This work was supported by generous funding to S.N.T. from the US National Institutes of Health (K99/R00 HL1431149; R01HL157803) and American Heart Association (18CDA34110049). We also gratefully acknowledge funding from the US National Institute of Health (R01DK134590; R24OD034189), National Science Foundation (EEC 1941543), Harvard Medical School Eleanor and Miles Shore Fellowship, Polsky Family Foundation, the Claflin Distinguished Scholar Award on behalf of the MGH Executive Committee on Research, and Shriners Children&#8217;s Boston (Grant #BOS-85115).

This study is conducted following the Institutional Animal Care and Use Committee (IACUC) of Massachusetts General Hospital.
1. System design
<ol>
	<li>Assemble the system with the three double-jacketed components, including a bubble trap, a reservoir, an oxygenator, a peristaltic pump, and a water circulator.</li>
	<li>Attach all jacketed components to one clamp stand. Connect the components in sequence with silicone tubing in two different sequences (Figure 1A).
	<ol>
		<li>Sequence 1 - Flow pattern of water through the jacket (solid lines in Figure 1A):
		<ol>
			<li>Connect the outflow of the water circulator to the bottom inlet of the bubble trap jacket using 36 G tubing. This will guarantee that the perfusate is kept at the proper temperature (37 &#176;C) before reaching the heart, as the water will lose heat as it travels through the other components of the system.</li>
			<li>Connect the top inlet of the bubble trap jacket to the bottom inlet of the reservoir jacket using the same size tubing.</li>
			<li>Subsequently, connect the top inlet of the reservoir jacket to the bottom inlet of the oxygenator jacket.</li>
			<li>Finally, connect the oxygenator&#39;s top inlet to the water circulator&#39;s inflow.</li>
		</ol>
		</li>
		<li>Sequence 2 - Flow pattern of the perfusate through the system (dashed lines in Figure 1A)
		<ol>
			<li>Attach the luer connectors to both sides of 16 G tubing. Attach the first end to the base of the reservoir and feed it through the peristaltic pump head. Connect the other end to one of the inlets of the silicone coil within the oxygenator.</li>
			<li>Connect a second piece of 16 G tubing, fitted with luer lock connectors on both ends, at the second inlet of the silicone coil of the oxygenator to the inlet in the bubble trap with the long protrusion.</li>
			<li>Connect a shorter piece of 16 G tubing, fitted with luer connectors, to the unused outlet of the bubble trap to a three-way luer lock valve.</li>
			<li>On the opposite side of the three-way valve, connect a piece of 16 G tubing with a second luer valve on the other end.&#160;This second valve lies immediately above the reservoir. Connect the opposite side of the valve to more 16 G tubing, followed by the pressure sensor.</li>
			<li>Connect a smaller diameter tube (&#732;3.7 mm) to the vertical port of the three-way valve with a connector to the cannula (14 G angiocath). The perfusate flows from the reservoir to the oxygenator through the bubble trap before recirculating back into the reservoir through the aortic cannula connection.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Perfusate preparation
<ol>
	<li>Prepare base perfusate, 0.96% Krebs-Henseleit Buffer, 9.915 mM Dextran, 25 mM Sodium Bicarbonate, 1.054 mM Bovine Serum Albumin, 1% Pen Strep, 0.13% Insulin, 0.02% Hydrocortisone, 0.5% Heparin, and 2.75 mM Calcium Chloride and bring to volume with distilled water.</li>
</ol>
3. Perfusion system setup
<ol>
	<li>Connect two 10 mL syringes to the top and side vent ports on the bubble trap.</li>
	<li>Add the base perfusate (75 mL) to the reservoir. Turn on the peristaltic pump and set the water circulator to 37 &#176;C.</li>
	<li>Connect an oxygen (95% O2 and 5% CO2) line to the third inlet in the oxygenator and oxygenate the perfusate to a minimum pO2 of 400 mmHg.</li>
	<li>Secure an injection port to the vertical port of the three-way valve immediately past the bubble trap. Connect a winged infusion needle with a 1 mL syringe to the injection port (used for sampling). Gently tap the injection port or draw perfusate with the 1 mL syringe to clear any bubbles introduced to the circuit.</li>
	<li>Once the base perfusate is up to temperature and oxygen level, perform an initial reading of the biochemical parameters to ensure correct ion concentration (Table 1) and proper oxygenation. 
	NOTE: The ion and pH levels must be read after the solution has been brought up to temperature (37 &#176;C ) and has been oxygenated with the proper gas mix (95% O2, 5% CO2).</li>
	<li>Zero the pressure sensor by unclamping the connected tube and allow the perfusate flow through the open sensor and the cannula to equilibrate. Once equilibrated, press the zero button in the sensor box and re-clamp the tubing. Record base pressures prior to the heart&#39;s attachment to the system for flows ranging from 1 mL/min to 15 mL/min.</li>
	<li>Low pressure perfusion
	<ol>
		<li>Adenosine drip: Make an initial 20 mM stock of adenosine in the base perfusate. Dissolve the adenosine by placing the tube in a warm water bath and mixing by inversion.</li>
		<li>Dilute the stock adenosine further to a concertation of 0.06 mg/mL in base perfusate and add it to a 50 mL syringe.</li>
		<li>Attach a winged infusion needle to the 50 mL syringe and connect it to the injection port in the three-way valve. Secure the syringe to a syringe pump and set it to an infusion speed of 166.6 &#181;L/min. 
		NOTE: Bubbles are released from the infusion port by lightly tapping or flicking the port.</li>
	</ol>
	</li>
	<li>High pressure perfusion:
	<ol>
		<li>Packed red blood cells (pRBCs) isolation:
		<ol>
			<li>Collect 10-12 mL of whole rat blood via cardiac puncture of a donor rat.</li>
			<li>Centrifuge the blood at 2000 x g for 10 min.</li>
			<li>Remove the plasma and buffy coat layer via pipetting.</li>
			<li>Resuspend the pRBCs in perfusate without calcium chloride at a 1:1 ratio (e.g., 5 mL of pRBCs: 5 mL of perfusate) by inversion mixing.</li>
			<li>Repeat steps 3.8.1.2-3.8.1.4 twice for a total of 3 washes.</li>
			<li>After the last wash, resuspend the cells in perfusate at a 1:1 ratio and add the mixture to the perfusion system, already containing the 75 mL of base perfusate.</li>
		</ol>
		</li>
		<li>Allow the cells to distribute evenly through the system, and measure the hematocrit of the perfusate using a hematology machine. The hematocrit ranges from 5%-7%.</li>
	</ol>
	</li>
</ol>
4. Cardiac graft procurement preparation
<ol>
