Name: using an isolated working rat heart model to test donor heart preservation strategies
Uploaded: 2024-10-04T13:50:00.000+00:00
Duration: 6 min 56 s
Description: Optimization of donor heart preservation solutions has played a key role in reducing ischemia-reperfusion injury in donor hearts during organ retrieval, ...

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><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&amp;nbsp;</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&amp;nbsp;</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&amp;nbsp;</td><td>26764.26</td><td>To make Krebs Buffer</td></tr><tr><td>Potassium phosphate monobasic&amp;nbsp;</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&quot;)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 &amp;micro;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

402 Views.  Victor Chang Cardiac Research Institute. Ons onderzoek maakt gebruik van een geïsoleerd hartperfusiecircuit van ratten om suppletiestrategieën voor het behoud van donorharten te testen. We streven ernaar conserveringsstrategieën te identificeren die het functionele herstel van donorharten kunnen verbeteren na lange perioden van koude opslag of lange perioden van warme ischemie na donatie na overlijden van de bloedsomloop. Het geïsoleerde werk in het hartprotocol voor ratten werd in eerste instan...

Video: Auteur in de schijnwerpers: verbetering van het behoud van het donorhart door middel van geïsoleerde onderzoeken naar de perfusie van het rattenhart

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 ...

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. ...

An Immunological Model for Heterotopic Heart and Cardiac Muscle Cell Transplantation in Rats

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications have been reported since the first description in 1964. After 30 years of performing heterotopic heart transplantation in rats, we have developed a simplified surgical approach, which can be easily taught and performed without further surgical training or background.
After dissection of the ascending aorta and the pulmonary artery and ligation of superior and inferior caval and pulmonary veins, the donor heart is harvested and subsequently perfused with ice-cold saline solution supplemented with heparin. After clamping and incising the recipient abdominal vessels, the donor ascending aorta and pulmonary artery are anastomosed to the recipient abdominal aorta and inferior vena cava, respectively, using continuous running sutures.
Depending on different donor-recipient combinations, this model allows analyses of either acute or chronic rejection of allografts. The immunological significance of this model is further enhanced by a novel approach of in-ear injection of vital cardiac muscle cells and subsequent analysis of draining cervical lymphatic tissue.

We describe a model of heterotopic abdominal heart transplantation in rats, implying modifications of current strategies, which lead to a simplified surgical approach. Additionally, we describe a novel rejection model by in-ear injection of vital cardiac muscle cells, allowing further transplant immunological analyses in rats.

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications ...

Immunology and Infection

Pre-clinical Model of Cardiac Donation after Circulatory Death

Cardiac transplantation demand is on the rise; nevertheless, organ availability is limited due to a paucity of suitable donors. Organ donation after circulatory death (DCD) is a solution to address this limited availability, but due to a period of prolonged warm ischemia and the risk of tissue injury, its routine use in cardiac transplantation is seldom seen. In this manuscript we provide a detailed protocol closely mimicking current clinical practices in the context of DCD with continuous monitoring of heart function, allowing for the evaluation of novel cardioprotective strategies and interventions to decrease ischemia-reperfusion injury.
In this model, the DCD protocol is initiated in anesthetized Lewis rats by stopping ventilation to induce circulatory death. When systolic blood pressure drops below 30 mmHg, the warm ischemic time is initiated. After a pre-set warm ischemic period, hearts are flushed with a normothermic cardioplegic solution, procured, and mounted onto a Langendorff ex vivo heart perfusion system. Following 10 min of initial reperfusion and stabilization, cardiac reconditioning is continuously evaluated for 60 min using intraventricular pressure monitoring. A heart injury is assessed by measuring cardiac troponin T and the infarct size is quantified by histological staining. The warm ischemic time can be modulated and tailored to develop the desired amount of structural and functional damage. This simple protocol allows for the evaluation of different cardioprotective conditioning strategies introduced at the moment of cardioplegia, initial reperfusion and/or during ex vivo perfusion. Findings obtained from this protocol can be reproduced in large models, facilitating clinical translation.

This protocol shows a simple and flexible approach for the evaluation of new conditioning agents or strategies to increase the feasibility of cardiac donation after circulatory death.

Cardiac transplantation demand is on the rise; nevertheless, organ availability is limited due to a paucity of suitable donors. Organ donation after ...

