The authors would like to thank Ph.D. Bettina Sommer Cervantes for her support in the writing of this manuscript, and Kitzia Elena Lara Safont for her support with the illustrations.

The isolated perfusion system in the rabbit model has been widely used in the Bronchial Hyperresponsiveness Laboratory at the Instituto Nacional de Enfermedades Respiratorias. The protocol includes New Zealand rabbits with an approximate weight of 2.5-3 kg. All animals were kept in standard vivarium conditions and ad libitum feeding in compliance with the official Mexican guidelines for laboratory animals (NOM 062-ZOO-1999) and under the Guide for the Care and Use of Laboratory Animals (8th edition, 2011). All the animal procedures and animal care methods presented in this protocol were previously approved by the Ethics Committee of the Instituto Nacional de Enfermedades Respiratorias.
NOTE: The preparation of the isolated lung perfusion system involves the deliberate death of an animal under anesthesia and via euthanasia.
1. Equipment and preparation of apparatus.
<ol>
	<li>Equipment arrangement:
	<ol>
		<li>Set up an operating table with size according to the weight of the rabbit.</li>
		<li>Mount the cover of the artificial thorax on the steel column with the glass chamber underneath and the ventilator with a roller pump by the sides.</li>
		<li>Ensure that the cover is easily inclined to have the tracheal cannula in line with the trachea to allow a faster connection.</li>
	</ol>
	</li>
	<li>Artificial thorax: 
	NOTE: It is an essential part of the system. It consists of a water-jacketed glass chamber sealed by a special cover. The cover works as the organ holder with the connections to cannulate the trachea and vessels embedded in it.
	<ol>
		<li>Set up a venturi jet operated by compressed air to generate the negative pressure inside the artificial thorax. 
		NOTE: The ventilation control module (VCM) allows separate adjustments of inspiratory and end-expiratory pressures as well as respiration rate and the ratio of inspiratory duration to total cycle duration.</li>
	</ol>
	</li>
	<li>Apparatus:
	<ol>
		<li>Ensure that a normally working apparatus consists of a main steel column mounted on a base plate holding the artificial thorax, with the pneumotachometer and weight transducer located above it and behind the preheating coil with a bubble trap.</li>
		<li>Connect one differential pressure transducer to the pneumotachometer and another to the chamber pressure. Set a different couple of pressure transducers behind the thorax to measure perfusion and venous pressures.</li>
		<li>Connect the changeover stock below the oxygenator with a level electrode and the ventilation system beside the apparatus.</li>
	</ol>
	</li>
</ol>
2. Surgical extraction of the cardiopulmonary block.
<ol>
	<li>Anesthesia:
	<ol>
		<li>Use a combination of a sedative (xylazine) and a barbiturate (pentobarbital). 
		NOTE: Different anesthetic cocktails can be used with no effect on experimental outcomes.</li>
		<li>First, sedate the healthy New Zealand rabbits with a single intramuscular injection of xylazine hydrochloride (3-5 mg/kg). Ensure that the rabbits remain calm and relaxed to allow further manipulation after a few minutes of the injection.</li>
		<li>Following sedation, use the marginal (lateral) ear veins as access to anesthetize the rabbits with an intravenous injection of pentobarbital sodium (28 mg/kg).</li>
	</ol>
	</li>
	<li>Monitoring:
	<ol>
		<li>To avoid insufficient anesthesia or excessive depression of cardiac and respiratory functions, monitor the following parameters. To assess the depth of anesthesia, perform a toe pinch test.</li>
		<li>Ensure that the mucous membrane is pink. Blue or gray shades indicate hypoxia.</li>
		<li>Ensure that the heart rate is between 120-135 beats/min, and that the body temperature does not drop below 36.5 &#176;C.</li>
	</ol>
	</li>
	<li>Animal placement:
	<ol>
		<li>Shave the rabbit&#39;s torso and place the animal on the operating table in supine position. Place the ventilation system near the table, behind the rabbit&#39;s head, to permit connecting the cannulae quickly after tracheotomy to avoid tissular damage.</li>
	</ol>
	</li>
	<li>Incision and tracheotomy:
	<ol>
		<li>Dissect the skin with a ventral median line incision of 3-5 cm from the diaphragm up to the neck.</li>