	<li>Completely prime the perfusion system before initiation of procurement to minimize cold ischemia time.</li>
	<li>Prepare the surgical tools. Surgical tools include blue pads, surgical tape, silk 5-0 sutures, cotton swabs, saline syringes (50 mL and 10 mL), operating scissors, forceps, micro scissors, micro forceps, Halstead clamp, 30 U of heparin, 16 G tubing for portal flush, 14 G tubing for canulation of heart, 16 G angiocath, modified 14 G angiocath with cuff, pressure sensor, ice bucket with ice, 47 mm Petri dish, gauze.</li>
	<li>Create a modified cannula by inserting a thin ring of tubing (inner diameter [ID] 0.167 mm, outer diameter [OD] 2.42 mm) on the 14 G cannula, creating a cuff effect.
	<ol>
		<li>Remove the needle of the cannula and add a drop of super glue below the ring. Carefully slide the ring to 1/4 inches above the base of the cannula. Allow glue to dry before use.</li>
		<li>Cut the cannula as close as possible to the cuff at an angle and remove sharp edges.</li>
	</ol>
	</li>
	<li>Fill a 60 mL syringe with heparinized saline (0.03 U/mL) for portal vein flush. Connect the syringe to the pressure sensor, followed by the 16 G flushing tube.</li>
	<li>Connect a 10 mL syringe of heparinized saline (0.03 U/mL) to the 14 G tube. Connect the other end of the tube to the aortic cannula and flush to remove any air bubbles.</li>
	<li>Place the aortic cannula in a 47 mm Petri dish with gauze and filled with saline. Leave the petri dish on ice until the heart is connected to the perfusion system.</li>
</ol>
5. Cardiac graft procurement
<ol>
	<li>Anesthetize the rats in an anesthetic chamber with 3% isoflurane.</li>
	<li>Once reflexes are not noticed, remove the rat from the chamber, place it into the surgical space, and deliver continuous isoflurane (3%) via face mask.</li>
	<li>Perform a toe pinch test, then heparinize the animal through the penile vein with 30 U of heparin.</li>
	<li>Shave the rat across the entire abdomen and upper chest area. Remove the fur shavings from the surgical field. Tape each limb to ensure no movement during surgery.</li>
	<li>Perform a toe pinch test once again to confirm the absence of reflexes. Then, make a horizontal midline incision in the skin of the lower abdomen, exposing the abdominal muscles.</li>
	<li>Make a second horizontal midline incision in the abdominal muscles exposing the internal organs.</li>
	<li>Reveal the sternum, secure it with a hemostat, and retract it cranially to expose the liver and portal vein. Cannulate the portal vein using a 16-gauge angiocath.</li>
	<li>Attach the 60 mL syringe of heparinized saline to the angiocath and create an incision in the inferior vena cava and abdominal aorta for venting. Flush the full amount of saline through the portal vein. 
	NOTE:&#160;The flushing pressure should remain around 10 mmHg.</li>
	<li>Make a horizontal cut in the diaphragm, followed by a proximal cut through the ribs on both sides of the sternum to reveal the thoracic cavity.</li>
	<li>Remove the heart from the cavity and immediately place it on the Petri dish with saline on ice.</li>
	<li>Identify the aortic arch, clamp with hemostats, and expose the descending aorta by cleaning any remaining connective tissue.</li>
	<li>Make a horizontal cut halfway across the descending aorta and cannulate with the 14 G angiocath. 
	NOTE: Do not breach the aortic valve with the cannula.</li>
	<li>Secure the cannula via a suture above the cuff and release the hemostat.</li>
	<li>Let the heart remain on ice until placed in the perfusion system.</li>
</ol>
6. Perfusion initiation
<ol>
	<li>Set the flow of the peristaltic pump to 1.0 mL/min. 
	NOTE: The aortic cannula is always handled at a 90&#730; angle with respect to the heart to avoid the introduction of bubbles into the coronaries (Figure 1B).</li>
	<li>Weigh the heart with the cannula before attaching the heart to the system. 
	NOTE: The aortic cannula should be completely free of any air bubbles.</li>
	<li>Attach the cannula to the connector in the system and start a timer.</li>
	<li>Once the heart has a full contraction, increase the flow by 0.2 mL/min increments while closely watching the pressures.</li>
	<li>Stop increases in flow when the desired pressures are reached or until a minimum of 3.5 mL/min is reached.
	<ol>
		<li>Low pressure perfusion
		<ol>
			<li>For pressures between 30-35 mmHg, use a flow of 4.5 mL/min.</li>
			<li>Start the adenosine syringe pump.</li>
		</ol>
		</li>
		<li>High pressure perfusion
		<ol>
			<li>For pressures between 70-80 mmHg, use a minimum flow of 5.0 mL/min.</li>
			<li>Start the adenosine syringe pump.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
7. Intraventricular balloon:
<ol>
	<li>Attach a small (50 &#181;L) latex balloon to a balloon catheter (2 mm diameter, 15 cm long) with a tapered tip (1.4 mm diameter).</li>
	<li>Connect the catheter to a pressure sensor via a luer lock connector, and secure the entire setup to a clamp stand.</li>
	<li>Fill the balloon/catheter/pressure sensor with approximately 200 &#181;L of saline via a syringe attached to the top end of the pressure sensor, and remove bubbles from inside the sensor, catheter, and balloon.</li>
	<li>Calibrate the pressure sensor using a sphygmometer.</li>
	<li>Make a small horizontal incision above the left atrium. Deflate the balloon by drawing the syringe at the top of the pressure sensor and inserting it into the left ventricle.</li>
	<li>Initiate data acquisition and inflate the balloon until diastolic pressures read 0 mmHg.</li>
</ol>
8. Sampling
<ol>
	<li>Collect heart rate, aortic flow, and coronary pressures after the first 20 min of perfusion and every hour thereafter.</li>
</ol>
9. End/cleanup
<ol>
	<li>At the end of the perfusion, remove the heart from the system and weigh it for edema estimation.</li>
	<li>Cut the apex of the heart via a circumferential cut and flash freeze in liquid nitrogen for post-perfusion analyses.</li>
	<li>Cut a circumferential piece of the heart for histological imaging and staining. Dispose of the rest of the heart and canula.</li>
	<li>Rinse all the components of the system by adding copious quantities of deionized (DI) water into the reservoir and running the peristaltic pump. Collect the water in an external bucket.</li>
	<li>Repeat step 9.4 two to three times.</li>
	<li>Rinse all sample ports and pressure sensor tubing thoroughly.</li>
	<li>Fill the reservoir with 600 mL of DI water and 3 mL of laboratory-grade detergent pump throughout the system.</li>
	<li>Turn off the water heater, oxygen tank, and peristaltic pump.</li>
</ol>

