The studies described here have been funded by grants awarded to P.S Macdonald from the National Health and Medical Research Council Australia, the St Vincent&#39;s Clinic Foundation, and the NSW Health Cardiovascular Research Capacity Senior Investigator Grant.

All animals received humane care in compliance with the National Health and Medical Research Council (Australia) guidelines. All animal procedures were approved by the Animal Ethics Committee of the Garvan Institute of Medical Research (Sydney, Australia).
1. Preparation of Krebs-Henseleit (KH) buffer
<ol>
	<li>Prepare 2 L of a 20x stock solution of KH buffer containing 2.36 M NaCl, 0.094 M KCl, 0.024 M MgSO4, 0.024 M KH2PO4. Aliquot into 50 mL tubes and store at -20 &#176;C.</li>
	<li>Prior to the experiment, prepare two 2 L Erlenmyer flasks of 1x KH buffer containing 1900 mL of deionized water, 100 mL of 20x stock KH buffer, 4.20 g of NaHCO3 (25 mM), and 3.96 g of glucose (11 mM).
	<ol>
		<li>Place buffer flasks in a 37 &#176;C water bath and gas the solution with carbogen (95% oxygen, 5% carbon dioxide, flow rate set to 2 L/min) via air stones (slip-on inlet filter) for 5 min, then add 2.8 mL of 1 M CaCl2 (1.4 mM). Continue bubbling the solution for at least 1 h prior to the start of the experiment.&#160;Use of an air stone assists with uniform aeration of the buffer.</li>
		<li>Place a small volume (~50 mL) of buffer into a round metal dish at -20 &#176;C to cool and form a layer of iced buffer.</li>
	</ol>
	</li>
</ol>
2. Preparation of perfusion circuit
<ol>
	<li>Ensure all tubing and glassware are clean with no visible growth/contamination.</li>
	<li>Set up heights of glass chambers to correspond to required perfusion pressures based on 1 cm H2O being equivalent to 1.36 mmHg: Langendorff-to-heart 100 cm (approximately 75 mmHg), pre-load working side-to-heart 15 cm (approximately 11 mmHg), heart-to-afterload chamber 100 cm (approximately 75 mmHg).</li>
	<li>Start circulation of hot water around the circuit at least 10 min before heart perfusion.</li>
	<li>Place microfiber filters (1.2 &#181;m) into filter holders and secure them tightly.</li>
	<li>Prime the circuit with KH buffer, ensuring all air is removed, and use a flow control tubing clamp to adjust the flow on the Langendorff side (Figure 1A, B) to ~50 mL/min. The slower flow allows for ease of aortic cannulation and prevents the heart from being dislodged before tying. Place a thermometer within the circuit via a T-shaped tubing connector to monitor the temperature of the buffer at ~37 &#176;C.</li>
</ol>
3. Setting up of Lab Chart (data analysis) software
<ol>
	<li>Ensure Powerlab (data acquisition system) and flowmeter are switched on and connected to the computer prior to opening the data analysis software.</li>
	<li>Open a new file.</li>
	<li>Set up data collection by selecting the Settings drop-down menu and selecting Channel Settings. Select for Aortic Pressure, Aortic Flow, and Heart Rate (Figure 2A).</li>
	<li>Set up each channel with the settings shown in Figure 2B.</li>
</ol>
4. Preparation of animal for rat heart isolation
NOTE: The methodology for rat heart isolation and perfusion for non-DBD and non-DCD hearts for cold static storage is outlined below. The methods for heart instrumentation onto the perfusion circuit are relatively similar regardless of which donation method is used. The differences in the protocol for DBD and DCD rat heart donation primarily occur from the time the animal anesthesia has taken effect to when the heart is excised from the animal.
<ol>
	<li>Weigh the animal, ensuring that its weight is &#62;330 g. The ideal weight range for male rats is 330-420 g.</li>
	<li>Administer an intraperitoneal injection of ketamine (80 mg/kg) and xylazine (10 mg/kg). Ensure adequate surgical plane of anesthesia based on the absence of a pedal reflex (toe-pinch reflex) and eye blink reflex.</li>
	<li>Place the animal in a supine position, securing the limbs with tape. Administer oxygen to the animal via a nose cone.</li>
	<li>Have the KH buffer ice-slush metal container and a small metal container with ice at the dissecting station. Soak a 7.5 cm x 7.5 cm gauze in KH buffer and place it on the container with ice.</li>
	<li>Once adequate surgical plane of anesthesia is confirmed, shave the skin at the surgical incision site and clean the skin with aseptic such as betadine or 70% alcohol. Make a lateral incision across the abdomen (laparotomy) and move the stomach and intestines to the side to allow access to the left renal vein. Inject 1500 IU heparin and apply pressure to the injection site until the bleeding stops. Allow 1-2 min for heparin to circulate before continuing with heart retrieval.</li>
	<li>Open the chest cavity by cutting through the rib cage on either side and cutting across the diaphragm. Carefully hold and lift the heart-lung bloc while cutting along the descending aorta, ensuring that the aortic arch remains intact. This will allow for enough aorta length to remain for cannulation.</li>
	<li>The rat is euthanized by the removal of the heart and lungs while under anesthesia. Submerge the heart-lung bloc in the KH buffer ice slush to allow the heart to arrest.</li>
</ol>
5. Cannulating the heart onto the isolated working heart perfusion circuit
<ol>
	<li>Transfer the heart-lung bloc to a gauze on ice. Dissect excess fat around the aorta, trim the aorta to leave 3-5 mm length for cannulation, and make a small incision ~1-2 mm in the pulmonary artery. This will allow a path for the coronary effluent.</li>
	<li>Place the trimmed heart-lung bloc (while still on ice) directly underneath the cannulas. Adjust the flow from the Langendorff side using a flow control tubing clamp to facilitate aortic cannulation. Carefully cannulate the aorta onto the left cannula, hold it with a black clamp, and tie it with a 2-0 silk suture.</li>
	<li>Increase flow from Langendorff perfusion by completely opening the flow control tubing clamp. Open the afterload clamp. Ensure that there is continuous flow from the Langendorff chamber. Start monitoring heart activity with software. 
	NOTE: if the aortic flow value on the Flowmeter display is &#60;10 mL/min, ensure that the pulmonary arteriotomy is of sufficient size. A low aortic flow trace/value could also indicate that the aortic cannula has been inserted too far.</li>