		<li>With the operating scissors, cut the anterior 2/3 of the trachea between two cartilage rings to insert the tracheal cannula through the tracheal fibrous membrane.</li>
		<li>Insert a 5 mm (outer diameter; OD) tracheal cannula through the tracheal fibrous membrane and use a 4-0 silk suture to fix it carefully.</li>
		<li>Place either forceps or tweezers underneath the trachea to ensure the cannula did not bend against the trachea.</li>
	</ol>
	</li>
	<li>Positive-pressure ventilation:
	<ol>
		<li>As long as the lungs remain outside the artificial thorax, use a small species respiration pump to ventilate a positive pressure in order to avoid lung collapse during the surgery.</li>
		<li>Initiate ventilation through the tracheal cannula connected to the respiration pump quickly after tracheotomy and before the thorax is opened.</li>
		<li>Set the tidal volume at 10 mL/kg. 
		NOTE: Depending on the experiment setup and artificial thorax model, provide positive-pressure ventilation by either the same ventilation pump used to provide negative-pressure or a different one, granting a quick re-cannulation.</li>
	</ol>
	</li>
	<li>Thoracotomy and exsanguination:
	<ol>
		<li>To access the thoracic cavity, use a scalpel or scissors to open the thorax wall and perform a medial sternotomy up to the upper third of the thorax.</li>
		<li>Hold the thorax halves open by two retractors. Several lung flaps usually surround the heart.</li>
		<li>Localize the superior and inferior vena cava and refer them with threads.</li>
		<li>Prior to the exsanguination of the animal, identify the right ventricle and inject 1000 UI/kg of heparin.</li>
		<li>Immediately after the injection, ligate the superior and inferior vena cava with the pre-looped thread and perform exsanguination.</li>
	</ol>
	</li>
	<li>Heart-lung harvest:
	<ol>
		<li>Harvest the cardiopulmonary block gently and quickly. Use direct digital dissection or spring scissors to separate the connective tissue so as to remove the lungs from the thorax.</li>
		<li>Dissect the vasculature in the area, as well as the esophagus.</li>
		<li>Cut through the manubrium sterni to extend the medial sternotomy towards the tracheal cannula, releasing the trachea on both sides from connecting tissue.</li>
		<li>Now, resect the trachea above the tracheal cannula. Gently pull up the cannula in a craniocaudal axis as the dorsal fixation of the trachea and lungs is resected.</li>
	</ol>
	</li>
	<li>Cannulation:
	<ol>
		<li>Lift the isolated lungs out of the thorax and carefully place them over a sterile gauze on a petri dish.</li>
		<li>To prevent atelectasis, ventilate the lungs using positive-pressure ventilation with positive end-expiratory pressure (PEEP) set at 2 cmH2O.</li>
		<li>Remove the ventricles by cutting them off the heart at the level of the atrioventricular groove.</li>
		<li>After opening the two ventricles, introduce the OD 4.6 mm pulmonary artery cannula for the rabbit with a basket through the pulmonary artery and introduce the OD 5.9 mm left atrium cannula for the rabbit with the basket through the mitral valve into the left atrium.</li>
		<li>Use a 4-0 silk suture in the pulmonary artery and left atrium to fix the cannulaes. Include the surrounding tissues in the ligatures of the pulmonary artery and left atrium to avoid the distension of these structures.</li>
		<li>Inject 250 mL of saline isotonic solution through the arterial cannulae to flush the remaining blood from the vascular bed.</li>
	</ol>
	</li>
</ol>
3. Perfusion technique.
<ol>
	<li>Setup:
	<ol>
		<li>Place the isolated lungs carefully into the lung chamber.</li>
		<li>Attach the trachea to the transductor on the cover of the chamber.</li>
		<li>Connect the cannulated vessels to the perfusion system.</li>
		<li>Close the chamber and secure it with the rotary lock. 
		NOTE: The recirculating perfusion circuit consists of an open venous reservoir, a peristaltic pump, a heat exchanger, and a bubble trap.</li>
		<li>At this point, attach the chamber lid and switch over a stopcock to switch from positive to negative pressure ventilation. To check the negative pressure ventilation of the lungs and airtight closure of the chamber, inspect the respiratory excursion of the lung and chamber pressure on the pressure gauge.</li>
		<li>Perfuse the lungs with 200 mL of artificial blood-free perfusate (a Krebs-Ringer bicarbonate buffer containing 2.5% of bovine albumin).</li>