Enhancing CVD Research with Modified Langendorff Perfusion

<ol>
	<li>Cardiovascular Diseases (cvds). World Health Organization Available from: <a target="_blank" href="https://www.who.int/news-room/fact-sheets/detail/cardiovascular-diseases-(cvds)">https://www.who.int/news-room/fact-sheets/detail/cardiovascular-diseases-(cvds)</a> (2021)</li><li>Amini, M., Zayeri, F., Salehi, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Trend+analysis+of+cardiovascular+disease+mortality,+incidence,+and+mortality-to-incidence+ratio:+Results+from+global+burden+of+disease+study+2017.">Trend analysis of cardiovascular disease mortality, incidence, and mortality-to-incidence ratio: Results from global burden of disease study 2017.</a> BMC Public Health. 21 (1), 401 (2021).</li><li>Aune, S. E., Yeh, S. T., Zelinski, D. P., Angelos, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+hydrogen+peroxide+and+oxidant+stress+in+a+recirculating+whole+blood-perfused+rat+heart+model.">Measurement of hydrogen peroxide and oxidant stress in a recirculating whole blood-perfused rat heart model.</a> Resuscitation. 82 (2), 222-227 (2011).</li><li>Lateef, R., Al-Masri, A., Alyahya, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Langendorff's+isolated+perfused+rat+heart+technique:+A+review.">Langendorff's isolated perfused rat heart technique: A review.</a> Int J Basic Clin Pharmacol. 4 (6), 1314-1322 (2015).</li><li>Herr, D. J., Aune, S. E., Menick, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+and+assessment+of+ischemia-reperfusion+injury+in+Langendorff-perfused+rat+hearts.">Induction and assessment of ischemia-reperfusion injury in Langendorff-perfused rat hearts.</a> J Vis Exp. (101), e52908 (2015).</li><li>Vervoorn, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extended+normothermic+ex+situ+heart+perfusion+without+functional+decline.">Extended normothermic ex situ heart perfusion without functional decline.</a> J Heart Lung Transplant. 43 (4), S156 (2024).</li><li>Moeslund, N., 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=Ex-situ+oxygenated+hypothermic+machine+perfusion+in+donation+after+circulatory+death+heart+transplantation+following+either+direct+procurement+or+in-situ+normothermic+regional+perfusion.">Ex-situ oxygenated hypothermic machine perfusion in donation after circulatory death heart transplantation following either direct procurement or in-situ normothermic regional perfusion.</a> J Heart Lung Transplant. 42 (6), 730-740 (2023).</li><li>Testai, L., Martelli, A., Cristofaro, M., Breschi, M. C., Calderone, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardioprotective+effects+of+different+flavonoids+against+myocardial+ischaemia/reperfusion+injury+in+Langendorff-perfused+rat+hearts.">Cardioprotective effects of different flavonoids against myocardial ischaemia/reperfusion injury in Langendorff-perfused rat hearts.</a> J Pharm Pharmacol. 65 (5), 750-756 (2013).</li><li>Watanabe, M., Okada, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Langendorff+perfusion+method+as+an+ex+vivo+model+to+evaluate+heart+function+in+rats.">Langendorff perfusion method as an ex vivo model to evaluate heart function in rats.</a> Methods Mol Biol. 1816, 107-116 (2018).</li><li>Chang, X., 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=Cardioprotective+effects+of+salidroside+on+myocardial+ischemia-reperfusion+injury+in+coronary+artery+occlusion-induced+rats+and+Langendorff-perfused+rat+hearts.">Cardioprotective effects of salidroside on myocardial ischemia-reperfusion injury in coronary artery occlusion-induced rats and Langendorff-perfused rat hearts.</a> Int J Cardiol. 215, 532-544 (2016).</li><li>Neely, J. R., Liebermeister, H., Battersby, E. J., Morgan, H. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+pressure+development+on+oxygen+consumption+by+isolated+rat+heart.">Effect of pressure development on oxygen consumption by isolated rat heart.</a> Am J Physiol. 212 (4), 804-814 (1967).</li><li>Matsuura, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Positive+inotropic+effects+of+atp+released+via+the+maxi-anion+channel+in+Langendorff-perfused+mouse+hearts+subjected+to+ischemia-reperfusion.">Positive inotropic effects of atp released via the maxi-anion channel in Langendorff-perfused mouse hearts subjected to ischemia-reperfusion.</a> Front Cell Dev Biol. 9, 597997 (2021).</li><li>Louradour, 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=Simultaneous+assessment+of+mechanical+and+electrical+function+in+Langendorff-perfused+ex-vivo+mouse+hearts.">Simultaneous assessment of mechanical and electrical function in Langendorff-perfused ex-vivo mouse hearts.</a> Front Cardiovasc Med. 10, 1293032 (2023).</li><li>Ueoka, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Testosterone+does+not+shorten+action+potential+duration+in+Langendorff-perfused+rabbit+ventricles.">Testosterone does not shorten action potential duration in Langendorff-perfused rabbit ventricles.</a> Heart Rhythm. 19 (11), 1864-1871 (2022).</li><li>Reichelt, M. E., Willems, L., Hack, B. A., Peart, J. N., Headrick, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+and+coronary+function+in+the+Langendorff-perfused+mouse+heart+model.">Cardiac and coronary function in the Langendorff-perfused mouse heart model.</a> Exp Physiol. 94 (1), 54-70 (2009).</li><li>Abel, R. M., Reis, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+coronary+blood+flow+and+perfusion+pressure+on+left+ventricular+contractility+in+dogs.">Effects of coronary blood flow and perfusion pressure on left ventricular contractility in dogs.</a> Circ Res. 27 (6), 961-971 (1970).</li><li>Arnold, G., Morgenstern, C., Lochner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+autoregulation+of+the+heart+work+by+the+coronary+perfusion+pressure.">The autoregulation of the heart work by the coronary perfusion pressure.</a> Pflugers Arch. 321 (1), 34-55 (1970).</li><li>Iwamoto, T., Bai, X. J., Downey, H. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coronary+perfusion+related+changes+in+myocardial+contractile+force+and+systolic+ventricular+stiffness.">Coronary perfusion related changes in myocardial contractile force and systolic ventricular stiffness.</a> Cardiovasc Res. 28 (9), 1331-1336 (1994).</li><li>Rossello, X., Hall, A. R., Bell, R. M., Yellon, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+the+Langendorff+perfused+isolated+mouse+heart+model+of+global+ischemia-reperfusion+injury:+Impact+of+ischemia+and+reperfusion+length+on+infarct+size+and+LDH+release.">Characterization of the Langendorff perfused isolated mouse heart model of global ischemia-reperfusion injury: Impact of ischemia and reperfusion length on infarct size and LDH release.</a> J Cardiovasc Pharmacol Ther. 21 (3), 286-295 (2016).</li><li>Headrick, J. P., Peart, J., Hack, B., Flood, A., Matherne, G. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+properties+and+responses+to+ischaemia-reperfusion+in+Langendorff+perfused+mouse+heart.">Functional properties and responses to ischaemia-reperfusion in Langendorff perfused mouse heart.</a> Exp Physiol. 86 (6), 703-716 (2001).</li><li>Noly, P. E., Naik, S., Tang, P., Lei, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+ex+vivo+murine+biventricular+function+in+a+Langendorff+model.">Assessment of ex vivo murine biventricular function in a Langendorff model.</a> J Vis Exp. (190), e64384 (2022).</li><li>Garcia-Manzano, A., Gonzalez-Llaven, J., Lemini, C., Rubio-Poo, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Standardization+of+rat+blood+clotting+tests+with+reagents+used+for+humans.">Standardization of rat blood clotting tests with reagents used for humans.</a> Proc West Pharmacol Soc. 44, 153-155 (2001).</li><li>Lewis, J. H., Van Thiel, D. H., Hasiba, U., Spero, J. A., Gavaler, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+hematology+and+coagulation:+Studies+on+rodentia+(rats).">Comparative hematology and coagulation: Studies on rodentia (rats).</a> Comp Biochem Physiol A Comp Physiol. 82 (1), 211-215 (1985).</li><li>Motayagheni, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modified+Langendorff+technique+for+mouse+heart+cannulation:+Improved+heart+quality+and+decreased+risk+of+ischemia.">Modified Langendorff technique for mouse heart cannulation: Improved heart quality and decreased risk of ischemia.</a> MethodsX. 4, 508-512 (2017).</li></ol>

