Portions of this work were supported by a generous contribution by the Fondation Marcel et Rolande Gosselin and Fondation Mr Stefane Foumy. Nicolas Noiseux is scholar of the FRQ-S.
The authors wish to thanks Josh Zhuo Le Huang, Gabrielle Gascon, Sophia Ghiassi, and Catherine Scalabrini for their support in data collection.

All animal care and experimental protocols conformed to the Guide for the Care and Use of Laboratory Animals and were approved by the institutional animal care and use committee of the Centre Hospitalier de l&#8217;Universit&#233; de Montr&#233;al Research Center.
1. Preliminary Preparations
<ol>
	<li>Turn on the water bath to heat the cardioplegia delivery system (Figure 1A) and the Langendorff ex vivo perfusion system (Figure 1B). Set the water temperature to 38.5 &#176;C for a solution temperature of 37 &#176;C. Setup photographs can be seen in Supplementary Figure 1A,B.</li>
	<li>Prepare 1 L of cardioplegic solution. Add 1 mL of 2% lidocaine hydrochloride and 10 mL of 2 mM KCl (final concentration 20 mM) to 1 L of Plasma-Lyte A (140 mM Na, 5 mM K, 1.5 mM Mg, 98 mM Cl, 27 mM acetate, 23 mM gluconate). Correct pH to 7.4 using 6 N HCl. 
	CAUTION: This model is highly sensitive to pH. A wrong pH correction (outside the 7.3-7.4 physiological range) or pH unstable solutions may compromise the experiment or provide unreliable data.</li>
	<li>Prepare 4 L of Krebs solution (113 mM NaCl, 4.5 mM KCl, 1.6 mM NaH2PO4, 1.25 mM CaCl2, 1 mM MgCl2&#8729;6H2O, 5.5 mM D-Glucose, 25 mM NaHCO3). Substrate masses per 1 L of solution should be as follows: 6.1 g of NaCl, 0.3355 g of KCl, 0.2035 g of MgCl2&#8729;6H2O, 0.192 g of NaH2PO4, 0.1387 g of CaCl2, 0.99 g of D-Glucose, 2.1 g of NaHCO3, final volume of 1 L in ultrapure deionized water. Add the NaHCO3 last to avoid precipitation. Filter the solution using a 0.22 &#181;m filter and store overnight. Correct the pH to 7.4 when the solution is at 37 &#176;C and bubble with 5% CO2/95% O2.</li>
	<li>Fill the Langendorff circuit with Krebs solution and start the system pump. Make sure that no bubbles are left inside the tubing. Adjust the peristaltic pump speed to 80 rpm (equivalent to 1 L/min). Using the two way stop cock, adjust the flow to maintain a slow drip through the aortic cannula until the heart is attached (Figure 1B). Keep a sample of Krebs solution (15 mL) in a 50 mL conic tube on ice for heart transportation.</li>
	<li>Fill the cardioplegia delivery system with the cardioplegic solution. Once the bubbles are removed, switch the circuit to saline using a 3 way stop cock (Figure 1A). Adjust the drip rate. Saline must be slowly dripping from the tip of the catheter to assure that no cardioplegic solution is injected before the animal&#8217;s death.</li>
</ol>
2. Animal Preparation
<ol>
	<li>Using an inhalation chamber, induce anesthesia with 3% isoflurane. Once the animal is unresponsive, perform an intraperitoneal injection of ketamine (75 mg/kg) and xylazine (5 mg/kg) or similarly suitable anesthetic, following local regulations, to maintain anesthesia for the rest of the procedure. Ensure the depth of anesthesia by no reaction to toe pinch and palpebral reflex.</li>
	<li>Intubate the animal using a 14 G, 2-inch I.V. catheter. Start ventilation at 50 breaths per min, with airway pressure limited to 20 cmH2O.</li>
	<li>Place the animal on a heating pad set to &#8220;medium&#8221; and cover with an absorbent pad to maintain body temperature. Insert a rectal temperature probe and attach a transdermal pulse oximeter sensor to one of the feet. Maintain rectal temperature at 37 &#176;C throughout the procedure.</li>
	<li>Vascular access
	<ol>
		<li>Make a 3 to 4 cm midline skin incision in the neck using scissors. Using blunt tip curved scissors, blunt dissect the subcutaneous tissue and expose the right sternohyoid muscle. Using non traumatic forceps, move the muscle laterally until the right carotid artery (pulsating), jugular vein (non-pulsating) and the vagus nerve (white) are visually identified (Supplementary Figure 2A). Carefully separate the vagus nerve from the carotid artery using blunt tip curved scissors.</li>
		<li>Inject heparin (2,000 IU/kg) via the right jugular vein. Apply pressure to the injection site after needle retraction to avoid blood leakage.</li>
		<li>Using curved forceps, pass two 5-0 silk sutures around the carotid artery. Firmly attach a distal suture to occlude the carotid artery at the superior aspect of the exposed artery. Keep the proximal suture untied. Pulling of the proximal suture will be used for bleeding control in the next step (Supplementary Figure 2B). The distance between sutures should be approximately 2 cm.</li>
		<li>Using a stereomicroscope for better visualization, carefully make a 1 mm incision with microsurgery scissors over the anterior wall of the carotid artery. Insert a 22 G, 1-inch closed I.V. catheter towards the aortic arch. The catheter is connected to a 2 way stop cock, allowing for connection to a pressure transducer for constant monitoring, with the possibility of injecting saline or cardioplegia via the cardioplegia delivery system (Figure 1A).</li>
	</ol>
	</li>
</ol>
3. Initiation of Cardiac Donation After Circulatory Death (DCD) Protocol
NOTE: A complete protocol timeline can be seen in Figure 2.
<ol>
	<li>Re-asses the anesthetic depth by performing a toe pinch and evaluating palpebral reflex. If reaction is observed, perform an intraperitoneal injection of Ketamine (37.5 mg/Kg) and xylazine (2.5 mg/Kg). Re-evaluate after 5 minutes. If no response is observed continue the procedure. Tracheal clamp should only be performed in adequately anesthetized animals.</li>
	<li>Turn off the ventilator and extubate the animal. Using mosquito forceps, clamp the trachea. This moment is considered as the start of the agonal phase. Start counting the functional warm ischemic time (WIT) when the peak systolic blood pressure drops below 30 mmHg, or if asystole or ventricular fibrillation appears, whatever comes first (Figure 3). 