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of transplant with the potential to deliver durable heart valves capable of somatic growth with no requirement for anticoagulation. However, the immunobiology of heart valve transplants remains unexplored, highlighting the need for animal models to study this new type of transplant. Previous rat models for heterotopic aortic valve transplantation into the abdominal aorta have been described, though they are technically challenging and costly. For addressing this challenge, a renal subcapsular transplant model was developed in rodents as a practical and more straightforward method for studying heart valve transplant immunobiology. In this model, a single aortic valve leaflet is harvested and inserted into the renal subcapsular space. The kidney is easily accessible, and the transplanted tissue is securely contained in a subcapsular space that is well vascularized and can accommodate a variety of tissue sizes. Furthermore, because a single rat can provide three donor aortic leaflets and a single kidney can provide multiple sites for transplanted tissue, fewer rats are required for a given study. Here, the transplantation technique is described, providing a significant step forward in studying the transplant immunology of heart valve transplantation.

This protocol describes a simple and efficient method for the transplantation of aortic valve leaflets under the renal capsule to allow for the study of alloreactivity of heart valves.

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of ...

Rodent Working Heart Model for the Study of Myocardial Performance and Oxygen Consumption

Isolated working heart models have been used to understand the effects of loading conditions, heart rate and medications on myocardial performance in ways that cannot be accomplished in vivo. For example, inotropic medications commonly also affect preload and afterload, precluding load-independent assessments of their myocardial effects in vivo. Additionally, this model allows for sampling of coronary sinus effluent without contamination from systemic venous return, permitting assessment of myocardial oxygen consumption. Further, the advent of miniaturized pressure-volume catheters has allowed for the precise quantification of markers of both systolic and diastolic performance. We describe a model in which the left ventricle can be studied while performing both volume and pressure work under controlled conditions. 
In this technique, the heart and lungs of a Sprague-Dawley rat (weight 300-500 g) are removed en bloc under general anesthesia. The aorta is dissected free and cannulated for retrograde perfusion with oxygenated Krebs buffer. The pulmonary arteries and veins are ligated and the lungs removed from the preparation. The left atrium is then incised and cannulated using a separate venous cannula, attached to a preload block. Once this is determined to be leak-free, the left heart is loaded and retrograde perfusion stopped, creating the working heart model. The pulmonary artery is incised and cannulated for collection of coronary effluent and determination of myocardial oxygen consumption. A pressure-volume catheter is placed into the left ventricle either retrograde or through apical puncture. If desired, atrial pacing wires can be placed for more precise control of heart rate. This model allows for precise control of preload (using a left atrial pressure block), afterload (using an afterload block), heart rate (using pacing wires) and oxygen tension (using oxygen mixtures within the perfusate).

Isolated working heart models can be used to measure the effect of loading conditions, heart rate, and medications on myocardial performance and oxygen consumption. We describe methods for preparation of a rodent left heart working model that permits study of systolic and diastolic performance and oxygen consumption under various conditions.

Isolated working heart models have been used to understand the effects of loading conditions, heart rate and medications on myocardial performance in ways ...

Biology

A Heterotopic Rat Heart Transplantation Model using Circulatory Death Donor Hearts

The objective of this protocol is to set up a rat heterotopic heart transplantation model with donation after circulatory death (DCD) donor hearts. There are two setups for this protocol: heart donor setup and recipient setup. In the heart donor setup, Sprague Dawley rats are anesthetized, endotracheally intubated, and ventilated. The right carotid artery is cannulated to deliver heparin and the paralytic agent vecuronium-bromide. The DCD process is initiated by terminating the ventilation. After 20 min, the heart is exposed and the aorta distal to the brachiocephalic branch is clamped. At 25 min from terminating the ventilator, ice-cold University of Wisconsin (UW) solution is perfused through the carotid catheter to flush the heart. The heart is procured by dividing the aorta, pulmonary artery, venae cavae, and pulmonary veins and stored in UW solution for implantation. In the recipient setup, the Lewis rat is anesthetized with isoflurane. Slow-release buprenorphine is administered subcutaneously to facilitate a smooth postoperative recovery. Through a midline abdominal incision, the infra-renal aorta and the inferior vena cava are isolated and clamped with an atraumatic vascular clamp. The donor heart aorta and pulmonary artery are sutured to the recipient abdominal aorta and vena cava, respectively, with a running 8-0 Prolene. The vascular clamp is removed to reperfuse the heart. The abdominal wall is closed and the rat is recovered. After a set interval (24 h to 2 weeks), the recipient rat is anesthetized, the transplanted heart is exposed, and a balloon-tip-catheter is inserted into the left ventricle via the apex to record developed pressure and dP/dt using a data acquisition system. The heart tissue is collected for histology, immunology, or molecular analysis. A successful DCD donor rat heart transplantation model will allow further studies on the cardioprotective approaches to improve heart transplantation outcomes from DCD donors.