SNT has patent applications relevant to this study and serves on the Scientific Advisory Board for Sylvatica Biotech Inc., a company focused on developing organ preservation technology. All competing interests are managed by the MGH and Partners HealthCare in accordance with their conflict-of-interest policies.

Langendorff perfusion is an extremely pliable technique that allows impressive tailoring and adjustment to meet a wide array of experimental needs. This tailoring is permitted by the significant adjustability of most perfusion parameters, including perfusion pressures. Due to the retrograde nature of Langendorff, perfusion pressures are equivalent to coronary perfusion pressures, which play an essential role in cardiac function. Coronary perfusion pressures (CPP) are known to directly control cardiac work, as a wide array of cardiac indices (i.e., left ventricular pressure, contractility (dP/dtmax), wall tension, ventricular stiffness) are directly proportional to CPP16,17,18. Historically, the field has utilized perfusion pressures, and in effect CPP, between 60 mmHg and 80 mmHg in an attempt to mimic physiological conditions5,8,15,19,20,21. However, the non-physiological nature of retrograde ex vivo machine perfusion, in combination with the high demand for work, leads to an over-time loss of cardiac function (Figure 3). Alternatively, lower perfusion pressures (30-35 mmHg), despite not accurately replicating the physiological conditions of rat hearts in vivo, inherently decrease cardiac work demand and achieve extended perfusion times (4 h) without the over-time loss of function (Figure 3), and decreased graft edema (Figure 2C). The use of lower perfusion pressures, although it signifies a deviation from physiological CPP, seems to provide important advantages over the use of physiological perfusion pressures, as the elimination of existent technique-dependent loss of function during Langendorff perfusion improves the technique into a more accurate and predictable model system with significant potential to advance cardiovascular research. Particularly, the research areas that benefit and/or require extended perfusion times to reach scientific relevance (i.e., drug treatments, immunological response analysis, gene editing, normothermic graft preservation, etc.) are becoming increasingly important in the battle against CVDs.
Langendorff perfusion is undisputable an essential tool for the field of cardiovascular research. Therefore, along with the significant benefits this scientific technique poses to the research community, it comes with an important level of scientific complexity. In effect, there are several critical steps within this protocol that require careful standardization, primarily to avoid cardiac graft damage prior to, during, and immediately after initiating perfusion. The first chance of graft damage is inconspicuous during the portal vein flush. This flush with heparinized saline aims to remove as much whole blood as possible from the cardiac graft with a double purpose. First, it serves as a way of euthanasia via exsanguination. Second, it minimizes the chances of coagulation within the cardiac graft during retrieval, cannulation, and transportation, as rat whole blood is known to have extremely short clothing times22,23. However, after hundreds of successful cardiac perfusions, it became apparent that the pressure applied to the rat organism during flushing is of utter importance, with the ideal flush pressure being around 10 mmHg. Higher portal vein flush pressures seem to result in damage to the vasculature of the cardiac graft, leading to increased vascular resistance (<img alt="Equation 1" src="/files/ftp_upload/66815/66815eq01.jpg" style="margin: auto;" />). Higher vascular resistance in effect results in target perfusion pressures being reached at lower flow rates. This imbalance between pressure and coronary flow is conveyed in the produced left ventricular pulse pressure (LVPP), resulting in significant variability.
The next instance of possible cardiac graft damage is during the connection of the graft to the system via the introduction of air bubbles into the coronaries. Air bubbles can be easily introduced by mishandling the cannulated heart (Figure 1B) or improper bubble removal from the perfusion system upstream of the bubble trap24. Due to the retrograde nature of this setup, any introduction of air will result in cardiac air embolism, leading to ischemic insults, fibrillation, and, very commonly, graft death. Finally, the last critical step to ensure protocol success occurs during the initiation of perfusion. Differently from the large majority of manuscripts that report utilizing Langendorff as a technique, the initiation of perfusion in this protocol is performed at relatively low flows (1 mL/min) with incremental increases (+0.2 mL/min), which warrant complete control over perfusion pressures5,8,15,19,20,21. This incremental increase in flow, and therefore pressure, is critical as abrupt changes in pressure irreversibly increase vascular resistance and alter the delicate flow/pressure balance.
High vascular resistance in pressure-controlled Langendorff perfusions is very consequential, as target perfusion pressures are reached at lower flows, and grafts result under-perfused. The large reliance on this perfect balance between flow and pressure is likely the largest limitation of this protocol, as any prior graft damage, intentional (i.e., extended cold preservation, warm ischemia insult, myocardial infarction, etc.) or unintentional, leads to increased vascular resistance. In effect, this protocol is particularly useful for research where the experiment starts after the initiation of perfusion (i.e., drug treatments, immunological response analysis, gene editing, normothermic graft preservation, etc.) but not prior. This limitation is a perfect example of one Langendorff not fitting all purposes and special care should be taken to tailor perfusion parameters to better meet experimental needs.