	NOTE: Damage extent should be proportional to WIT. Experiments are needed to optimize WIT time according to anesthetic used, animal strain, sex and weight chosen. In control animals, immediately after carotid vascular access is secured, cardioplegia is injected and the heart is procured as described in the next step (Figure 2). The start of perfusion with cardioplegia is considered as the end of WIT.</li>
	<li>At the end of WIT, perform a medial sternotomy. Keep the thorax open by using an Alm retractor. Using scissors, open the inferior vena cava and both atria to avoid myocardial distension or cardioplegia recirculation (Supplementary figure 3). Clamp the aorta above the diaphragm. Through the previously catheterized carotid artery, infuse the cardioplegic solution at a constant pressure of 60 mmHg for 5 min using the cardioplegia delivery system. Infusion pressure can be modified by altering the height of the water column.</li>
	<li>At the end of cardioplegic infusion, dissect the ascending proximal aorta from the pulmonary artery using curved forceps (Supplementary Figure 4A). Cut the aorta distal to the left subclavian artery. Ensure an aortic length of at least 0.5 cm for cannulation for the Langendorff apparatus.</li>
	<li>Holding the heart from the aorta, complete the cardiectomy by separating the heart from the pulmonary veins and other thoracic structures (Supplementary Figure 4B). Rapidly, submerge the heart in to ice-cold Krebs solution for rapid transportation to the ex vivo system. Keep the dissection and transport times as short as possible (5 min).</li>
</ol>
4. Ex Vivo Heart Perfusion System (EVHP) and Cardiac Functional Assessment
<ol>
	<li>Open the aortic lumen using forceps. Deair the aorta by filling the lumen with the dripping Krebs solution to avoid forcing bubbles in to the coronary vessels. Lower the cannula into the aorta, taking care not to pass the aortic root or damage the aortic valve leaflets. Fix the setup with a small clamp.</li>
	<li>Using the 2 way stopcock, increase the flow to search for possible leaks in the aorta. If none are detected, tightly fix the aorta to the cannula using a 2-0 silk suture. Fully open the flow to the cannula. Maintain aortic pressure at a physiological pressure of 60-70 mmHg (adjusted by changing the height of the system). At this moment the initial reperfusion and stabilization time is initiated. Aortic pressure can be modified according to the investigator&#8217;s experimental plan.</li>
	<li>Rotate the heart so the base of the heart (atria) is facing the pressure sensor. Widen the left ventricular atrial opening by dissecting the pulmonary veins. Insert the latex balloon connected to a pressure sensor. Make sure that the balloon is fully positioned inside the ventricle by visual inspection. Slowly fill the balloon with saline until end diastolic pressure (EDP) is set to 15 mmHg. Adjust as needed to keep EDP constant (pre-determined physiological EDP). The EDP can be adjusted according to the experimental objectives of each investigator.</li>
	<li>Insert the pacing electrode in the anterior face of the heart (right ventricular outflow tract). Avoid puncturing the coronary vessels. Once spontaneous beating is observed, initiate pacing at 300 beats per min. Required voltage may vary between experiments and rat strains.</li>
	<li>After 10 min of stabilization, initiate continuous intraventricular pressure measurement recording. This moment is considered the beginning of the reconditioning and assessment phase (time 0) that will last for 1 h (Figure 2). Reconditioning may be prolonged, but a time-dependent decrease in contractility is expected in all hearts.</li>
	<li>At the start of reconditioning, collect cardiac effluent dropping from the cardiac veins for 5 min for baseline coronary flow assessment and biochemical analyses. For troponin T repeat every 15 min (times 0, 15, 30, 45 and 60 min). For other analyses individualization of collection times is needed (Figure 2).</li>
</ol>
5. End of Experience
<ol>
	<li>Remove the heart from the Langendorff apparatus.</li>
	<li>Using a straight high carbon steel blade (microtome blade or similar), remove the base of the heart (including aorta and pulmonary artery).</li>
	<li>With the right ventricle facing down, cut transverse ventricular slides of 1-2 mm thickness. In one representative section (normally the third) excise the right ventricle and snap freeze the left ventricle. This sample can be used for biochemical analyses.</li>
	<li>Submerge the remaining sections in to freshly prepared 5% 2,3,5-triphenyl-tetrazolium chloride in commercial phosphate buffer saline pH 7.4 for 10 min at 37 &#176;C. Viable tissues are colored red brick.</li>
	<li>Wash twice with phosphate buffer saline pH 7.4 and fix with 10% formalin at 4 &#176;C overnight. Wash twice with phosphate buffered saline pH 7.4 and keep each slice submerged.</li>
	<li>Withdraw excess liquid and weight each slide. Take digital color images of both sides. Use planimetric analyses to calculate percent infarct size and correct for slice and total ventricular weight. Coloration fades with time. Photos must be taken as soon as possible.</li>
</ol>
6. Data analyses
<ol>
	<li>Save all pressure data in a new file per animal.</li>
	<li>For pressure analyses, select at least 200 pressure cycles per time points. Analyses can be performed off-line (after completion of the experiment) using dedicated software (i.e., LabChart). Common cardiovascular parameters available include: Maximal generated pressure, end diastolic pressure, +dP/dt (steepest slope during the upstroke of the pressure curve, an indicator of ventricular contractile ability), -dP/dt (steepest slope during the downstroke of the pressure curve, an indicator of ventricular relaxation capacity) among others. 
	NOTE: For troponin analyses, an increase in troponin release at reperfusion is expected. After 1 h of reperfusion in the EVHP system, troponin levels may decrease to baseline, stressing the need for careful timing in the collection and handling of these samples.</li>
</ol>