		<li>Start the perfusate flow at 3 mL/min/kg, then slowly step up the flow over a 5-min period to 5 mL/min/kg. Reach a flow of 8 mL/min/kg over the next 5 min and then after another 5-min period reach a maximum flux of 10 mL/min/kg. Take care of avoiding air from getting into the circuit. 
		NOTE: Maintain the pH and the temperature of the perfusate within physiological ranges (pH 7.4-7.5; temperature, 37 &#176;C-38 &#176;C). To adjust the pH, add NaHCO3&#160;(1N) or increase the flow of carbon dioxide. Alternatively, use HCl (0.1N) to acidify.</li>
	</ol>
	</li>
	<li>Parameters:
	<ol>
		<li>Check whether the predetermined perfusion and ventilation parameters are set as required.</li>
		<li>Ventilate the lungs with humidified air at a frequency of 30 bpm, a tidal volume of 10 mL/kg, and an end-expiratory pressure (Pe) of 2 cmH2O. 
		NOTE: The pulmonary arterial pressure (0-20 mmHg) corresponds to the height of the liquid level in the oxygenator or reservoir in centimeters above the pulmonary trunk, while the pulmonary venous pressure corresponds to the height of the pressure equilibration chamber above the left atrium. Both values can be modified. Note that left atrium pressure is also 0-20 mmHg.</li>
	</ol>
	</li>
	<li>Achieving zone 3 conditions:
	<ol>
		<li>Use the two catheters connected to side ports of the cannulae secured in the pulmonary artery, left atrium, and pressure transducers to measure the arterial (Pa) and venous (Pv) pressures.</li>
		<li>Set the baseline pressures at the level of the lung hilum (Zero-reference).</li>
		<li>Conduct the experiments under zone 3 ventilation conditions. To achieve this, wait for 10-15 min to obtain an equilibrium characterized by an isogravimetric state.</li>
		<li>Ensure that the venous pressure is higher than the alveolar pressure (Palv) and the arterial pressure remains higher than both (Pa &#62; Pv &#62; Palv) for Zone 3 conditions to occur.</li>
		<li>Ensure that the lungs&#39; weight remains constant and arterial and left atrial pressures are stable to achieve zone 3 conditions to open up a maximum number of pulmonary vessels and maintain the microvascular bed content during the experiment. 
		NOTE: The measurement of Kfc as an indicator of pulmonary edema has no variation between a manual and an automatic perfusion system.</li>
	</ol>
	</li>
	<li>Electronic control and signal processing: Ensure that the respiratory flow, weight changes, microvascular pressure, tidal volume, vascular resistance, among others, are registered on a multiple central electronics unit that integrates signals coming from the transducers and displays them on the evaluation system.</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ex+vivo+lung+perfusion:+a+comprehensive+review+of+the+development+and+exploration+of+future+trends.">Ex vivo lung perfusion: a comprehensive review of the development and exploration of future trends.</a> Transplantation. 96 (6), 509-518 (2013).</li><li>Delaunois, A., Gustin, P., Ansay, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple+muscarinic+receptor+subtypes+mediating+pulmonary+oedema+in+the+rabbit.">Multiple muscarinic receptor subtypes mediating pulmonary oedema in the rabbit.</a> Pulmonary Pharmacology. 7 (3), 185-193 (1994).</li><li>Delaunois, A., Gustin, P., Vargas, M., Ansay, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protective+effect+of+various+antagonists+of+inflammatory+mediators+against+paraoxon-induced+pulmonary+edema+in+the+rabbit.">Protective effect of various antagonists of inflammatory mediators against paraoxon-induced pulmonary edema in the rabbit.</a> Toxicology and Applied Pharmacology. 132 (2), 343-345 (1995).</li><li>Barr, H. 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C., Torres-Villalobos, G., Aguilar-Blas, N., Fr&#237;as-Guill&#233;n, J., Guerra-Mora, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+study+of+capillary+filtration+coefficient+(Kfc)+determination+by+a+manual+and+automatic+perfusion+system.+Step+by+step+technique+review.">Comparative study of capillary filtration coefficient (Kfc) determination by a manual and automatic perfusion system. Step by step technique review.</a> Physiological Research. 68 (6), 901-908 (2019).</li><li>Pereira, M. 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The authors declare no conflicts of interest.