The field of cardiovascular research has seen important advancements in the diagnosis and treatment of cardiovascular diseases (CVDs). However, despite the general decrease in incidence and mortality rates, CVDs remain the leading cause of death globally1,2. This alarming fact highlights the need for increased research and scientific advancement, which is undoubtedly dependent on the accuracy and predictability of the available research tools. As a result, there is a constant need for innovation, improvement and/or repurposing of the research toolset. For instance, retrograde or Langendorff heart perfusion, a technique available to the field for over a century, can be easily modified to cover a larger scope of scientific needs and achieve a broader range of applications.
The isolation of the cardiac graft from the rest of the organism during Langendorff perfusion provides an important degree of control over a wide array of experimental parameters, including temperature, circulating solution, coronary perfusion pressures, etc.3,4,5,6,7. The manipulation of these parameters facilitates the simulation of a large number of cardiac scenarios that can be leveraged to further scientific advancements5,8,9,10. Among these parameters, perfusion pressure is likely the most overlooked experimental setting11.
During Langendorff, perfusion pressures exhibit a direct correlation with heart rate, peak systolic/diastolic pressures, and oxygen consumption11. This correlation provides direct and precise control over the amount of work produced by the cardiac grafts, which can be adjusted to meet individual experimental needs. Despite this valuable control capability, the field has historically gravitated towards the use of higher perfusion pressures (60-80 mmHg), subjecting all cardiac grafts to high work demand irrespective of experimental needs8,12,13,14,15. The consequences of this unnecessarily high demand for work arise from the overarching principle that overwork tends to result in premature failure. This seems to be particularly true for cardiac grafts perfused via Langendorff, as the non-physiological nature of this method and the lack of recovery support present in vivo seem to exacerbate graft failure. This premature loss of graft function limits perfusion and experimental times significantly. In effect, in circumstances where longer perfusion times are more relevant to the experimental needs (i.e., drug treatments, immunological response analysis, gene editing, graft preservation, etc.), lower cardiac work can be afforded in exchange for increased graft durability.
This protocol demonstrates the feasibility of utilizing lower perfusion pressures (30-35 mmHg) during Langendorff, as well as the significant effect these pose for cardiac graft function over time when compared to higher perfusion pressures (60-80 mmHg). Furthermore, the findings in this manuscript highlight the importance of prioritizing the customization of the wide array of perfusion parameters to better meet experimental needs.

<table><tbody><tr><td>5-0 Suture</td><td>Fine Scientific Tools</td><td>18020-50</td><td></td></tr><tr><td>14 G Angiocath</td><td>Becton Dickinson</td><td>381867</td><td></td></tr><tr><td>16 G Angiocath</td><td>Becton Dickinson</td><td>381957</td><td></td></tr><tr><td>24 mm Heart Chamber adaptors</td><td>Radnoti</td><td>140132</td><td></td></tr><tr><td>Balloon Catheter&amp;nbsp;</td><td>Radnoti</td><td>170423</td><td></td></tr><tr><td>BD&amp;nbsp;Slip Tip Sterile Syringes- 10 mL</td><td>Fisher Scientific</td><td>14-823-16E</td><td></td></tr><tr><td>BD&amp;nbsp;Slip Tip Sterile Syringes- 1 mL</td><td>Fisher Scientific</td><td>14-823-434</td><td></td></tr><tr><td>BD&amp;nbsp;Slip Tip Sterile Syringes- 50 mL</td><td>Fisher Scientific</td><td>14-820-11</td><td></td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma</td><td>A7906</td><td></td></tr><tr><td>Bubble Trap Compliance Chamber</td><td>Radnoti</td><td>130149</td><td></td></tr><tr><td>Calcium Chloride</td><td>Sigma</td><td>C7902</td><td></td></tr><tr><td>Clamp Holder</td><td>United Scientic</td><td>RTCLMP1</td><td></td></tr><tr><td>Dextran</td><td>Sigma</td><td>31389</td><td></td></tr><tr><td>DIN8 Extension Cable</td><td>Iworx</td><td>SKU&amp;nbsp;C-DIN-EXT</td><td></td></tr><tr><td>Falcon High Clarity 50 mL conical tubes</td><td>Fisher Scientific</td><td>14-432-22</td><td></td></tr><tr><td>GSC Go Science Crazy&amp;nbsp;Cast Iron Support Ring Stand</td><td>Fisher Scientific</td><td>S13748</td><td></td></tr><tr><td>Heart&amp;nbsp; Chamber</td><td>Radnoti</td><td>140160</td><td></td></tr><tr><td>Heated Water Circulator bath</td><td>Cole Parmer</td><td>N/A</td><td></td></tr><tr><td>Heparin sodium Injection</td><td>Medplus</td><td>G-0409-2720-0409-2721</td><td></td></tr><tr><td>Hydrocortisone</td><td>Solu-Cortef</td><td>MGH Pharmacy</td><td></td></tr><tr><td>Insulin</td><td>Humulin R</td><td>MGH Pharmacy</td><td></td></tr><tr><td>Insvasive Fluid Filled Blood Pressure Sensor</td><td>Iworx</td><td>SKU&amp;nbsp;BP-10x</td><td></td></tr><tr><td>Iworx Data Acquisition System</td><td>Iworx</td><td>IX-RA-834</td><td></td></tr><tr><td>Krebs-Henseleit Buffer</td><td>Sigma</td><td>K3753</td><td></td></tr><tr><td>Left Ventricular Pressure Balloon</td><td>Radnoti</td><td>170404</td><td></td></tr><tr><td>Masterflex L/S Easy-Load II Pump Head for Precision Tubing, PPS Housing, SS Rotor</td><td>VWR</td><td>MFLX77200-60</td><td></td></tr><tr><td>Masterflex L/S Standard Digital Pump Systems</td><td>VWR</td><td>MFLX07551-30</td><td></td></tr><tr><td>Membrane Oxygenating Chamber</td><td>Radnoti</td><td>130144</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>ThermoFisher Scientific</td><td>15140122</td><td></td></tr><tr><td>Polyethylene Tubing</td><td>Fisher Scientific</td><td>14-170-12H</td><td></td></tr><tr><td>Precision Pump Tubing-16</td><td>VWR</td><td>MFLX96410-16</td><td></td></tr><tr><td>Sodium Bicarobonate</td><td>Sigma</td><td>5761</td><td></td></tr><tr><td>Standard PHD ULTRA CP Syringe Pump</td><td>Harvard Aparatus</td><td>88-3015</td><td></td></tr><tr><td>Tygon Transfer Tubing</td><td>VWR</td><td>MFLX95702-03</td><td></td></tr></tbody></table>

modified langendorff perfusion for extended perfusion times of rodent cardiac grafts