	<li>Adjust the heart-lung bloc so that the anterior surface of the lungs is facing the operator. Tie the left lung lobe with a 2-0 silk suture and cut off excess left lung tissue. Tie the remaining lung lobes in one sweep with a 2-0 silk suture and cut the tissue. Rotate the heart so that the tied lungs are towards the back.</li>
	<li>Using small Vannas scissors, cut a 2-3 mm edge of the left atrial appendage, ensuring that a small opening into the atria is achieved. Carefully cannulate the left atrial appendage to the left cannula, hold it with a black clamp, and tie it with a 2-0 silk suture. Figure 3A shows the placement of both cannulas.</li>
	<li>Adjust the heart chamber water jacket so that the heart sits within the center of the chamber.</li>
	<li>Open the pressure transducer to air and zero the pressure transducer in the software by selecting the Aortic Pressure drop-down menu &#62; Bridge Amp &#62; Zero. Click OK.</li>
	<li>Perfuse the heart in Langendorff mode for 10 min. Ensure that the Langendoff aortic flow measures between 10-20 mL/min (ideal 20 mL/min).</li>
	<li>Prior to switching the working mode, ensure that the air stone (slip-on inlet filter) from the Langendorff KH buffer reservoir is placed in the working KH buffer reservoir.</li>
	<li>Switch the heart into working mode by closing the large clamp leading to the aortic cannula and opening the flow leading to the left atrial appendage. Open three-way tap leading from working perfusate. Transfer the common return tubing (taking KH buffer back to the reservoir) from the Langendorff reservoir to the working reservoir.</li>
	<li>Perfuse the heart in working mode for 15 min. Aortic flow values must ideally be &#62;30 mL/min at the end of 15 min perfusion. Once the heart has stabilized, start recording data in data analysis software.</li>
	<li>Measure coronary effluent (coronary flow) at 1 min, 5 min, 10 min, and 15 min. 
	NOTE: Coronary flow values should be between 10-20 mL/min. Coronary flow greater than 22 mL/min generally indicates a leak originating from an untied pulmonary vessel. Heart rate should be between 200-300 bpm. Coronary effluent can be collected for downstream analysis of tissue injury markers (e.g., troponin-I, lactate dehydrogenase release).</li>
</ol>
6. Administering cardioplegia for cold static storage of the heart
<ol>
	<li>Fill the cardioplegia chamber (positioned at a height of 60 cm to yield perfusion pressure of approximately 40 mmHg) with 50 mL of designated cardiac preservation solution (with or without supplementation).</li>
	<li>Lower the water jacket and close the three-way tap on the working mode side.</li>
	<li>Stop the flow towards the left atrial cannula by closing the clamp.</li>
	<li>Open the flow to the aortic cannula.</li>
	<li>Close the clamp from the Langendoff side and on the afterload.</li>
	<li>Open the clamp from the cardioplegia line and flush 15 mL of cardioplegia out from the pressure line.&#160;Close the tap at the pressure&#160;line.</li>
	<li>Allow the heart to be flushed with cardioplegia via the aortic cannula and collect the coronary effluent. Run cardioplegia for 3 min and record the volume of cardioplegia administered.</li>
	<li>Stop recording data in the software and save the file.</li>
	<li>After 3 min, close the cardioplegia clamp and the small blue clamp leading to the aortic cannula.</li>
	<li>Place a bulldog clamp on the aortic cannula and a second bulldog clamp on the left cannula. Disconnect the heart-cannula block from the circuit and place it in a beaker containing cardioplegia in an ice box (Figure 3B, C).</li>
	<li>Store the heart in the fridge for 6 h (or desired cold storage time).</li>
</ol>
7. Heart reperfusion after cold static storage
<ol>
	<li>Prepare KH buffer as per section 1.</li>
	<li>Prime the perfusion circuit as per section 2.</li>
	<li>Open the saved file from baseline perfusion.</li>
	<li>Carefully remove the heart-cannula block from the ice box and secure it onto the perfusion circuit. Before reconnecting the aortic cannula, use a blunt-end 18 G syringe filled with KH buffer to remove any air from the cannula. Connect the aortic cannula first, open the tubing clamp on the Langendorff side, and ensure the afterload tubing clamp is open.</li>
	<li>Reconnect the left atrial cannula, ensuring that air has been removed from the cannula before connecting to the perfusion circuit. Keep the working side clamp closed until it is time to switch from Langendorff to working mode.</li>
	<li>Perfuse the heart in Langendorff mode for 15 min. Ensure to start recording data at the start of Langendorff reperfusion.</li>
	<li>Prior to switching to the working mode, place both bubbling stones in the KH buffer, open the 3-way tap on the working side, and transfer common return tubing to the working reservoir.</li>
	<li>Close the white Langendorff clamp, open the white working side clamp, and perfuse the heart for 30 min. Measure and collect coronary effluent at designated time points for downstream analysis (e.g., cardiac injury markers). Ensure to stop-restart software recording to start a new segment (or alternatively, add a comment).</li>
	<li>At approximately 15 min working mode reperfusion, close the three-way tap.</li>
	<li>At the end of reperfusion, save the data file, stop the flow through the circuit, and disconnect the heart block. Collect left ventricle samples for histological processing and/or snap frozen for future studies (e.g., protein extraction for western blot studies).</li>
	<li>Remove the filters from the filter holders, drain the buffer from the circuit, and flush twice with 5 L of distilled water. 
	NOTE: Although not necessary, it does help to pre-warm the distilled water in a large container within the water bath.</li>
</ol>
8. Analysis of functional recovery
<ol>
	<li>Open the LabChart file.</li>
	<li>Highlight a region of at least 30 s of recording, then click Command &#62; Add data selection OR Multiple add to data. This will add the desired parameters to a DataPad file within the software.</li>
	<li>Select Window drop-down menu &#62; DataPad.</li>
	<li>Highlight and copy the recovery values from DataPad to a spreadsheet for functional recovery analysis. Calculate functional recovery as a percentage of its respective baseline value1,6,7.</li>
</ol>