This work displays a general view of the isolated lung perfusion system, an essential technique in pulmonary physiology research. The isolated lung perfusion system offers a great degree of versatility in its uses and allows the evaluation of several parameters relevant in the testing of a wide range of hypotheses15. An isolated lung system is a tool with worldwide presence that, in the last decade, has further established its relevance for organ-specific evaluations and also expanded its usefulne...

Brodie and Dixon first described the ex-vivo lung perfusion system in 1903 1. Since then, it has become a gold standard tool for studying the physiology, pharmacology, toxicology, and biochemistry of the lungs2,3. The technique offers a consistent and reproducible way to evaluate the viability of lung transplants, and to determine the effect of inflammatory mediators such as histamine, arachidonic acid metabolites, and substance P, among others, as well as their interactions during pulmonary phenomena such as bronchoconstriction, atelectasis, and pulmonary edema. The isolated lung system has been a key technique in unveiling the important role of the lungs in the elimination of biogenic amines from general circulation4,5. Additionally, the system has been used to evaluate the biochemistry of pulmonary surfactant6. Over the last few decades, the ex-vivo lung perfusion system has become an ideal platform for lung transplantation research7. In 2001 a team lead by Stig Steen described the first clinical application of the ex-vivo lung perfusion system by using it to recondition the lungs of a 19-year-old donor, who was initially rejected by transplantation centers due to its injuries. The left lung was harvested and perfused for 65 min; afterward, it was successfully transplanted into a 70-year-old man with COPD8. Further research into lung reconditioning using the ex-vivo perfusion led to developing the Toronto technique for extended lung perfusion to assess and treat injured donor lungs9,10. Clinically, the ex-vivo lung perfusion system has shown to be a safe strategy to increase donor pools by treating and reconditioning sub-standard donor lungs, presenting no significant difference in risks or outcomes against standard criteria donors10. 
The main advantage of the isolated lung perfusion system is that the experimental parameters can be evaluated in a complete functional organ that preserves its physiological function under an artificial laboratory setup. Furthermore, it allows the measurement and manipulation of pulmonary mechanical ventilation to analyze the components of pulmonary physiology such as airway resistance, total vascular resistance, gas exchange, and edema formation, which to date cannot be measured precisely in vivo on lab animals2. Notably, the composition of the solution with which the lung is perfused can be fully controlled, enabling the addition of substances to evaluate their effects in real-time and sample collection from perfusion for further study11. Researchers working with the isolated lung system should bear in mind that mechanical ventilation causes decay of the pulmonary tissue shortening its useful time. This progressive fall in mechanical parameters can be significantly delayed by hyperinflating the lungs occasionally during the time of the experiment4. Still, the preparation cannot usually last more than eight hours. Another consideration for the ex-vivo lung perfusion system is the absence of central nervous regulation and lymphatic drainage. The effects of their absence are not yet fully understood and could potentially be a source of bias in certain experiments. 
The isolated lung perfusion system technique can be performed in the rabbit model with a high degree of consistency and reproducibility. This work describes the technical and surgical procedures for the implementation of the ex-vivo isolated lung perfusion technique as developed for the rabbit model at Instituto Nacional de Enfermedades Respiratorias in Mexico City, intending to share the insights and provide a clear guide on key steps in the application of this experimental model.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2-Stop Tygon E-Lab Tubing, 3.17 mm ID, 12/pack, Black/White</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-1864</td>
			<td></td>
		</tr>
		<tr>
			<td>Adapter for Positive Pressure Ventilation on IPL-4</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4312</td>
			<td></td>
		</tr>
		<tr>
			<td>Adapter for Positive Pressure Ventilation on IPL-4</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4312</td>
			<td></td>
		</tr>
		<tr>
			<td>Alternative Pressure-Free Gas Supply for IPL-4: To supply the trachea with gas mixture different from room air during negative ventilation</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4309</td>
			<td></td>
		</tr>
		<tr>
			<td>Base Unit for the Rabbit to Fetal Pig Isolated Perfused Lung</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4138</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum A2:D41albumin lyophilized powder</td>
			<td>sigma</td>
			<td>3912</td>
			<td>500 g</td>
		</tr>
		<tr>
			<td>Calcium chloride, CaCl2&#183;2H2O.</td>
			<td>JT Baker</td>
			<td>10035-04-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryogenic vials</td>
			<td>Corning</td>
			<td>430659</td>