Despite important advancements in the diagnosis and treatment of cardiovascular diseases (CVDs), the field is in urgent need of increased research and scientific advancement. As a result, innovation, improvement and/or repurposing of the available research toolset can provide improved testbeds for research advancement. Langendorff perfusion is an extremely valuable research technique for the field of CVD research that can be modified to accommodate a wide array of experimental needs. This tailoring can be achieved by personalizing a large number of perfusion parameters, including perfusion pressure, flow, perfusate, temperature, etc. This protocol demonstrates the versatility of Langendorff perfusion and the feasibility of achieving longer perfusion times (4 h) without graft function loss by utilizing lower perfusion pressures (30-35 mmHg). Achieving extended perfusion times without graft damage and/or function loss caused by the technique itself has the potential to eliminate confounding elements from experimental results. In effect, in scientific circumstances where longer perfusion times are relevant to the experimental needs (i.e., drug treatments, immunological response analysis, gene editing, graft preservation, etc.), lower perfusion pressures can be key for scientific success.

Hearts from adult male Lewis rats (250-300 g body weight) were harvested and perfused at high (70-80 mmHg) or low (30-35 mmHg) perfusion pressures (n = 3 per group). The effects of perfusion pressure on overall cardiac function and health were determined by collecting heart rate, edema, and left ventricular function.
A clear correlation between heart rate and perfusion pressures was determined (Figure 2). Heart rate was statistically higher in high-pressure hearts when compared to low-pressure hearts for all time points, except the first one (60 min, Figure 2A,B). Interestingly, low-pressure hearts seem to undergo a period of adjustment at the beginning of perfusion, where it took about 30 min for the heart rate to stabilize and reach the levels that were maintained through the rest of the perfusion (Figure 2A). A large difference in left ventricular pulse pressure (LVPP) was also observed between the groups, with the LVPP of high-pressure hearts being statistically higher than low-pressure hearts at every timepoint (Figure 3B). This sustained high demand for work resulted in a progressive loss of function in high-pressure hearts with a statistical decrease in LVPP seen after 2 h of perfusion (Figure 3A,B). Alternatively, no loss of function was present in hearts perfused with low pressures, with LVPP remaining unchanged throughout&#160;perfusion time (Figure 3A,B). Similar to LVPP, high-pressure hearts exhibited higher cardiac muscle contraction (dP/dtmax) and relaxation (dP/dtmin) throughout the perfusion time when compared to low-pressure hearts (Figure 3C,D). In accordance, high-pressure hearts underwent a progressive loss of contractility and relaxation capacity, with both parameters being statistically higher 1 h into the perfusion time when compared to the last hour of perfusion. Differently, cardiac muscle contractility and relaxation capabilities were comparably low in the low-pressure group and remained unchanged over 4 h of perfusion time (Figure 3C,D). In addition to the functional effects, high perfusion pressures over extended periods of time also exacerbate interstitial fluid retention within the cardiac grafts, leading to edema. This edema was semi-quantified in percent weight change and resulted in high-pressure hearts having statistically higher weight gain when compared to hearts perfused at low pressures (Figure 2C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66815/66815fig01.jpg" /> 
Figure 1: Perfusion system setup. (A) Overall perfusion setup. Dashed lines represent the order in which the components of the system were connected to optimize perfusate circulation. Solid, colored lines represent the order in which the components were connected to optimize perfusate temperature. (B) The proper way of handling the heart after cannulation to avoid emptying of the catheter and introducing air into the coronaries. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/66815/66815fig01large.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/66815/66815fig02.jpg" /> 
Figure 2: Effects of pressure on heart rate and edema. (A) Heart rate obtained from the intraventricular balloon measurements. The solid line is the median of the experimental groups. The shaded area is the interquartile range. (B) Area under the curve (AUC) of heart rate data for every hour of perfusion. (C) Percent weight gained after 4 h of perfusion at low and high pressures. All data are expressed as median &#177; interquartile range (IQR). *p &#60; 0.01, **p &#60; 0.05, ***p &#60; 0.001. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/66815/66815fig02large.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/66815/66815fig03.jpg" /> 
Figure 3: Effects of pressure on left ventricular function. (A) Maximum systolic pressure plotted over time, denoted as left ventricular pulse pressure (LVPP). The solid line is the median of the experimental groups. The shaded area is the interquartile range. (B) The area under the LVPP curve (AUC) for every hour of perfusion. (C) Cardiac muscle contractility quantified from the maximum derivative of the pressure pulse. (D) Cardiac muscle relaxation quantified from the minimum derivative of the pressure pulse. All data are expressed as median &#177; interquartile range. *p &#60; 0.01, **p &#60; 0.05, ***p &#60; 0.001, ****p &#60; 0.0001. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/66815/66815fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Ion&#160;</td>
			<td>Concentration (mmol/L)&#160;</td>
		</tr>
		<tr>
			<td>Na+&#160;</td>
			<td>135&#8211;145&#160;</td>
		</tr>
		<tr>
			<td>K +&#160;</td>
			<td>&#60;6.00&#160;</td>
		</tr>
		<tr>
			<td>Ca +2&#160;</td>
			<td>1.0&#8211;1.3&#160;</td>
		</tr>
		<tr>
			<td>Cl -&#160;</td>
			<td>96&#8211;106&#160;</td>
		</tr>
	</tbody>
</table>
Table 1: Acceptable range of ion concentration in the perfusate.

Author Spotlight: Advancing Cardiovascular Research &#8212; Tailored Langendorff Perfusion Techniques for Improved Experimental Outcomes

This article demonstrates the feasibility of achieving longer perfusion times (4 h) of murine cardiac grafts without function loss by employing lower (30-35 mmHg) than physiological (60-80 mmHg) perfusion pressures during Langendorff.

Modified Langendorff Perfusion for Extended Perfusion Times of Rodent Cardiac Grafts

Despite important advancements in the diagnosis and treatment of cardiovascular diseases (CVDs), the field is in urgent need of increased research and ...

modified-langendorff-perfusion-for-extended-perfusion-times-rodent

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

1.1K Views.  Harvard Medical School and Massachusetts General Hospital. This article demonstrates the feasibility of achieving longer perfusion times (4 h) of murine cardiac grafts without function loss by employing lower (30-35 mmHg) than physiological (60-80 mmHg) perfusion pressures during Langendorff. 