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The authors report no proprietary or commercial interest in any product mentioned or concept discussed in this article.

The protocol presented here introduces a simple, convenient and versatile model of cardiac DCD, offering the opportunity to assess cardiac functional recovery, tissue damage and the use of post-conditioning cardioprotective agents to improve recovery of donor hearts otherwise discarded for transplantation. Ex vivo heart perfusion systems (EVHP) systems have been optimized to provide a platform for evaluating cardiac function and offer a unique opportunity to deliver and test modified solutions supplemented with post-conditioning pharmacological agents to preserve and repair DCD hearts in small15 and large animals16,17 models of cardiac DCD. Nevertheless the protocols are often insufficiently detailed and not always clinically relevant, making clinical translation difficult.
In the realm of DCD models, ex vivo DCD models, like the one described by Sanz18, lack an agonal phase. By inducing cardiac arrest by stopping mechanical ventilation, the sympathetic nervous system is overactivated, leading to a &#8220;catecholamine storm&#8221;19. This increase in catecholamines modifies the characteristics of the donor organs, and has been linked to a reduced functional status of experimental DCD organs19. Additionally, the progressive decline in function prior to asystole leads to right ventricular distension and consequent injury. In our protocol, we have induced circulatory death using a clinically relevant asphyxiation model, which maintains these responses.
Two main in vivo cardiac DCD models are described in the literature: open chest15 and closed chest20 models. Cardiac physiology is altered by the open chest approach by reducing the mechanical lung/heart interaction and preload. Furthermore, in open chest procedures, body heat loss is accelerated, further affecting functional outcomes21. Therefore it is preferable to maintain a closed chest approach preventing heat loss. Another refinement is to minimize the variability of time to circulatory death. Kearns et al. reported that time to death (time to non-pulsatile or mean blood pressure less than 30 mmHg) was between 3 to 11 min. In the 10 and 20 min WIT, 40% and 60% of hearts did not recover function, respectively, on an ex vivo working heart apparatus, making data interpretation more difficult15. An alternative to reduce the time to circulatory death is to use paralytic agents20; nevertheless, some evidence points towards direct cardiac effects of vecuronium, due to its effects on sympathetic and parasympathetic innervation22. To increase reproducibility, we elected for tracheal clamping, combined with a precise arterial pressure monitorization, allowing for a more homogeneous agonal time (&#60;5 min). It is known that organ damage starts before the moment of circulatory death; with some authors considering a cut-off systolic blood pressure below 50 mmHg as the beginning of functional WIT6, explaining the reluctance to transplant organs following a long period form withdraw of life sustaining measures until reperfusion. In this protocol, the WIT definition used follows the current experimental standard15, nevertheless, further studies are needed to clarify the exact set of hemodynamic parameters that mark the induction of organ damage in order to improve WIT calculation, thus offering better information for clinical practice.
The infusion of cardioplegic solution at constant physiological pressure and temperature offers a unique opportunity to initiate heart conditioning and tissue protection with any pharmacological agent or by other means. Technical refinements include clamping the thoracic aorta, limiting perfusion to the heart and thereby reducing the amount of solution needed for each essay. Once the heart is on the EVHP system, standardized functional evaluation is necessary. It has been shown that the use of an EVHP system has the potential to improve resuscitation of hearts previously considered not transplantable23,24. Interestingly, the clinically available EVHP system evaluates cardiac viability only by using serial lactate measurements8,23. Lactate measurements are not related to cardiac performance of DCD hearts24,25, thus additional measurements to evaluate transplantability are necessary. This experimental setup allows for a complete functional evaluation, including generated pressures and myocardial contractility measurements including +dP/dT and &#8211;dP/dT, allowing for a more thorough evaluation of cardiac function before the final transplant decision is made. Additionally, measurements of cardiac troponin, a marker of myocardial damage directly correlated to ischemic infarct size26, and release kinetics are related to the extent of cardiac ischemia in a Langendorff ischemia/reperfusion system. In particular, with long ischemic times (60 min), troponin levels are maintained after 1 h reperfusion, while LDH and creatinine kinase significantly decrease, and being non related to the extent of cardiac damage27,28, thus the use of serial troponin measures ensure a complete evaluation of organ viability before transplant. A major confounding variable in cardiac functional evaluation is the heart rate. Spontaneous heart rate is inversely related to length of ischemia29, and heart rate directly correlates with +dP/dt in isolated rat hearts30 and in animal models31. Interestingly, in recently published work on rodent models of DCD hearts and EVHP conditioning, pacing was not used and cardiac rates were variable and recorded in their protocols15,18,20. To maintain physiological heart rate, pacing was used once the heart had recovered rhythmic contraction. The chosen 300 bpm frequency is similar to those of healthy, non-stressed rats32.
Limitations of this protocol include the use of volatile anesthetic for induction. These agents have been shown to confer ischemic preconditioning33. Nevertheless, the short time of inhaled anesthetic use had no observable effect in this protocol and progressive myocardial dysfunction was still noted with increasing WIT. The use of normothermic cardioplegia can be also viewed as a limitation. Using normothermic cardioplegia allows optimal translation from the in vitro conditions used for the development of pharmacological conditioning agents, since cells are usually maintained at 37 &#176;C. Nevertheless, in this setup cardioplegia temperature can be easily regulated according to the requirements of the investigator. On the other hand, the use of a Langendorff preparation versus a working heart preparation for reconditioning might also be seen as a limitation. The working heart preparation allows for continuous recording of a pressure/volume loop12,15, with controlled pre and afterload, allowing for complete functional evaluation. The main advantage of a Langendorff preparation is that it maintains a constant aortic and perfusion pressure, especially during initial reperfusion, when generated pressure is minimal. In addition, the evaluation setup is simpler for the Langendorff heart compared to a working heart preparation. Nevertheless, this setup can be converted into a working heart preparation if deem necessary. Alternatively, cardiac reanimation can be performed in situ using normothermic regional perfusion, with cardiac performance being measured directly by the use of a Millar catheter34, allowing comprehensive hemodynamic and myocardial functional evaluation before organ procurement. In humans, both in situ and ex vivo reconditioning strategies have been described6, thus the development of both models allows for experimental comparisons that may translate in to optimization of clinical practice. Finally, the small size and high heart rate of this animal model may be considered as a limitation due to the potential technical difficulties observed while performing these experiments, and the inevitable physiological differences between rat and human hearts. If the EVHP evaluation is already standardized, a researcher can be familiarized with this technique by performing as little as 3 experiments. On the other hand, the use of this small animal model allows for convenient screening at a reasonable cost, reserving larger and more costly animal models such as the porcine model, to therapies with high human translational potential.
In conclusion, the protocol described here takes into account the best practices emanating from several groups researching DCD hearts. This protocol grants full control of WIT, allowing for a comprehensive structural and functional evaluation of cardioprotective conditioning treatment strategies in rats. This protocol can be upscaled and transferred to large animal models, allowing to translate research findings to clinical reality and ultimately allowing development of novel therapies increasing the quality and availability of lifesaving organs much needed by patients.