			<td>2 mL</td>
		</tr>
		<tr>
			<td>D-glucosa, C6H12O6.</td>
			<td>sigma</td>
			<td>G5767</td>
			<td></td>
		</tr>
		<tr>
			<td>Differential Low Pressure Transducer DLP2.5, Range +- 2.5 cmH2O, HSE Connector</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-3882</td>
			<td></td>
		</tr>
		<tr>
			<td>Differential Pressure Transducer MPX, Range +- 100 cmH2O, HSE Connector</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-0064</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf tubes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol absolute HPLC grade</td>
			<td>Caledon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon tubes</td>
			<td></td>
			<td></td>
			<td>14 mL</td>
		</tr>
		<tr>
			<td>Harvard Peristaltic Pump P-230 (Complete with Control Box and P-230 Motor Drive)</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>70-7001</td>
			<td></td>
		</tr>
		<tr>
			<td>Heated Linear Pneumotachometer 0 to 10 L/min flow range</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>59-9349</td>
			<td></td>
		</tr>
		<tr>
			<td>Heater Controller for Single Pneumotachometer 230 VAC, 50 Hz</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>59-9703</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>PISA</td>
			<td></td>
			<td>5000 UI</td>
		</tr>
		<tr>
			<td>HPLC Column (C18 100A 5U)</td>
			<td>Alltech</td>
			<td>98121213</td>
			<td>150 mm x 4.6 mm</td>
		</tr>
		<tr>
			<td>Hydrophilic Syringe Filter</td>
			<td>Millex</td>
			<td>SLLGR04NL</td>
			<td>4 mm</td>
		</tr>
		<tr>
			<td>IPL-4 Core System for Isolated Rabbit to Fetal Pig Lung, 230</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4296</td>
			<td></td>
		</tr>
		<tr>
			<td>IPL-4 Core System for Isolated Rabbit to Fetal Pig Lung, 230 V</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4296</td>
			<td></td>
		</tr>
		<tr>
			<td>Jacketed Glass Reservoir for Buffer Solution, with Frit and Tubing, 6.0 L</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-0322</td>
			<td></td>
		</tr>
		<tr>
			<td>Lauda Thermostatic Circulator, Type E-103, 230 V/50 Hz, 3 L Bath Volume, Temperature Range 20 to 150&#176;C</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-0125</td>
			<td></td>
		</tr>
		<tr>
			<td>Left Atrium Cannula for Rabbit with Basket, OD 5.9 mm</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4162</td>
			<td></td>
		</tr>
		<tr>
			<td>Low Range Blood Pressure Transducer P75 for PLUGSYS Module</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-0020</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate heptahydrate, MgSO4&#183;7H2O</td>
			<td>JT Baker</td>
			<td>10034-99-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge Tube</td>
			<td>Corning</td>
			<td>430909</td>
			<td></td>
		</tr>
		<tr>
			<td>Negative Pressure Ventilation Control Option with Pressure Regulator for IPL-4</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4298</td>
			<td></td>
		</tr>
		<tr>
			<td>New Zeland rabbits</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PISABENTAL (Pentobarbital sodium)</td>
			<td>PISA</td>
			<td>Q-7833-215</td>
			<td></td>
		</tr>
		<tr>
			<td>PLUGSYS Case, Type 603* 7</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-0045</td>
			<td></td>
		</tr>
		<tr>
			<td>PLUGSYS TCM Time Counter Module</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-1750</td>
			<td></td>
		</tr>
		<tr>
			<td>PLUGSYS Transducer Amplifier Module (TAM-A)</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-0065</td>
			<td></td>
		</tr>
		<tr>
			<td>PLUGSYS Transducer Amplifier Module (TAM-D)</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-1793</td>
			<td></td>
		</tr>
		<tr>
			<td>PLUGSYS VCM-4R Ventilation Control Module with Pressure Regulator</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-1755</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride, KCl.</td>
			<td>JT Baker</td>
			<td>3040-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium dihydrogen phosphate, KH2PO4</td>
			<td>JT Baker</td>
			<td>7778-77-0</td>
			<td></td>
		</tr>
		<tr>
			<td>PROCIN (Xylacine clorhydrate)</td>
			<td>PISA</td>
			<td>Q-7833-099</td>
			<td></td>
		</tr>
		<tr>
			<td>Pulmonary Artery Cannula for Rabbit with Basket, OD 4.6 mm</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4161</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel knife</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Serotonin 5-HT</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Servo Controller for Perfusion (SCP</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-2806</td>
			<td></td>
		</tr>
		<tr>
			<td>Snap Cap Microcentrifuge Tube</td>
			<td>Costar</td>
			<td>3620</td>
			<td>1.7 mL</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate, NaHCO3</td>
			<td>sigma</td>
			<td>S6014</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride, NaCl.</td>
			<td>sigma</td>
			<td>S9888</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical gloves No. 7 1/2</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical gloves No. 8</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Taygon tubes</td>
			<td>Masterflex</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tracheal Cannula for Rabbit, OD 5.0 mm</td>
			<td>Hugo Sachs Elektronik (HSE)</td>
			<td>73-4163</td>
			<td></td>
		</tr>
	</tbody>
</table>

