Ischemia-Reperfusion Injury by Simulating Hemodynamic Changes in Rat Liver Transplant Model

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, including ischemia-reperfusion injury (IRI) of extrahepatic organs. This model requires numerous experiments and devices. The duration of anhepatic phase is closely related to the time to develop IRI after transplantation. In this experiment, we used hemodynamic changes to induce extrahepatic organ damage in rats and determined the maximum tolerance time. The time until the most severe organ injury varied for different organs. This method can easily be replicated and can also be used to study IRI of the extrahepatic organs after liver transplantation.

This paper provides a detailed description of how to build an animal model of the anhepatic phase (liver ischemia) in rats to facilitate basic research into ischemia-reperfusion injury after liver transplantation.

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, ...

A Rat Lung Transplantation Model of Warm Ischemia/Reperfusion Injury: Optimizations to Improve Outcomes

From our experience with rat lung transplantation, we have found several areas for improvement. Information in the existing literature regarding methods for choosing appropriate cuff sizes for the pulmonary vein (PV), pulmonary artery (PA), or bronchus (Br) are varied, thus making the determination of proper cuff size during rat lung transplantation an exercise of trial and error. By standardizing the cuffing technique to use the smallest effective cuff appropriate for the size of the vessel or bronchus, one can make the transplantation procedure safer, faster, and more successful. Since diameters of the PV, PA, and Br are related to the body weight of the rat, we present a strategy to choosing an appropriate size using a weight-based guide. Since lung volume is also related to body weight, we recommend that this relationship should also be considered when choosing the proper volume of air for donor lung inflation during warm ischemia as well as for the proper volume of PBS to be instilled during bronchoalveolar lavage (BAL) fluid collection. We also describe methods for 4th intercostal space dissection, wound closure, and sample collection from both the native and transplanted lobes.

Here, we present optimizations to a rat lung transplantation model that serve to improve outcomes. We provide a size guide for cuffs based on body weight, a measurement strategy to ascertain the 4th intercostal space, and methods of wound closure and BAL (bronchoalveolar lavage) fluid and tissue collection.

From our experience with rat lung transplantation, we have found several areas for improvement. Information in the existing literature regarding methods ...