Solid organ transplantation in general and cardiac transplantation, in particular, are on the rise worldwide1,2. The standard method of organ procurement is donation after brain death (DBD). Given the strict inclusion criteria of DBD, less than 40% of the offered hearts are accepted3, thereby limiting the offer in face of increasing demand and extending the organ waiting list. To address this issue, the use of organs donated after circulatory death (DCD) is considered a potential solution4.
In DCD donors, however, an agonal phase following withdrawal of care and a period of unprotected warm ischemia before resuscitation are inevitable5. The potential organ injury after circulatory death can lead to organ dysfunction, explaining the reluctance to routinely adopt DCD heart transplantations. It is reported that only 4 centers use DCD hearts clinically, with stringent criteria that includes very short warm ischemia times and young donors without chronic pathologies6,7. For ethical and legal reasons, limited or no cardioprotective interventions can be applied in donors prior to circulatory death5,8,9. Thus, any mitigation to alleviate the ischemia-reperfusion (IR) injury is limited to cardioprotective therapies initiated during early reperfusion with cardioplegic solutions, and do not allow for proper functional assessment. Ex vivo heart perfusion (EVHP) and reconditioning of the DCD heart using dedicated platforms has been proposed as an alternative solution and studied by various scholars10,11,12,13. EVHP offers a unique opportunity to deliver post-conditioning agents to DCD hearts to improve functional recovery. However, for efficient clinical translation, many technical and practical issues remain to be addressed, and this is further compounded by a lack of consensus on a range of perfusion and functional criteria to determine transplantability6,8.
Herein we report the development of a reproducible pre-clinical small animal DCD protocol combined with an ex vivo heart perfusion system that can be used to investigate organ post-conditioning initiated at the time of procurement, during initial reperfusion, and/or throughout EVHP.