Optimization of Donor Heart Preservation Using a Working Rat Heart Perfusion Model

<ol>
	<li>Kwan, J. C., Gao, L., Macdonald, P. S., Hicks, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardio-protective+signalling+by+glyceryl+trinitrate+and+cariporide+in+a+model+of+donor+heart+preservation.">Cardio-protective signalling by glyceryl trinitrate and cariporide in a model of donor heart preservation.</a> Heart Lung Circ. 24 (3), 306-318 (2015).</li><li>Watson, A. J., 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=Enhanced+preservation+of+the+rat+heart+after+prolonged+hypothermic+ischemia+with+erythropoietin-supplemented+Celsior+solution.">Enhanced preservation of the rat heart after prolonged hypothermic ischemia with erythropoietin-supplemented Celsior solution.</a> J Heart Lung Transplant. 32 (6), 633-640 (2013).</li><li>Chew, H. C., 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=Outcomes+of+donation+after+circulatory+death+heart+transplantation+in+Australia.">Outcomes of donation after circulatory death heart transplantation in Australia.</a> J Am Coll Cardiol. 73 (12), 1447-1459 (2019).</li><li>Dhital, K. K., 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=Adult+heart+transplantation+with+distant+procurement+and+ex-vivo+preservation+of+donor+hearts+after+circulatory+death:+a+case+series.">Adult heart transplantation with distant procurement and ex-vivo preservation of donor hearts after circulatory death: a case series.</a> Lancet. 385 (9987), 2585-2591 (2015).</li><li>Joshi, Y., 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=Heart+transplantation+from+DCD+donors+in+Australia:+Lessons+learned+from+the+first+74+cases.">Heart transplantation from DCD donors in Australia: Lessons learned from the first 74 cases.</a> Transplantation. 107 (2), 361-371 (2023).</li><li>Villanueva, J. E., 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=Functional+recovery+after+dantrolene-supplementation+of+cold+stored+hearts+using+an+ex+vivo+isolated+working+rat+heart+model.">Functional recovery after dantrolene-supplementation of cold stored hearts using an ex vivo isolated working rat heart model.</a> PLoS One. 13 (10), e205850 (2018).</li><li>Villanueva, J. E., 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=The+cardioprotective+potential+of+the+sodium-glucose+cotransporter+2+inhibitor+empagliflozin+in+donor+heart+preservation.">The cardioprotective potential of the sodium-glucose cotransporter 2 inhibitor empagliflozin in donor heart preservation.</a> J Heart Lung Transplant. 39 (10), 1151-1153 (2020).</li><li>Gao, L., 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=Critical+role+of+the+STAT3+pathway+in+the+cardioprotective+efficacy+of+zoniporide+in+a+model+of+myocardial+preservation+-+the+rat+isolated+working+heart.">Critical role of the STAT3 pathway in the cardioprotective efficacy of zoniporide in a model of myocardial preservation - the rat isolated working heart.</a> Br J Pharmacol. 162 (3), 633-647 (2011).</li><li>Cropper, J. R., Hicks, M., Ryan, J. B., Macdonald, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardioprotection+by+cariporide+after+prolonged+hypothermic+storage+of+the+isolated+working+rat+heart.">Cardioprotection by cariporide after prolonged hypothermic storage of the isolated working rat heart.</a> J Heart Lung Transplant. 22 (8), 929-936 (2003).</li><li>Gao, L., 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=Enhanced+functional+recovery+of+the+heart+donated+after+circulatory+death+determination+with+antemortem+heparin.">Enhanced functional recovery of the heart donated after circulatory death determination with antemortem heparin.</a> J Heart Lung Transplant. 39 (6), 607-609 (2020).</li><li>Villanueva, J. E., 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=The+effect+of+increasing+donor+age+on+myocardial+ischemic+tolerance+in+a+rodent+model+of+donation+after+circulatory+death.">The effect of increasing donor age on myocardial ischemic tolerance in a rodent model of donation after circulatory death.</a> Transplant Direct. 7 (6), e699 (2021).</li><li>Kumarasinghe, G., 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=Improved+heart+function+from+older+donors+using+pharmacologic+conditioning+strategies.">Improved heart function from older donors using pharmacologic conditioning strategies.</a> J Heart Lung Transplant. 35 (5), 636-646 (2016).</li></ol>

The authors have no conflicts of interest to disclose.