isolated lung perfusion system in the rabbit model

The isolated lung perfusion system has been widely used in pulmonary research, contributing to elucidate the lungs&#39; inner workings, both micro and macroscopically. This technique is useful in the characterization of pulmonary physiology and pathology by measuring metabolic activities and respiratory functions, including interactions between circulatory substances and the effects of inhaled or perfused substances, as in drug testing. While in vitro methods involve the slicing and culturing of tissues, the isolated ex vivo lung perfusion system allows to work with a complete functional organ making possible the study of a continuous physiological function while recreating ventilation and perfusion. However, it should be noted that the effects of the absence of central innervation and lymphatic drainage still have to be fully assessed. This protocol aims to describe the assembly of the isolated lung apparatus, followed by the surgical extraction and cannulation of lungs and heart from experimental lab animals, as well as to display the perfusion technique and signal processing of data. The average viability of the isolated lung ranges between 5-8 h; during this period, the pulmonary capillary permeability increases, causing edema and lung injury. The functionality of preserved pulmonary tissue is measured by the capillary filtration coefficient (Kfc), used to determine the extent of pulmonary edema through time.

The isolated lung perfusion system allows organ manipulation for biopsy, sample collection from perfusion, and real-time data collection of physiological parameters. The isolated system can be used to test many hypotheses involving different functions and lung phenomena, from metabolic and enzymatic activity to edema formation and preservation periods for lung transplants.
Figure 1&#160;displays a diagram of the fully assembled isolated lung perfusion system along...