Here, we describe the preparation and technical details of murine heterotopic heart transplantation utilizing a circulatory death donor heart.

The objective of this protocol is to set up a rat heterotopic heart transplantation model with donation after circulatory death (DCD) donor hearts. There ...

Large-Animal Model of Donation after Circulatory Death and Normothermic Regional Perfusion for Cardiac Assessment

The increase in demand for cardiac transplantation throughout the years has fueled interest in donation after circulatory death (DCD) to expand the organ donor pool. However, the DCD process is associated with the risk of cardiac tissue injury due to the inevitable period of warm ischemia. Normothermic regional perfusion (NRP) allows for an in situ organ assessment, allowing the procurement of hearts determined to be viable. Here, we describe a clinically relevant large-animal model of DCD followed by NRP. Circulatory death is established in anesthetized pigs by stopping mechanical ventilation. After a preset warm ischemia period, an extracorporeal membrane oxygenator (ECMO) is used for a NRP period lasting at least 30 min. During this reperfusion period, the model allows the collection of various myocardial biopsies and blood samples for initial cardiac evaluation. Once NRP is weaned, biochemical, hemodynamic, and echocardiographic assessments of cardiac function and metabolism can be performed before organ procurement. This protocol closely simulates the clinical scenario previously described for DCD and NRP in heart transplantation and has the potential to facilitate studies aimed at decreasing ischemia-reperfusion injury and enhance cardiac functional preservation and recovery.

The protocol describes a large-animal (porcine) model of donation after circulatory death, followed by thoracoabdominal normothermic regional perfusion that closely simulates the clinical scenario in heart transplantation, and has the potential to facilitate therapeutic studies and strategies.

The increase in demand for cardiac transplantation throughout the years has fueled interest in donation after circulatory death (DCD) to expand the organ ...

Simultaneous Isolation of High Quality Cardiomyocytes, Endothelial Cells, and Fibroblasts from an Adult Rat Heart

The rat is an important animal model used in cardiovascular research, and rat cardiac cells are used routinely for in vitro analysis of the molecular mechanisms of cardiovascular disease progression such as cardiac hypertrophy, fibrosis, and atherosclerosis. Although several attempts with variable success have been made to develop immortalized cell lines from the cardiovascular system to understand these cellular mechanisms, primary cells offer a more natural and close to in vivo environment for such studies. Therefore, different laboratories working on a particular cell type have developed protocols to isolate individual types of rat cardiac cells of interest. A protocol that allows the isolation of more than one cell type, however, is missing. Here an optimized protocol is described that allows the isolation of high-quality major cardiac cell types (cardiomyocytes, endothelial cells, and fibroblasts) from a single preparation and enables their use for cellular analyses. This permits the most efficient use of available resources, which may save time and reduce research costs.

Several protocols have been developed and described for the isolation of different cardiac cell types from a rat heart. Here, an optimized protocol is described that allows the isolation of high-quality major cardiac cell types (cardiomyocytes, endothelial cells, and fibroblasts) from a single preparation, reducing the experimental costs.

The rat is an important animal model used in cardiovascular research, and rat cardiac cells are used routinely for in vitro analysis of the molecular ...

Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation

Heart transplantation is the most effective therapy for end-stage heart failure. Despite the improvements in therapeutic approaches and interventions, the number of heart failure patients waiting for transplantation is still increasing. The normothermic ex situ preservation technique has been established as a comparable method to the conventional static cold storage technique. The main advantage of this technique is that donor hearts can be preserved for up to 12 h in a physiologic condition. Moreover, this technique allows resuscitation of the donor hearts after circulatory death and applies required pharmacologic interventions to improve donor function after implantation. Numerous animal models have been established to improve normothermic ex situ preservation techniques and eliminate preservation-related complications. Although large animal models are easy to handle compared to small animal models, it is costly and challenging. We present a rat model of normothermic ex situ donor heart preservation followed by heterotopic abdominal transplantation. This model is relatively cheap and can be accomplished by a single experimenter.

Here, we present an assessment protocol of a heterotopically implanted heart after normothermic ex situ preservation in the rat model.

Heart transplantation is the most effective therapy for end-stage heart failure. Despite the improvements in therapeutic approaches and interventions, the ...

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

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