Given the sensitive nature of heart baseline function, care must be taken to maintain the perfusion rig clean and with compatible components. For example, the correct PVC tubing must be used. Some tubing materials, such as silicone, can contribute to lower aortic flow and contractility, which may be due to loss of oxygen via the tubing. While there are isolated working rat heart perfusion circuits commercially available, most of these are primarily used for Langendorff perfusion only. Generally, functional measurements can be obtained in a Langendoff perfusion model via the insertion of a balloon catheter into the left ventricle to obtain left ventricular diastolic pressure (LVDP) measurements. However, the ability to measure raw aortic flow measurements and cardiac output in a Langendoff perfusion system is not possible. A main advantage of the IWRH protocol is the ability to obtain aortic flow and cardiac output functional measurements without the need for additional interventions of the heart (i.e., balloon catheter). There are commercial options for a working rat heart perfusion system and although these can be customizable for individual study requirements, these systems can be costly.
There are critical steps in the protocol to ensure optimal baseline function of the rodent heart during perfusion. Preparation of KHB buffer stock solutions should be no older than 4 weeks. The perfusion circuit must be free of any air bubbles when priming. Care must be taken when cannulating the left atrial appendage. Excessive handling of the tissue can increase the risk of tearing and can also add to functional irritability of the heart.
The IWRH perfusion circuit performance quality can generally be determined by the baseline readings obtained in hearts (AF &#62; 30 mL/min, CF ~20 mL/min). Further quality control testing in a CSS model can be performed by arresting and storing the heart in Celsior preservation solution supplemented with 1 &#181;mol/L zoniporide plus 0.1 mg/mL glyceryl trinitrate. Hearts arrested with Celsior supplemented with glyceryl trinitrate plus zoniporide guarantees immediate Langendorff reperfusion recovery and a final functional recovery between 40-60% after 6 h cold storage. If in doubt as to whether the rig is performing well, this quality control supplementation is useful to test.
The methods for using rodent hearts procured via donation after circulatory death have been previously described10,11. Briefly, rats are anesthetized, the carotid artery cannulated, and pressure readings obtained. Heparin (500 IU) is injected via the carotid artery cannula. The trachea is ligated to initiate withdrawal of life support. Once the required warm ischemic time is reached, the heart can be excised and cannulated onto the perfusion circuit. For administration of preservation solution into the DCD heart, 60 mL of preservation solution is flushed through the aorta (~20 mL/min) via the pressure line on the circuit. The pulmonary vessels are tied, the lungs are dissected, the left cannula is inserted, and then 40 mL of preservation solution is flushed via the pressure line. The heart is then perfused as per the protocol above. Functional recovery is taken as a raw value of functional hemodynamics, and statistical analysis is performed by comparing the values to those hearts that were sham-operated (no DCD, however, flushed on the rig).
The methods for using rodent hearts procured after brain death have been previously described12. Briefly, animals are anesthetized, intubated, and ventilated, with hemodynamic monitoring as per DCD rodent protocol. A burr hole is performed in the temporoparietal skull, and a 3F embolectomy catheter is placed sub-durally. Sham controls have burr holes but no catheter insertion. The balloon is inflated with 0.3 mL water over 2 min and kept inflated for 1 h. After 1 h of hemodynamic observation, hearts are explanted and placed onto the isolated working heart perfusion circuit, baseline measurements are recorded, cardioplegia is administered, and hearts are cold stored as described above.
The main limitation of using the IWRH perfusion circuit is the inability to use a blood-based perfusion buffer to circulate through the system. It would be impossible to source enough rodent blood to add to the perfusate; therefore, blood-based perfusion is not feasible. Alternative models, such as rodent heterotopic heart transplant or large animal orthotopic heart transplant studies, would likely need to be performed to test the impact of blood-based perfusion. However, these alternative methods require expertise in microsurgical skills and/or large animal surgical facilities and experienced anesthetists, perfusionists, and cardiothoracic surgeons.