Watch this Scientific Journal Video about Isolated Lung Perfusion System in the Rabbit Model at JoVE.com

Isolated Lung Perfusion System in the Rabbit Model

The isolated rabbit lung preparation is a gold standard tool in pulmonary research. This publication aims to describe the technique as developed for the study of physiological and pathological mechanisms involved in airway reactivity, lung preservation, and preclinical research in lung transplantation and pulmonary edema.

The isolated lung perfusion system has been widely used in pulmonary research, contributing to elucidate the lungs&#39; inner workings, both micro and ...

isolated-lung-perfusion-system-in-the-rabbit-model

Research

JoVE Journal

Medicine

3.5K Views.  Instituto Nacional de Enfermedades Respiratorias. The isolated rabbit lung preparation is a gold standard tool in pulmonary research. This publication aims to describe the technique as developed for the study of physiological and pathological mechanisms involved in airway reactivity, lung preservation, and preclinical research in lung transplantation and pulmonary edema. 

Video: Isolated Lung Perfusion System in the Rabbit Model

NADH Fluorescence Imaging of Isolated Biventricular Working Rabbit Hearts

Since its inception by Langendorff1, the isolated perfused heart remains a prominent tool for studying cardiac physiology2. However, it is not well-suited for studies of cardiac metabolism, which require the heart to perform work within the context of physiologic preload and afterload pressures. Neely introduced modifications to the Langendorff technique to establish appropriate left ventricular (LV) preload and afterload pressures3. The model is known as the isolated LV working heart model and has been used extensively to study LV performance and metabolism4-6. This model, however, does not provide a properly loaded right ventricle (RV). Demmy et al. first reported a biventricular model as a modification of the LV working heart model7, 8. They found that stroke volume, cardiac output, and pressure development improved in hearts converted from working LV mode to biventricular working mode8. A properly loaded RV also diminishes abnormal pressure gradients across the septum to improve septal function. Biventricular working hearts have been shown to maintain aortic output, pulmonary flow, mean aortic pressure, heart rate, and myocardial ATP levels for up to 3 hours8.
When studying the metabolic effects of myocardial injury, such as ischemia, it is often necessary to identify the location of the affected tissue. This can be done by imaging the fluorescence of NADH (the reduced form of nicotinamide adenine dinucleotide)9-11, a coenzyme found in large quantities in the mitochondria. NADH fluorescence (fNADH) displays a near linearly inverse relationship with local oxygen concentration12 and provides a measure of mitochondrial redox state13. fNADH imaging during hypoxic and ischemic conditions has been used as a dye-free method to identify hypoxic regions14, 15 and to monitor the progression of hypoxic conditions over time10.
The objective of the method is to monitor the mitochondrial redox state of biventricular working hearts during protocols that alter the rate of myocyte metabolism or induce hypoxia or create a combination of the two. Hearts from New Zealand white rabbits were connected to a biventricular working heart system (Hugo Sachs Elektronik) and perfused with modified Krebs-Henseleit solution16 at 37 &deg;C. Aortic, LV, pulmonary artery, and left &amp; right atrial pressures were recorded. Electrical activity was measured using a monophasic action potential electrode. To image fNADH, light from a mercury lamp was filtered (350&plusmn;25 nm) and used to illuminate the epicardium. Emitted light was filtered (460&plusmn;20 nm) and imaged using a CCD camera. Changes in the epicardial fNADH of biventricular working hearts during different pacing rates are presented. The combination of the heart model and fNADH imaging provides a new and valuable experimental tool for studying acute cardiac pathologies within the context of realistic physiological conditions.

The objective is to monitor the mitochondrial redox state of isolated hearts within the context of physiologic preload and afterload pressures. A biventricular working rabbit heart model is presented. High spatiotemporal resolution fluorescence imaging of NADH is used to monitor the mitochondrial redox state of epicardial tissue.

Since its inception by Langendorff1, the isolated perfused heart remains a prominent tool for studying cardiac physiology2. However, it is not well-suited ...