Video: Author Spotlight: Advancing Cardiovascular Research — Tailored Langendorff Perfusion Techniques for Improved Experimental Outcomes

Optical Mapping of Langendorff-perfused Rat Hearts

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in experimental models that range in scale from cell cultures to whole-organs[1, 2]. Using state-of-the-art optical imaging systems, generation and propagation of action potentials during normal cardiac rhythm or throughout initiation and maintenance of arrhythmias can be visualized almost instantly[1]. The latest commercially-available systems can provide information at exceedingly high spatiotemporal resolutions and were based on custom-built equipment initially developed to overcome the obstacles imposed by more conventional electrophysiological methods[1]. Advancements in high-resolution and high-speed complementary metal-oxide-semiconductor (CMOS) cameras and intensely-bright, light-emitting diodes (LEDs) as well as voltage-sensitive dyes, optics, and filters have begun to make electrical signal acquisition practical for cardiovascular cell biologists who are more accustomed to working with microscopes. Although the newest generation of CMOS cameras can acquire 10,000 frames per second on a 16,384 pixel array, depending on the type of sample preparation, long-established fluorescence acquisition technologies such as photodiode arrays, laser scanning systems, and cooled charged-coupled device (CCD) cameras still have some distinct advantages with respect to dynamic range, signal-to-noise ratio, and quantum efficiency[1, 3]. In the present study, Lewis rat hearts were perfused ex vivo with a crystalloid perfusate (Krebs-Henseleit solution) at 37&deg;C on a modified Langendorff apparatus. After a 20 minute stabilization period, the hearts were intermittently perfused with 11 mMol/L 2,3-butanedione monoxime to eliminate contraction-associated motion during image acquisition. For optical mapping, we loaded hearts with the fast-response potentiometric probe di-8-ANEPPS[4] (5 &mu;Mol/L) and briefly illuminated the preparation with 475&plusmn;15 nm excitation light. During a typical 2 second period of illumination, &gt;605 nm light emitted from the cardiac preparation was imaged with a high-speed CMOS camera connected to a horizontal macroscope. For this demonstration, hearts were paced at 300 beats per minute with a coaxial electrode connected to an isolated electrical stimulation unit. Simultaneous bipolar electrographic recordings were acquired and analyzed along with the voltage signals using readily-available software. In this manner, action potentials on the surface of Langendorff-perfused rat hearts can be visualized and registered with electrographic signals.

This article describes a high temporal and spatial resolution technique to optically image action potential movement on the surface of Langendorff-perfused rat hearts using a potentiometric dye (di-8-ANEPPS).

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in ...

Biology

Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts

The success of liver transplantation has resulted in a dramatic organ shortage. In most transplant regions 20-30% of patients on the waiting list for liver transplantation die without receiving an organ transplant or are delisted for disease progression. One strategy to increase the donor pool is the utilization of marginal grafts, such as fatty livers, grafts from older donors, or donation after cardiac death (DCD). The current preservation technique of cold static storage is only poorly tolerated by marginal livers resulting in significant organ damage. In addition, cold static organ storage does not allow graft assessment or repair prior to transplantation.
These shortcomings of cold static preservation have triggered an interest in warm perfused organ preservation to reduce cold ischemic injury, assess liver grafts during preservation, and explore the opportunity to repair marginal livers prior to transplantation. The optimal pressure and flow conditions, perfusion temperature, composition of the perfusion solution and the need for an oxygen carrier has been controversial in the past.
In spite of promising results in several animal studies, the complexity and the costs have prevented a broader clinical application so far. Recently, with enhanced technology and a better understanding of liver physiology during ex vivo perfusion the outcome of warm liver perfusion has improved and consistently good results can be achieved.
This paper will provide information about liver retrieval, storage techniques, and isolated liver perfusion in pigs. We will illustrate a) the requirements to ensure sufficient oxygen supply to the organ, b) technical considerations about the perfusion machine and the perfusion solution, and c) biochemical aspects of isolated organs.

Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts

Marginal grafts, such as fatty livers, grafts from older donors, or livers retrieved after cardiac death (DCD) tolerate conventional, cold static storage only poorly. We developed a novel model of subnormothermic ex vivo liver perfusion for preservation, assessment, and repair of marginal liver grafts prior to transplantation.

The success of liver transplantation has resulted in a dramatic organ shortage. In most transplant regions 20-30% of patients on the waiting list for ...

Medicine

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing modified cannulas and techniques adapted from conventional commercial ex vivo perfusion equipment. The system was utilized to evaluate the preservation outcomes following 12 h of perfusion. C57BL/6J mice served as liver donors, and the livers were explanted by cannulating the portal vein (PV) and bile duct (BD), and subsequently flushing the organ with warm (37 &#176;C) heparinized saline. Then, the explanted livers were transferred to the perfusion chamber and subjected to normothermic oxygenated machine perfusion (NEVLP). Inlet and outlet perfusate samples were collected at 3 h intervals for perfusate analysis. Upon completion of the perfusion, liver samples were obtained for histological analysis, with morphological integrity assessed using modified Suzuki-Score through Hematoxylin-Eosin (HE) staining. The optimization experiments yielded the following findings: (1) mice weighing over 30 g were deemed more suitable for the experiment due to the larger size of their bile duct (BD). (2) a 2 Fr (outer diameter = 0.66 mm) polyurethane cannula was better suited for cannulating the portal vein (PV) when compared to a polypropylene cannula. This was attributed to the polyurethane material&#39;s enhanced grip, resulting in reduced catheter slippage during the transfer from the body to the organ chamber. (3) for cannulation of the bile duct (BD), a 1 Fr (outer diameter = 0.33 mm) polyurethane cannula was found to be more effective compared to the polypropylene UT - 03 (outer diameter = 0.30 mm) cannula. With this optimized protocol, mouse livers were successfully preserved for a duration of 12 h without significant impact on the histological structure. Hematoxylin-Eosin (HE) staining revealed a well-preserved morphological architecture of the liver, characterized by predominantly viable hepatocytes with clearly visible nuclei and mild dilation of hepatic sinusoids.

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

A normothermic ex vivo liver perfusion (NEVLP) system was created for mouse livers. This system requires experience in microsurgery but allows for reproducible perfusion results. The ability to utilize mouse livers facilitates the investigation of molecular pathways to identify novel perfusate additives and enables the execution of experiments focused on organ repair.

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing ...