The success of a heart transplant can be greatly influenced by the preservation method used for the donor heart. Donor hearts are subjected to ischemic insults throughout the organ retrieval, transport, and transplantation process. The degree of ischemic damage to the donor organ can be mitigated by selecting an appropriate donor heart preservation strategy. Cold static storage (CSS) remains the most feasible and common method for donor heart preservation; however, CSS may not necessarily be the best option for all heart donation pathways. For example, hearts procured via the donation after circulatory death (DCD) pathway are maintained and assessed for viability using normothermic machine perfusion, whereas hearts procured via the donation after brain death (DBD) pathway can be preserved using CSS or hypothermic machine perfusion. Another important determinant for the preservation strategy used is the amount of either warm ischemia (for DCD) or cold ischemia (for DBD) the heart is subjected to during organ procurement and transportation.
Many centers use additional pharmacological supplementation of the heart preservation solution to increase ischemic tolerance and improve post-transplant heart function. Rodent studies from our laboratory showed that supplementation of cardioplegia with erythropoietin and glyceryl trinitrate was able to significantly improve cardiac functional recovery after prolonged cold storage of the donor heart1,2. These studies then progressed to porcine studies of donor heart preservation and provided the pre-clinical groundwork that led to the incorporation of erythropoietin and glyceryl trinitrate into the cardioplegia solution of hearts procured via a DCD pathway utilizing normothermic machine perfusion3,4,5. Currently, supplementation with erythropoietin and glyceryl trinitrate is also used in Australian transplant units for clinical DBD heart retrievals incorporating CSS. The overall goal of the isolated working rat heart (IWRH) perfusion protocol outlined below is to test donor heart preservation strategies in a laboratory setting and test their efficacy for preserving donor heart function after reperfusion. The isolated working rat heart protocol described has been used for over two decades within our laboratory and has proven extremely useful in screening for potential supplements and/or strategies that can improve donor heart preservation2,7,8,9.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.5 mL colorless Eppendorff tube, 1000 per box</td>
			<td>Bio Strategy Pty Ltd</td>
			<td>0030.125.150</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL cent/tubes (S) Sleeve/25 PP ctn/500</td>
			<td>Sigma-aldrich Pty Ltd</td>
			<td>CLS430791-500ea</td>
			<td></td>
		</tr>
		<tr>
			<td>BP Transducer/Cable kit</td>
			<td>ADInstruments</td>
			<td>MLT1199</td>
			<td>Data Acquisition</td>
		</tr>
		<tr>
			<td>Bridge Amp</td>
			<td>ADInstruments</td>
			<td>FE221</td>
			<td>Data Acquisition</td>
		</tr>
		<tr>
			<td>Carbogen 555G2</td>
			<td>BOC</td>
			<td>BOC002</td>
			<td></td>
		</tr>
		<tr>
			<td>Checktemp1 thermometer</td>
			<td>Hanna Instruments</td>
			<td>HI98509</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Clamp Pinch 1/4-7/16 PK 12</td>
			<td>Thomas Scientific</td>
			<td>2848Y40</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Clamp W/Extension Stem Med.</td>
			<td>Thomas Scientific</td>
			<td>8847T08</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Clamp W/Extension Stem Sm.</td>
			<td>Thomas Scientific</td>
			<td>8847T02</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Clips, Vessel, 60 g Pressure</td>
			<td>Coherent Scientific</td>
			<td>14121</td>
			<td>Surgical Equipment</td>
		</tr>
		<tr>
			<td>Closed Connector</td>
			<td>Thomas Scientific</td>
			<td>8847E25</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Covidien Sofsilk 2-0 black precut 45 cm box of 24</td>
			<td>Specialist Medical Supplies</td>
			<td>S195</td>
			<td>Surgical consumables</td>
		</tr>
		<tr>
			<td>Custom Made 250 mL Jacketed Degasser</td>
			<td>Custom Blown Glassware Pty Ltd</td>
			<td>N/A</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Custom Made 750 mL Reservoir</td>
			<td>Custom Blown Glassware Pty Ltd</td>
			<td>N/A</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>D-(+)-Glucose Anhydrous SigmaUltra</td>
			<td>Sigma-aldrich Pty Ltd</td>
			<td>G7528-1Kg</td>
			<td>To make Krebs Buffer</td>
		</tr>
		<tr>
			<td>Dual Channel Console</td>
			<td>ADInstruments</td>
			<td>TS402</td>
			<td>Data Acquisition</td>
		</tr>
		<tr>
			<td>Erlenmyer flasks 2 L&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Filter Microfibre type GF/C glass fibre 47 mm, 100</td>
			<td>Bio-strategy Pty Ltd</td>
			<td>1822047</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps, 15 cm, 0.3 mm, CRVD</td>
			<td>Coherent Scientific</td>
			<td>14114</td>
			<td>Surgical Equipment</td>
		</tr>
		<tr>
			<td>Four Prong Clamps with 9 mm x 115 mm long arm for holding 2-70 mm diam objects. Vinyl coated</td>
			<td>Met-App Australia Pty Ltd</td>
			<td>1352</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Heater Circulator. Digital Solid State Control. (1020 Watts/240 Volts)</td>
			<td>Thermoline Scientific</td>
			<td>TU3</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Heparin 5000 U/5 mL box 50 Pfizer 02112115</td>
			<td>Clffird Hallam Healthcare Pty Ltd</td>
			<td>1258693</td>
			<td>Drugs</td>
		</tr>
		<tr>
			<td>Ilium Xylazil 20 Inj 50 mL</td>
			<td>Cenvet Australia Pty Ltd</td>
			<td>X5010</td>
			<td>Anaesthetic</td>
		</tr>
		<tr>
			<td>Johns Hopkins Bulldog Clamp</td>
			<td>Coherent Scientific&#160;</td>
			<td>CS-WPI-14117</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine 100 mg/50 mL</td>
			<td>Provet (NSW) Pty Ltd</td>
			<td>KETAI1</td>
			<td>Anaesthetic</td>
		</tr>
		<tr>
			<td>Magnesium Sulphate heptahydrate AR 500 g Chemsupply</td>
			<td>Bio Strategy Pty Ltd</td>
			<td>MA048-500g</td>
			<td>To make Krebs Buffer</td>
		</tr>
		<tr>
			<td>Male/Female Hinged Adapter</td>
			<td>Thomas Scientific</td>
			<td>8847V08</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Masterflex L/S Easy-Load Head for Precision Tubing, PPS, CRS Rotor</td>
			<td>John Morris</td>
			<td>1015164</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Metzenbaum scissors, 11.5 cm curved</td>
			<td>Coherent Scientific</td>
			<td>WPI-501748</td>
			<td>Surgical Equipment</td>
		</tr>
		<tr>
			<td>Metzenbaum scissors, 14.5 cm straight</td>
			<td>Coherent Scientific</td>
			<td>WPI-501252</td>
			<td>Surgical Equipment</td>
		</tr>
		<tr>
			<td>Mounting Hardware F/2-HEADS SS</td>
			<td>John Morris</td>
			<td>1014414</td>
			<td>Mounting screws for pump heads</td>
		</tr>
		<tr>
			<td>Open-sided connector</td>
			<td>Thomas Scientific</td>
			<td>8847E05</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Paraformadehyde</td>
			<td>Sigma-Aldrich</td>
			<td>P6148-500G</td>
			<td>Sample processing</td>
		</tr>
		<tr>
			<td>Potassium Chloride (AnalaR NORMAPUR) 500 g</td>
			<td>VWR Chemicals&#160;</td>
			<td>26764.26</td>
			<td>To make Krebs Buffer</td>
		</tr>
		<tr>
			<td>Potassium phosphate monobasic&#160;</td>
			<td>Sigma-Aldrich Pty Ltd</td>
			<td>P5379-500g</td>
			<td>To make Krebs Buffer</td>
		</tr>
		<tr>
			<td>Powerlab 2/26, 2 channel recorder + Labchart software</td>
			<td>ADInstruments</td>
			<td>ML826</td>
			<td>Computer Hardware and Software</td>
		</tr>
		<tr>
			<td>Precision XN Inline Flowsensor, 3.2 mm (1/8&#34;)ID ME4PXN-KR37 XF</td>
			<td>ADInstruments</td>
			<td>ME4PXN</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Scalpel with handle disposable #11 pkt/10</td>
			<td>BSN Medical (Aust) Pty Ltd</td>
			<td>73252-36</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicone Gasket for Swinnex 47 mm 5/PK</td>
			<td>Merck Millipore</td>
			<td>SX0004701</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Silicone O-Ring 5-329 10/PK</td>
			<td>Merck Millipore</td>
			<td>XX1104707</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Single Buret Clamp</td>
			<td>Thomas Scientific</td>
			<td>8847T32</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Slip-on inlet Filter pore size 10 &#181;m (bubbler)</td>
			<td>Sigma-aldrich</td>
			<td>59277</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Sm. 360 Rotation Connector</td>
			<td>Thomas Scientific</td>
			<td>8847E35</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma-Aldrich Pty Ltd</td>
			<td>S6297 - 250g</td>
			<td>To make Krebs Buffer</td>
		</tr>
		<tr>
			<td>Swinnex Filter Holder, 47 mm</td>
			<td>Merck Millipore</td>
			<td>SX0004700</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>syringes 1 mL box/100</td>
			<td>Becton Dickinson Pty Ltd</td>
			<td>302100</td>
			<td></td>
		</tr>
		<tr>
			<td>Three Prong Clamps with 9 mm diameter x 125 mm long arm and twin screw for holding 5-80 mm</td>
			<td>Met-App Australia Pty Ltd</td>
			<td>1356</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Tubing Flowmeter Module TS410</td>
			<td>ADInstruments</td>
			<td>TS410</td>
			<td>Data Acquisition</td>
		</tr>
		<tr>
			<td>Tubing PVC 6.35 mm ID x 9.52 mm OD 50ft Roll 15.24m, clear , DEHP phthalate free, food grade meets REAC</td>
			<td>Thermo Fisher</td>
			<td>NAL 8701-0600</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Tubing PVC 7.94mm ID x 11.1mm OD 50ft Roll 15.24 m, clear , DEHP phthalate free, food grade meets REAC</td>
			<td>Thermo Fisher</td>
			<td>NAL 8701-0900</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Tubing PVC 9.52 mm ID x 12.7 mm OD 100ft Roll</td>
			<td>Thermo Fisher</td>
			<td>NAL 8701-4120</td>
			<td>Rig Construction</td>
		</tr>
		<tr>
			<td>Vannas scissors, 8.5 cm, Straight, 7 mm Blades</td>
			<td>Coherent Scientific</td>
			<td>WPI-500-086</td>
			<td>Surgical Equipment</td>
		</tr>
		<tr>
			<td>Water Bath 30 Litre with Suspended Tray</td>
			<td>Thermoline Scientific</td>
			<td>TLWB-30</td>
			<td>Rig Construction</td>
		</tr>
	</tbody>
</table>