<table><tbody><tr><td>0.9% Sodium Chloride. 1 L bag</td><td>Baxter</td><td></td><td>Electrolyte solution for flushing in the modified Langendorff system.</td></tr><tr><td>14 G 2&quot; I.V catheter</td><td>Jelco</td><td>4098</td><td>To act as endotracheal tube.</td></tr><tr><td>2,3,5-Triphenyltetrazolium chloride</td><td>Milipore-Sigma</td><td>T8877</td><td>Vital coloration</td></tr><tr><td>22 G 1&quot; I.V catheter</td><td>BD</td><td>383532</td><td>I.V catheter with extension tube that facilitates manipulation for carotid catheterization</td></tr><tr><td>Adson Dressing Fcp, 4 3/4&quot;, Serr</td><td>Skalar</td><td>50-3147</td><td>Additional forceps for tissue manipulation</td></tr><tr><td>Alm Self-retaining retractor 4x4 Teeth Blunt 2-3/4&quot;</td><td>Skalar</td><td>22-9027</td><td>Tissue retractor used to maintain the chest open.</td></tr><tr><td>Bridge amp</td><td>ADinstruments</td><td>FE221</td><td>Bridge amp for intracarotid blood pressure measurement</td></tr><tr><td>Calcium chloride</td><td>Milipore-Sigma</td><td>C1016</td><td>CaCl&lt;sub&gt;2&lt;/sub&gt; anhydrous, granular, &amp;le;7.0 mm, &amp;ge;93.0% Part of the Krebs solution</td></tr><tr><td>D-(+)-Glucose</td><td>Milipore-Sigma</td><td>G8270</td><td>D-Glucose &amp;ge;99.5% Part of the Krebs solution</td></tr><tr><td>DIN(8) to Disposable BP Transducer</td><td>ADinstruments</td><td>MLAC06</td><td>Adapter cable for link between bridge amp and pressure transducer</td></tr><tr><td>Disposable BP Transducer (stopcock)</td><td>ADinstruments</td><td>MLT0670</td><td>Pressure transducer for intracarotid blood pressure measurement</td></tr><tr><td>dPBS</td><td>Gibco</td><td>14190-144</td><td>Electrolyte solution without calcium or magnesium.</td></tr><tr><td>Eye Dressing Fcp, Str, Serr, 4&quot;</td><td>Skalar</td><td>66-2740</td><td>Additional forceps for tissue manipulation</td></tr><tr><td>Formalin solution, neutral buffered, 10%</td><td>Milipore-Sigma</td><td>HT501128</td><td>Fixative solution</td></tr><tr><td>Heating Pad</td><td>Sunbean</td><td>756-CN</td><td></td></tr><tr><td>Heparin sodium 1,000 UI/mL</td><td>Sandoz</td><td></td><td>For systemic anticoagulation</td></tr><tr><td>Hydrochloric Acid 36,5 to 38,0%</td><td>Fisher scientific</td><td>A144-500</td><td>Diluted 1:1 for pH correction</td></tr><tr><td>Ketamine</td><td>Bimeda</td><td></td><td>Anesthetic. 100 mg/mL</td></tr><tr><td>LabChart</td><td>ADinstruments</td><td></td><td>Control software for the Powerlab polygraph, allowing off-line analyses. Version 7, with blood pressure and PV loop modules enabled</td></tr><tr><td>Left ventricle pressure balloon</td><td>Radnoti</td><td>170404</td><td>In latex. Size 4.</td></tr><tr><td>Lidocaine HCl 2% solution</td><td>AstraZeneca</td><td></td><td>Antiarrhythmic for the cardioplegic solution</td></tr><tr><td>Magnesium Chloride ACS</td><td>ACP Chemicals</td><td>M-0460</td><td>MgCl&lt;sub&gt;2&lt;/sub&gt;+6H&lt;sub&gt;2&lt;/sub&gt;O &amp;ge;99.0% Part of the Krebs solution</td></tr><tr><td>Micro pressure sensor</td><td>Radnoti</td><td>159905</td><td>Micro pressure sensor and amplifier connected to the intraventricular balloon</td></tr><tr><td>Pacemaker</td><td>Biotronik</td><td>Reliaty</td><td>Set to generate a pulse each 200 ms for a heart rate of 300 bpm.</td></tr><tr><td>pH bench top meter</td><td>Fisher scientific</td><td>AE150</td><td></td></tr><tr><td>Physiological monitor</td><td>Kent Scientific</td><td>Physiosuite</td><td>For continuous monitoring of rodent temperature and saturation during the procedure</td></tr><tr><td>Plasma-Lyte A</td><td>Baxter</td><td></td><td>Electrolyte solution used as base to prepare cardioplegia</td></tr><tr><td>Potassium Chloride</td><td>Milipore-Sigma</td><td>P4504</td><td>KCl &amp;ge;99.0% Part of the Krebs solution</td></tr><tr><td>Potassium Chloride 2 meq/ml</td><td>Hospira</td><td></td><td>Part of the cardioplegic solution</td></tr><tr><td>PowerLab 8/30 Polygraph</td><td>ADinstruments</td><td></td><td>Electronic polygraph</td></tr><tr><td>Silk 2-0</td><td>Ethicon</td><td>A305H</td><td>Suture material for Langendorff apparatus</td></tr><tr><td>Silk 5-0</td><td>Ethicon</td><td>A302H</td><td>Suture material for carotid</td></tr><tr><td>Small animal anesthesia workstation</td><td>Hallowell EMC</td><td>000A2770</td><td>Small animal ventilator</td></tr><tr><td>Sodium bicarbonate</td><td>Milipore-Sigma</td><td>S5761</td><td>NaHCO&lt;sub&gt;3&lt;/sub&gt; &amp;ge;99.5% Part of the Krebs solution</td></tr><tr><td>Sodium Chloride</td><td>Milipore-Sigma</td><td>S7653</td><td>NaCl &amp;ge;99.5% Part of the Krebs solution</td></tr><tr><td>Sodium Hydroxide pellets</td><td>ACP chemicals</td><td>S3700</td><td>Diluted to 5 N (10 g in 50 mL) for pH correction</td></tr><tr><td>Sodium phosphate monobasic</td><td>Milipore-Sigma</td><td>S0751</td><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; &amp;ge;99.0% Part of the Krebs solution</td></tr><tr><td>Stevens Tenotomy Sciss, Str, Delicate, SH/SH, 4 1/2&quot;</td><td>Skalar</td><td>22-1240</td><td>Small scisors for atria and cava vein opening</td></tr><tr><td>Tissue slicer blades</td><td>Thomas scientific</td><td>6727C18</td><td>Straight carbon steel blades for tissue slicing at the end of the protocol</td></tr><tr><td>Tuberculin safety syringe with needle 25 G 5/8&quot;</td><td>CardinalHealth</td><td>8881511235</td><td>For heparin injection</td></tr><tr><td>Veterinary General Surgery Set</td><td>Skalar</td><td>98-1275</td><td>Surgery instruments including disection scisors and mosquito clamps</td></tr><tr><td>Veterinary Micro Set</td><td>Skalar</td><td>98-1311</td><td>Surgery instruments with microscisors used for carotid artery opening</td></tr><tr><td>Working Heart Rat/Guinea Pig/Rabbit system</td><td>Radnoti</td><td>120101BEZ</td><td>Modular working heart system modified for the needs of the protocol. Includes all the necesary tubbing, water jacketed reservoirs and valves, including 2 and 3 way stop cock</td></tr><tr><td>Xylazine</td><td>Bayer</td><td></td><td>Sedative. 20 mg/mL</td></tr></tbody></table>