Engineering

Induction and Assessment of Ischemia-reperfusion Injury in Langendorff-perfused Rat Hearts

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for myocardial infarction and cardiac complications of thoracic surgery. The ability of certain drugs to precondition the myocardium against ischemia reperfusion injury has led to multiple clinical trials, with little success. The isolated heart model allows acute observation of the functional effects of ischemia reperfusion injury in real time, including the effects of various pharmacological interventions administered at any time-point before or within the ischemia-reperfusion injury window. Since brief periods of ischemia can precondition the heart against ischemic injury, in situ aortic cannulation is performed to allow for functional assessment of non-preconditioned myocardium. A saline filled balloon is placed into the left ventricle to allow for real-time measurement of pressure generation. Ischemic injury is simulated by the cessation of perfusion buffer flow, followed by reperfusion. The duration of both ischemia and reperfusion can be modulated to examine biochemical events at any given time-point. Although the Langendorff isolated heart model does not allow for the consideration of systemic events affecting ischemia and reperfusion, it is an excellent model for the examination of acute functional and biochemical events within the window of ischemia reperfusion injury as well as the effect of pharmacological intervention on cardiac pre- and postconditioning. The goal of this protocol is to demonstrate how to perform in situ aortic cannulation and heart excision followed by ischemia/reperfusion injury in the Langendorff model.

The isolated rat heart is an enduring model for ischemia reperfusion injury. Here, we describe the process of harvesting the beating heart from a rat via in situ aortic cannulation, Langendorff perfusion of the heart, simulated ischemia-reperfusion injury, and infarct staining to confirm the extent of ischemic insult.

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for ...

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 ...

Porcine Liver Transplantation Without Veno-Venous Bypass As an Extended Criteria Donor Model

Liver transplantation is regarded as the gold standard for the treatment of a variety of fatal hepatic diseases. However, unsolved issues of chronic graft failure, ongoing organ donor shortages, and the increased use of marginal grafts call for the improvement of current concepts, such as the implementation of organ machine perfusion. In order to evaluate new methods of graft reconditioning and modulation, translational models are required. With respect to anatomical and physiological similarities to humans and recent progress in the field of xenotransplantation, pigs have become the main large animal species used in transplantation models. After the initial introduction of a porcine orthotopic liver transplant model by Garnier et al. in 1965, several modifications have been published over the past 60 years.
Due to specifies-specific anatomical traits, a veno-venous bypass during the anhepatic phase is regarded as a necessity to reduce intestinal congestion and ischemia resulting in hemodynamic instability and perioperative mortality. However, the implementation of a bypass increases the technical and logistical complexity of the procedure. Furthermore, associated complications such as air embolism, hemorrhage, and the need for a simultaneous splenectomy have been reported previously.
In this protocol, we describe a model of porcine orthotopic liver transplantation without the use of a veno-venous bypass. The engraftment of donor livers after static cold storage of 20 h - simulating extended criteria donor conditions - demonstrates that this simplified approach can be performed without significant hemodynamic alterations or intraoperative mortality and with regular uptake of liver function (as defined by bile production and liver-specific CYP1A2 metabolism). The success of this approach is ensured by an optimized surgical technique and a sophisticated anesthesiologic volume and vasopressor management.
This model should be of special interest for workgroups focusing on the immediate postoperative course, ischemia-reperfusion injury, associated immunological mechanisms, and the reconditioning of extended criteria donor organs.

In this protocol, a model of porcine orthotopic liver transplantation after static cold storage of donor organs for 20 h without the use of a veno-venous bypass during engraftment is described. The approach uses a simplified surgical technique with minimization of the anhepatic phase and sophisticated volume and vasopressor management.

Liver transplantation is regarded as the gold standard for the treatment of a variety of fatal hepatic diseases. However, unsolved issues of chronic graft ...

Assessment of Ex Vivo Murine Biventricular Function in a Langendorff Model

Primary graft dysfunction (PGD) remains the leading cause of early death following heart transplantation. Prolonged ischemic time during cold preservation is an important risk factor for PGD, and reliable evaluation of cardiac function is essential to study the functional responses of the donor heart after cold preservation. The accompanying video describes a technique to assess murine right and left ventricular function using ex vivo perfusion based in a Langendorff model after cold preservation for different durations. In brief, the heart is isolated and stored in a cold histidine-tryptophan-ketoglutarate (HTK) solution. Then, the heart is perfused with a Kreb buffer in a Langendorff model for 60 min. A silicone balloon is inserted into the left and right ventricle, and cardiac functional parameters are recorded (dP/dt, pressure-volume relationships). This protocol allows the reliable evaluation of cardiac function after different heart preservation protocols. Importantly, this technique allows the study of cardiac preservation responses specifically in native cardiac cells. The use of very small murine hearts allows access to an enormous array of transgenic mice to investigate the mechanisms of PGD.

Assessment of Ex Vivo Murine Biventricular Function in a Langendorff Model

Presented here is a protocol to reliably quantify the right and left ventricular function of donor hearts after cold preservation using an ex vivo perfusion system.

Primary graft dysfunction (PGD) remains the leading cause of early death following heart transplantation. Prolonged ischemic time during cold preservation ...

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 ...

Enhanced Cardiac Assessment in Ex Vivo Perfusion with Left Ventricular Loading

Cardiac Loading using Passive Left Atrial Pressurization and Passive Afterload for Graft Assessment

Ex vivo machine perfusion or normothermic machine perfusion is a preservation method that has gained great importance in the transplantation field. Despite the immense opportunity for assessment due to the beating state of the heart, current clinical practice depends on limited metabolic trends for graft evaluation. Hemodynamic measurements obtained from left ventricular loading have garnered significant attention within the field due to their potential as objective assessment parameters. In effect, this protocol provides an easy and effective manner of incorporating loading capabilities to established Langendorff perfusion systems through the simple addition of an extra reservoir. Furthermore, it demonstrates the feasibility of employing passive left atrial pressurization for loading, an approach that, to our knowledge, has not been previously demonstrated. This approach is complemented by a passive Windkessel base afterload, which acts as a compliance chamber to maximize myocardial perfusion during diastole. Lastly, it highlights the capability of capturing functional metrics during cardiac loading, including left ventricular pulse pressure, contractility, and relaxation, to uncover deficiencies in cardiac graft function after extended periods of preservation times (&#707;6 h).

Author Spotlight: Enhancing Graft Viability Assessment Through Quantitative Metrics and Innovative Reservoir Systems

The protocol describes a porcine ex vivo heart perfusion system in which direct loading of the left ventricle may serve as an assessment technique for graft health while simultaneously providing a holistic evaluation of graft function. A discussion of the system design and possible assessment metrics is also provided.

Ex vivo machine perfusion or normothermic machine perfusion is a preservation method that has gained great importance in the transplantation field. ...

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, ...

Modified Langendorff Perfusion for Extended Perfusion Times of Rodent Cardiac Grafts

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Cardiac Loading using Passive Left Atrial Pressurization and Passive Afterload for Graft Assessment

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