using an isolated working rat heart model to test donor heart preservation strategies

Optimization of donor heart preservation solutions has played a key role in reducing ischemia-reperfusion injury in donor hearts during organ retrieval, transportation, and transplantation. Previous work from our laboratory showed that the addition of glyceryl trinitrate and erythropoietin to donor heart preservation solutions could significantly improve cardiac functional recovery after prolonged cold storage and in donation after circulatory death hearts. This supplementation protocol has been implemented in clinical use in transplant units around Australia, Belgium, and the United Kingdom. Here, we outline a protocol for testing supplementation strategies using an ex vivo isolated working rat heart (IWRH) perfusion circuit. Using this methodology, supplementation strategies can be tested in the context of prolonged cold static storage, donation after brain death, and donation after circulatory death donor heart preservation. Cardiac functional recovery, measured by aortic flow, coronary flow, cardiac output, pulse pressure, and heart rate, can be used to determine whether a particular preservation strategy can minimize ischemia-reperfusion injury of the donor heart.

The results from the baseline perfusion will determine whether the initial experiment (pre-storage) was successful. The aortic flow displayed during Langendorff mode should be between 14 - 22 mL/min. Langendorff flow is displayed as a negative value on the flow meter due to the retrograde perfusion of the aorta. An example of an acceptable Langendorff trace is shown in Figure 4A.
Once switched to working mode, the aortic flow should be at least 30 mL/min once the heart settles. It is normal for the flow to initially surge to approximately 50 mL/min, then decrease to approximately 25 mL/min before increasing back to a stable baseline aortic flow. Once the baseline parameters are satisfactory (AF &#62; 30 mL/min, CF 10-20 mL/min, HR 200-300 bpm), the heart may be stored. An example of a good baseline trace is shown in Figure 4B.
Depending on the preservation solution tested, some hearts will not yield good recovery after prolonged cold static storage despite having an excellent baseline. An example of a trace indicating poor recovery in working mode is shown in Figure 4C.
The final functional recovery for aortic flow can be calculated based on the final aortic flow reading as a percentage of the baseline value. For example, if the heart baseline AF was 40 mL/min, and the AF at the end of the reperfusion after cold static storage was 20 mL/min, the AF recovery for this heart was 50%. An example of this data representation is shown in Figure 5.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66910/66910fig01.jpg" /> 
Figure 1: Working heart perfusion circuit. (A) Photo of the isolated working rat heart perfusion circuit. The red box indicates where the isolated heart is positioned. (B) Schematic of the isolated working heart perfusion circuit indicating the flow direction of KH buffer through the circuit for Langendorff (dotted line) and working mode (solid line) perfusion. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66910/66910fig01large.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/66910/66910fig02.jpg" /> 
Figure 2: LabChart software setup. (A) Setup of Channel Settings for Aortic Pressure, Aortic Flow, and Heart Rate. (B) Example of LabChart template set up with three channels. Windows on the right shows each individual channel set up. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66910/66910fig02large.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/66910/66910fig03.jpg" /> 
Figure 3: Cannulation and cardioplegia. (A) Heart cannulation prior to placing within the water jacket chamber. The left cannula is inserted into the aorta, and the right cannula is inserted into the left ventricle via the left atrial appendage. The pulmonary vessels have been tied, and the lung tissue dissected. (B) After cardioplegia has been administered, the heart is placed in a beaker containing the cardioplegia solution. The cannulas have been clamped with bulldog clips prior to disconnecting from the rig. (C) Example of hearts placed within an ice box for prolonged cold storage. Here, two hearts have been stored for reperfusion. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66910/66910fig03large.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/66910/66910fig04.jpg" /> 
Figure 4: Examples of LabChart traces. Traces for pressure (red graph), aortic flow (blue graph), and heart rate (green graph) for a heart with (A) Good Langendorff trace, (B) Good recovery during working mode reperfusion, (C) Poor recovery during working mode reperfusion. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66910/66910fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66910/66910fig05.jpg" /> 
Figure 5: Representative graphs for functional recovery. (A) aortic flow, (B) coronary flow, (C) cardiac output, and (D) heart rate. This figure has been reused with permission from Gao et al.8. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66910/66910fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Author Spotlight: Enhancing Donor Heart Preservation Through Isolated Rat Heart Perfusion Studies

We describe an ex vivo isolated working rat heart protocol to test donor heart preservation strategies. This paper describes the protocol for use in static cold storage of rodent donor hearts; however, the protocol can also be used for donor hearts obtained after donation after circulatory death and brain death.