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.

Following extubation, blood pressure rapidly drops in a predictable pattern (Figure 3). Expected time to death is less than 5 min.
Figure 4 shows an average pressure/time curve at the start of reconditioning following 0, 10 and 15 min of WIT. Contractile function will improve over time. The use of short periods of WIT will allow for contractility to return to normal, and morphological damage will not be detectable (Figure 5 and Figure 6).
Proof-of-concept use of a conditioning agent added with the cardioplegia and at the stabilization phase show that the damage generated by 15 min of WIT in this model are amenable to modulation by cardioprotective agents (Figure 4, Figure 5 and Figure 6).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59789/59676fig01.jpg" /> 
Figure 1: Required equipment schemas.&#160;Minimal requirements for a (A) cardioplegia delivery system and a (B) Langendorff ex vivo heart perfusion system. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676fig01large.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/59789/59676fig02.jpg" /> 
Figure 2: Protocol timeline.&#160;Timeline from the moment of extubation until the end of the protocol. In control animals, cardioplegia is initiated without DCD or warm ischemic time. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676fig02large.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/59789/59676fig03.jpg" /> 
Figure 3: Intracarotid blood pressure/time plot.&#160;Typical evolution of intracarotid blood pressure following extubation. Warm ischemia time stars when the peak systolic blood pressure drops below 30 mmHg, or if asystole or ventricular fibrillation appears, whatever comes first. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676fig03large.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/59789/59676fig04.jpg" /> 
Figure 4: Ex vivo average beat-to-beat ventricular pressure time curve.&#160;Image derived from analyses of data taken after 10 min stabilization and perfusion (time 0 in figure 2) with or without the use of an experimental pharmacological cardioprotective conditioning agent. Ischemic time refers to warm ischemic times (WIT). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676fig04large.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/59789/59676fig05.jpg" /> 
Figure 5: Ex vivo recovery and functional analyses.&#160;(A) Continuous ventricular pressure-time curve after 10 min stabilization and perfusion with or without the use of an experimental pharmacological cardioprotective conditioning agent. Arrows show artifacts due to manual modification of EDP. (B) Maximum (+dP/dt) and minimum (-dP/dt) rate of pressure change in the LV vs. time plot derived from (A) showing a time-dependant improvement in contractility without treatment (green line). Short WIT (red line) or treated (yellow) hearts show a pattern similar to the control group (blue line). Data points are the mean of at least 200 individual beats. Bars show the standard error of the mean of each data point. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59789/59676fig06.jpg" /> 
Figure 6: 2,3,5-Triphenyl-tetrazolium chloride coloration at the end of experiments.&#160;Infarct area observed following diverse warm ischemic times (WIT) and the use of a pharmacological cardioprotective conditioning agent. Brick red: viable tissue. Light yellow: non-viable tissue. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Setup photograph. (A) Photograph showing the setup for the cardioplegia delivery system. Numbered equipment corresponds to: cardioplegia container (1), bubble trap (2), pressure sensor and catheter (3), peristaltic pump (4), polygraph connected to the pressure sensor (5) and small animal ventilator (6). (B) Photograph showing the setup for the Langendorff ex vivo heart perfusion system. Numbered equipment corresponds to: Perfusate container (1), conditioning agent container (2) and heart chamber (3). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676suppfig01large.jpg" target="_blank">Please click here to download this figure.</a>
Supplementary Figure 2: Neck dissection. (A) Photography showing the exposed jugular vein (arrow) prior to heparin injection. (B) shows the dissected carotid artery (arrow) with the sutures placed for bleeding control. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676suppfig02large.jpg" target="_blank">Please click here to download this figure.</a>
Supplementary Figure 3: Opening of the atria to prevent recirculation. (A) Photography showing the opening of the left atrial appendage (1). On the background the aorta (2) is clamped above the diaphragm (3). (B) Shows the opening of the right atrial appendage (1). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676suppfig03large.jpg" target="_blank">Please click here to download this figure.</a>
Supplementary Figure 4: Heart procurement. (A) Photography showing the use of curved forceps to separate the aorta (arrow) and the pulmonary artery. (B) Photography showing cardiac dissection and procurement. The heart is hold by the aorta using forceps. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59789/59676suppfig04large.jpg" target="_blank">Please click here to download this figure.</a>