Using an Isolated Working Rat Heart Model to Test Donor Heart Preservation Strategies

Optimization of donor heart preservation solutions has played a key role in reducing ischemia-reperfusion injury in donor hearts during organ retrieval, ...

using-an-isolated-working-rat-heart-model-to-test-donor-heart

Nanyang Technological University

Research

JoVE Journal

Medicine

353 Views.  Victor Chang Cardiac Research Institute. We describe an ex vivo isolated working rat heart protocol to test donor heart preservation strategies. This paper describes the protocol for use in static cold storage of rodent donor hearts; however, the protocol can also be used for donor hearts obtained after donation after circulatory death and brain death. 

Video: Author Spotlight: Enhancing Donor Heart Preservation Through Isolated Rat Heart Perfusion Studies

Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant vasculopathy (TVP).
The commonly used animal model for cardiovascular chronic rejection studies is the heterotopic heart transplant model performed in laboratory rodents. This model is used widely in experiments since Ono and Lindsey (3) published their technique. To analyze the findings in the blood vessels, the heart has to be sectioned and all vessels have to be measured.
Another method to investigate chronic rejection in cardiovascular questionings is the aortic transplant model (1, 2). In the orthotopic aortic transplant model, the aorta can easily be histologically evaluated (2). The PVG-to-ACI model is especially useful for CAV studies, since acute vascular rejection is not a major confounding factor and Cyclosporin A (CsA) treatment does not prevent the development of CAV, similar to what we find in the clinical setting (4). A7-day period of CsA is required in this model to prevent acute rejection and to achieve long-term survival with the development of TVP.
This model can also be used to investigate acute cellular rejection and media necrosis in xenogeneic models (5).

This video demonstrates the orthotopic aortic transplant model as a simple model to study the development of transplant vasculopathy (TVP) in rats.

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant ...

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 ...

An Isolated Working Heart System for Large Animal Models

Since its introduction in the late 19th century, the Langendorff isolated heart perfusion apparatus, and the subsequent development of the working heart model, have been invaluable tools for studying cardiovascular function and disease1-15. Although the Langendorff heart preparation can be used for any mammalian heart, most studies involving this apparatus use small animal models (e.g., mouse, rat, and rabbit) due to the increased complexity of systems for larger mammals1,3,11. One major difficulty is ensuring a constant coronary perfusion pressure over a range of different heart sizes &#8211; a key component of any experiment utilizing this device1,11. By replacing the classic hydrostatic afterload column with a centrifugal pump, the Langendorff working heart apparatus described below allows for easy adjustment and tight regulation of perfusion pressures, meaning the same set-up can be used for various species or heart sizes. Furthermore, this configuration can also seamlessly switch between constant pressure or constant flow during reperfusion, depending on the user&#8217;s preferences. The open nature of this setup, despite making temperature regulation more difficult than other designs, allows for easy collection of effluent and ventricular pressure-volume data.

Most studies involving the Langendorff apparatus use small animal models due to the increased complexity of systems for larger mammals. We describe a Langendorff system for large animal models that allows for use across a range of species, including humans, and relatively easy data acquisition.

Since its introduction in the late 19th century, the Langendorff isolated heart perfusion apparatus, and the subsequent development of the working heart ...

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. As this technology expands and improves, the ability to potentially evaluate and improve the quality of substandard lungs prior to transplant is a critical need. In order to more rigorously evaluate these approaches, a reproducible animal model needs to be established that would allow for testing of improved techniques and management of the donated lungs as well as to the lung-transplant recipient. In addition, an EVLP animal model of associated pathologies, e.g., ventilation induced lung injury (VILI), would provide a novel method to evaluate treatments for these pathologies. Here, we describe the development of a rat EVLP lung program and refinements to this method that allow for a reproducible model for future expansion. We also describe the application of this EVLP system to model VILI in rat lungs. The goal is to provide the research community with key information and &#8220;pearls of wisdom&#8221;/techniques that arose from trial and error and are critical to establishing an EVLP system that is robust and reproducible.

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. Here, we describe the development of a rat EVLP program and refinements that allow for a reproducible model for future expansion.

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed ...

Murine Isolated Heart Model of Myocardial Stunning Associated with Cardioplegic Arrest

The following protocol is of use to evaluate impaired cardiac function or myocardial stunning following moderate ischemic insults. The technique is useful for modeling ischemic injury associated with numerous clinically relevant phenomenon including cardiac surgery with cardioplegic arrest and cardiopulmonary bypass, off-pump CABG, transplant, angina, brief ischemia, etc. The protocol presents a general method to model hypothermic hyperkalemic cardioplegic arrest and reperfusion in rodent hearts focusing on measurement of myocardial contractile function. In brief, a mouse heart is perfused in langendorff mode, instrumented with an intraventricular balloon, and baseline cardiac functional parameters are recorded. Following stabilization, the heart is then subject to brief infusion of a cardioprotective hypothermic cardioplegia solution to initiate diastolic arrest. Cardioplegia is delivered intermittently over 2 hr. The heart is then reperfused and warmed to normothermic temperatures and recovery of myocardial function is monitored. Use of this protocol results in reliable depressed cardiac contractile function free from gross myocardial tissue damage in rodents.

The goal of this protocol to assess myocardial stunning following ischemic cardioplegic arrest in rodents.

The following protocol is of use to evaluate impaired cardiac function or myocardial stunning following moderate ischemic insults. The technique is useful ...

Vein Interposition Model: A Suitable Model to Study Bypass Graft Patency

Bypass grafting is an established treatment method for coronary artery disease. Graft patency continues to be the Achilles heel of saphenous vein grafts. Research models for bypass graft failure are essential for a better understanding of pathobiological and pathophysiological processes during graft patency loss. Large animal models, such as pigs or sheep, resemble human anatomical structures but require special facilities and equipment. This video describes a rat vein interposition model to investigate vein graft patency loss. Rats are inexpensive and easy to handle. Compared to mouse models, the convenient size of rats permits better operability and enables a sufficient amount of material to be obtained for further diverse analysis. In brief, the inferior epigastric vein of a donor rat is harvested and used to replace a segment of the femoral artery. Anastomosis is conducted via single stitches and sealed with fibrin glue. Graft patency can be monitored non-invasively using duplex sonography. Myointimal hyperplasia, which is the main cause for graft patency loss, develops progressively over time and can be calculated from histological cross sections.

This video demonstrates a model to study the development of myointimal hyperplasia after venous interposition surgery in rats.

Bypass grafting is an established treatment method for coronary artery disease. Graft patency continues to be the Achilles heel of saphenous vein grafts. ...

The Inverted Heart Model for Interstitial Transudate Collection from the Isolated Rat Heart

The present protocol describes a unique approach that enables the collection of cardiac transudate (CT) from the isolated, saline-perfused rat heart. After isolation and retrograde perfusion of the heart according to the Langendorff technique, the heart is inverted into an upside-down position and is mechanically stabilized by a balloon catheter inserted into the left ventricle. Then, a thin latex cap &#8211; previously cast to match the average size of the rat heart &#8211; is placed over the epicardial surface. The outlet of the latex cap is connected to silicon tubing, with the distal opening 10 cm below the base level of the heart, creating slight suction. CT continuously produced on the epicardial surface is collected in ice-cooled vials for further analysis. The rate of CT formation ranged from 17 to 147 &#181;L/min (n = 14) in control and infarcted hearts, which represents 0.1-1% of the coronary venous effluent perfusate. Proteomic analysis and high performance liquid chromatography (HPLC) revealed that the collected CT contains a wide spectrum of proteins and purinergic metabolites.

This protocol describes a method to collect cardiac interstitial fluid from the isolated, perfused rat heart. To physically separate interstitial transudate from coronary venous effluent perfusate, the Langendorff perfused heart is inverted, and the transudate (interstitial fluid) formed on the cardiac surface is collected using a soft latex cap.

The present protocol describes a unique approach that enables the collection of cardiac transudate (CT) from the isolated, saline-perfused rat heart. ...

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.

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 Rat Model of Pressure Overload Induced Moderate Remodeling and Systolic Dysfunction as Opposed to Overt Systolic Heart Failure

In response to an injury, such as myocardial infarction, prolonged hypertension or a cardiotoxic agent, the heart initially adapts through the activation of signal transduction pathways, to counteract, in the short-term, for the cardiac myocyte loss and or the increase in wall stress. However, prolonged activation of these pathways becomes detrimental leading to the initiation and propagation of cardiac remodeling leading to changes in left ventricular geometry and increases in left ventricular volumes; a phenotype seen in patients with systolic heart failure (HF). Here, we describe the creation of a rat model of pressure overload induced moderate remodeling and early systolic dysfunction (MOD) by ascending aortic banding (AAB) via a vascular clip with an internal area of 2 mm2. The surgery is performed in 200 g Sprague-Dawley rats. The MOD HF phenotype develops at 8-12 weeks after AAB and is characterized noninvasively by means of echocardiography. Previous work suggests the activation of signal transduction pathways and altered gene expression and post-translational modification of proteins in the MOD HF phenotype that mimic those seen in human systolic HF; therefore, making the MOD HF phenotype a suitable model for translational research to identify and test potential therapeutic anti-remodeling targets in HF. The advantages of the MOD HF phenotype compared to the overt systolic HF phenotype is that it allows for the identification of molecular targets involved in the early remodeling process and the early application of therapeutic interventions. The limitation of the MOD HF phenotype is that it may not mimic the spectrum of diseases leading to systolic HF in human. Moreover, it is a challenging phenotype to create, as the AAB surgery is associated with high mortality and failure rates with only 20% of operated rats developing the desired HF phenotype.

We describe the creation of a rat model of pressure overload induced moderate remodeling and early systolic dysfunction where signal transduction pathways involved in the initiation of the remodeling process are activated. This animal model will aid in identifying molecular targets for applying early therapeutic anti-remodeling strategies for heart failure.

In response to an injury, such as myocardial infarction, prolonged hypertension or a cardiotoxic agent, the heart initially adapts through the activation ...

An Image Guided Transapical Mitral Valve Leaflet Puncture Model of Controlled Volume Overload from Mitral Regurgitation in the Rat

Mitral regurgitation (MR) is a widely prevalent heart valve lesion, which causes cardiac remodeling and leads to congestive heart failure. Though the risks of uncorrected MR and its poor prognosis are known, the longitudinal changes in cardiac function, structure and remodeling are incompletely understood. This knowledge gap has limited our understanding of the optimal timing for MR correction, and the benefit that early versus late MR correction may have on the left ventricle. To investigate the molecular mechanisms that underlie left ventricular remodeling in the setting of MR, animal models are necessary. Traditionally, the aorto-caval fistula model has been used to induce volume overload, which differs from clinically relevant lesions such as MR. MR represents a low-pressure volume overload hemodynamic stressor, which requires animal models that mimic this condition. Herein, we describe a rodent model of severe MR in which the anterior leaflet of the rat mitral valve is perforated with a 23G needle, in a beating heart, with echocardiographic image guidance. The severity of MR is assessed and confirmed with echocardiography, and the reproducibility of the model is reported.&#160;&#160;

A rodent model of left heart volume overload from mitral regurgitation is reported. Mitral regurgitation of controlled severity is induced by advancing a needle of defined dimensions into the anterior leaflet of the mitral valve, in a beating heart, with ultrasound guidance.&#160;

Mitral regurgitation (MR) is a widely prevalent heart valve lesion, which causes cardiac remodeling and leads to congestive heart failure. Though the ...

Using an Isolated Working Rat Heart Model to Test Donor Heart Preservation Strategies

Summary

Explore More Videos

Chapters in this video

Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

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

An Isolated Working Heart System for Large Animal Models

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

Murine Isolated Heart Model of Myocardial Stunning Associated with Cardioplegic Arrest

Vein Interposition Model: A Suitable Model to Study Bypass Graft Patency

The Inverted Heart Model for Interstitial Transudate Collection from the Isolated Rat Heart

A Chronic Cardiac Ischemia Model in Swine Using an Ameroid Constrictor

A Rat Model of Pressure Overload Induced Moderate Remodeling and Systolic Dysfunction as Opposed to Overt Systolic Heart Failure

An Image Guided Transapical Mitral Valve Leaflet Puncture Model of Controlled Volume Overload from Mitral Regurgitation in the Rat