Watch this Scientific Journal Video about Pre-clinical Model of Cardiac Donation after Circulatory Death at JoVE.com

Pre-clinical Model of Cardiac Donation after Circulatory Death

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 ...

pre-clinical-model-of-cardiac-donation-after-circulatory-death

Nanyang Technological University

Research

JoVE Journal

Medicine

7.9K Views.  Centre de recherche du Centre Hospitalier de l'Université de Montréal (CRCHUM). 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.

Video: Pre-clinical Model of Cardiac Donation after Circulatory Death

Pre-clinical Evaluation of Tyrosine Kinase Inhibitors for Treatment of Acute Leukemia

Receptor tyrosine kinases have been implicated in the development and progression of many cancers, including both leukemia and solid tumors, and are attractive druggable therapeutic targets. Here we describe an efficient four-step strategy for pre-clinical evaluation of tyrosine kinase inhibitors (TKIs) in the treatment of acute leukemia. Initially, western blot analysis is used to confirm target inhibition in cultured leukemia cells. Functional activity is then evaluated using clonogenic assays in methylcellulose or soft agar cultures. Experimental compounds that demonstrate activity in cell culture assays are evaluated in vivo using NOD-SCID-gamma (NSG) mice transplanted orthotopically with human leukemia cell lines. Initial in vivo pharmacodynamic studies evaluate target inhibition in leukemic blasts isolated from the bone marrow. This approach is used to determine the dose and schedule of administration required for effective target inhibition. Subsequent studies evaluate the efficacy of the TKIs in vivo using luciferase expressing leukemia cells, thereby allowing for non-invasive bioluminescent monitoring of leukemia burden and assessment of therapeutic response using an in vivo bioluminescence imaging system. This strategy has been effective for evaluation of TKIs in vitro and in vivo and can be applied for identification of molecularly-targeted agents with therapeutic potential or for direct comparison and prioritization of multiple compounds.

Receptor tyrosine kinases are ectopically expressed in many cancers and have been identified as therapeutic targets in acute leukemia. This manuscript describes an efficient strategy for pre-clinical evaluation of tyrosine kinase inhibitors for the treatment of acute leukemia.

Receptor tyrosine kinases have been implicated in the development and progression of many cancers, including both leukemia and solid tumors, and are ...

A Recovery Cardiopulmonary Bypass Model Without Transfusion or Inotropic Agents in Rats

Cardiopulmonary bypass (CPB) is indispensable in cardiovascular surgery. Despite the dramatic refinement of CPB technique and devices, multi-organ complications related to prolonged CPB still compromise the outcome of cardiovascular surgeries, and may worsen postoperative morbidity and mortality. Animal models recapitulating the clinical usage of CPB enable the clarification of the pathophysiological processes that occur during CPB, and facilitate pre-clinical studies to develop strategies protecting against these complications. Rat CPB models are advantageous because of their greater cost-effectiveness, convenient experimental processes, abundant testing methods at the genetic or protein levels, and genetic consistency. They can be used for investigating the immune system activation and synthesis of proinflammatory cytokines, compliment activation, and production of oxygen free radicals. The rat models have been refined and have gradually taken the place of large-animal models. Here, we describe a simple CPB model without transfusion and/or inotropic agents in a rat. This recovery model allows the study of the long-term multiple organ sequelae of CPB.

Here, we present a protocol to describe a simple recovery cardiopulmonary bypass model without transfusion or inotropic agents in a rat. This model allows the study of the long-term multiple organ sequelae of cardiopulmonary bypass.

Cardiopulmonary bypass (CPB) is indispensable in cardiovascular surgery. Despite the dramatic refinement of CPB technique and devices, multi-organ ...

Immunology and Infection

Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain death (BD) still represents the major source for solid transplants. Unfortunately, the irreversible loss of brain function is known to induce multiple pathophysiological changes, including hemodynamic as well as hormonal modifications, finally leading to a systemic inflammatory response. Models that allow a systematic investigation of these effects in vivo are scarce. We present a murine model of BD induction, which could aid investigations into the devastating effects of BD on allograft quality. After implementing intra-arterial blood pressure measurement via the common carotid artery and reliable ventilation via a tracheostomy, BD is induced by steadily increasing intracranial pressure using a balloon catheter. Four hours after BD induction, organs may be harvested for analysis or for further transplantation procedures. Our strategy enables the comprehensive analysis of donor BD in a murine model, therefore allowing an in-depth understanding of BD-related effects in solid organ transplantation and potentially paving the way to optimized organ preconditioning.

We present a murine model of brain death induction in order to evaluate the influence of its pathophysiological effects on organs as well as on consecutive grafts in the context of solid organ transplantation.

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain ...

Neuroscience

A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation

Fifty-years following the first successful report, cardiac transplantation remains the gold-standard treatment for eligible patients with advanced heart failure. Multiple small-animal models of heart transplantation have been used to study the acute and long-term effects of novel therapies. However, few are tested and demonstrated success in clinical trials. It is of critical importance to evaluate new therapies in a clinically relevant large-animal model for efficient and reliable translation of basic studies&#39;&#160;findings. Here, we describe a pre-clinical large-animal (porcine) model of orthotopic heart transplantation that has been firmly established and previously used to investigate novel cardioprotective strategies. This procedure focuses on acute ischemia-reperfusion injury and is a reliable method to investigate novel interventions which have been tested and validated in smaller experimental models, such as the murine model. We demonstrate its usefulness in assessing cardiac performance during the early post-transplantation period and other potential possibilities enabled by the model.

Here, we describe a pre-clinical large-animal (porcine) model of orthotopic heart transplantation that has been firmly established and utilized to investigate novel cardioprotective strategies.

Fifty-years following the first successful report, cardiac transplantation remains the gold-standard treatment for eligible patients with advanced heart ...

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 ...

Lung Rapid Recovery Procurement Combined with Abdominal Normothermic Regional Perfusion in Controlled Donation after Circulatory Death

Controlled donation after circulatory death (cDCD) has contributed to increasing donor numbers all over the world. Experiences published in the last years confirm that the outcomes after lung transplantation from cDCD are similar to those from brain death donors; however, the utilization of lungs from asystole donors remains low. Several reasons may be involved: different legal frameworks among countries and centers with different premortem interventions, inadequate lung donor care before procurement, or even poor experience with cDCD procedures and protocols.
Initially, the rapid recovery technique was commonly employed for the procurement of thoracic and abdominal organs in cDCD, but, in the last decade, abdominal normothermic regional perfusion (ANRP) with extracorporeal membrane oxygenation devices has become a useful method to restore blood flow to abdominal organs, allowing their quality improvement and their functional assessment prior to transplantation. This makes the donation procedure more complex and generates doubts about injury to the grafts due to dual temperature.
The aim of this article is to describe a protocol based on a single center experience with Maastricht III donors combining lung cooling rapid recovery in the thorax and abdominal normothermic regional perfusion. Tips and tricks focused on premortem interventions and lung procurement procedure techniques are explained. This may help to minimize the reluctance among professionals to use this combined technique and encourage other donor centers to use it, despite the increased complexity of the procedure.

The protocol combines a lung cooling rapid recovery technique with abdominal normothermic regional perfusion for abdominal graft procurement in controlled asystole donors, which is a safe and useful method to expand the donor pool.

Controlled donation after circulatory death (cDCD) has contributed to increasing donor numbers all over the world. Experiences published in the last years ...

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 ...

Extracorporeal Cardiopulmonary Resuscitation in Animal Model After Cardiac Arrest

Pediatric Animal Model of Extracorporeal Cardiopulmonary Resuscitation After Prolonged Circulatory Arrest

Congenital heart disease (CHD) is the most prevalent congenital malformation, with about one million births impacted worldwide per year. Comprehensive investigation of this disease requires appropriate and validated animal models. Piglets are commonly used for translational research due to their analogous anatomy and physiology. This work aimed to describe and validate a neonatal piglet model of cardiopulmonary bypass (CPB) with circulatory and cardiac arrest (CA) as a tool for studying severe brain damage and other complications of cardiac surgery. In addition to including a list of materials, this work provides a roadmap for other investigators to plan and execute this protocol. After experienced practitioners performed several trials, the representative results of the model demonstrated a 92% success rate, with failures attributed to small piglet size and variant vessel anatomy. Furthermore, the model allowed practitioners to select from a wide variety of experimental conditions, including varying times in CA, temperature alterations, and pharmacologic interventions. In summary, this method uses materials readily available in most hospital settings, is reliable and reproducible, and can be widely employed to enhance translational research in children undergoing heart surgery.

Author Spotlight: Simulating Pediatric Cardiac Surgery Using a Neonatal Piglet Model

This protocol describes a neonatal porcine model of cardiopulmonary bypass (CPB), with circulatory and cardiac arrest as a tool for studying severe brain damage and other complications secondary to CPB.

Congenital heart disease (CHD) is the most prevalent congenital malformation, with about one million births impacted worldwide per year. Comprehensive ...

Study of Experimental Organ Donation Models for Lung Transplantation

Experimental models are important tools for understanding the etiological phenomena involved in various pathophysiological events. In this context, different animal models are used to study the elements triggering the pathophysiology of primary graft dysfunction after transplantation to evaluate potential treatments. Currently, we can divide experimental donation models into two large groups: donation after brain death and donation after circulatory arrest. In addition, the deleterious effects associated with hemorrhagic shock should be considered when considering animal models of organ donation. Here, we describe the establishment of three different lung donation models (post-brain death donation, post-circulatory death donation, and post-hemorrhagic shock donation) and compare the inflammatory processes and pathological disorders associated with these events. The objective is to provide the scientific community with reliable animal models of lung donation for studying the associated pathological mechanisms and searching for new therapeutic targets to optimize the number of viable grafts for transplantation.

The present study shows the establishment of three different lung donation models (post-brain death donation, post-circulatory death donation, and post-hemorrhagic shock donation). It compares the inflammatory processes and pathological disorders associated with these events.

Experimental models are important tools for understanding the etiological phenomena involved in various pathophysiological events. In this context, ...

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

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.

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.

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