Optimized System for Cerebral Perfusion Monitoring in the Rat Stroke Model of Intraluminal Middle Cerebral Artery Occlusion

The translational potential of pre-clinical stroke research depends on the accuracy of experimental modeling. Cerebral perfusion monitoring in animal models of acute ischemic stroke allows to confirm successful arterial occlusion and exclude subarachnoid hemorrhage. Cerebral perfusion monitoring can also be used to study intracranial collateral circulation, which is emerging as a powerful determinant of stroke outcome and a possible therapeutic target. Despite a recognized role of Laser Doppler perfusion monitoring as part of the current guidelines for experimental cerebral ischemia, a number of technical difficulties exist that limit its widespread use. One of the major issues is obtaining a secure and prolonged attachment of a deep-penetration Laser Doppler probe to the animal skull. In this video, we show our optimized system for cerebral perfusion monitoring during transient middle cerebral artery occlusion by intraluminal filament in the rat. We developed in-house a simple method to obtain a custom made holder for twin-fibre (deep-penetration) Laser Doppler probes, which allow multi-site monitoring if needed. A continuous and prolonged monitoring of cerebral perfusion could easily be obtained over the intact skull.

Cerebral perfusion monitoring has been demonstrated to improve accuracy in ischemic stroke models. Technical difficulties often limit the use of this essential tool for cerebrovascular research. In this video, an optimized system is shown to obtain a single or multi-site hemodynamic monitoring during intraluminal middle cerebral artery occlusion in rats.

The translational potential of pre-clinical stroke research depends on the accuracy of experimental modeling. Cerebral perfusion monitoring in animal ...

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

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

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

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

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

Isolated Hepatic Perfusion as a Treatment for Liver Metastases of Uveal Melanoma

Isolated hepatic perfusion (IHP) is a procedure where the liver is surgically isolated and perfused with a high concentration of the chemotherapeutic agent melphalan. Briefly, the procedure starts with the setup of a percutaneous veno-venous bypass from the femoral vein to the external jugular vein. Via a laparotomy, catheters are then inserted into the proper hepatic artery and the caval vein. The portal vein and the caval vein, both supra- and infrahepatically, are then clamped. The arterial and venous catheters are connected to a heart lung machine and the liver is perfused with melphalan (1 mg/kg body weight) for 60 min. This way it is possible to locally perfuse the liver with a high dose of a chemotherapeutic agent, without leakage to the systemic circulation.
In previous studies including patients with isolated liver metastases of uveal melanoma, an overall response rate of 33-100% and a median survival between 9 and 13 months, have been reported. The aim of this protocol is to give a clear description of how to perform the procedure and to discuss IHP as a treatment option for liver metastases of uveal melanoma.

Here, we present a protocol how to perform an isolated liver perfusion (IHP) with melphalan and we also discuss IHP as a treatment option for liver metastases of uveal melanoma.

Isolated hepatic perfusion (IHP) is a procedure where the liver is surgically isolated and perfused with a high concentration of the chemotherapeutic ...

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute respiratory distress syndrome (ARDS), characterized by neutrophilic alveolitis, and injury to both alveolar epithelium and vascular endothelium. Clinically, ARDS is defined by acute onset of hypoxemia, bilateral patchy pulmonary infiltrates and non-cardiogenic pulmonary edema. Human studies have provided us with valuable information about the physiological and inflammatory changes in the lung caused by ARDS, which has led to various hypotheses about the underling mechanisms. Unfortunately, difficulties determining the etiology of ARDS, as well as a wide range of pathophysiology have resulted in a lack of critical information that could be useful in developing therapeutic strategies.
Translational animal models are valuable when their pathogenesis and pathophysiology accurately reproduce a concept proven in both in vitro and clinical settings. Although large animal models (e.g., sheep) share characteristics of the anatomy of human trachea-bronchial tree, murine models provide a host of other advantages including: low cost; short reproductive cycle lending itself to greater data acquisition; a well understood immunologic system; and a well characterized genome leading to the availability of a variety of gene deletion and transgenic strains. A robust model of low pH induced ARDS requires a murine ALI that targets mainly the alveolar epithelium, secondarily the vascular endothelium, as well as the small airways leading to the alveoli. Furthermore, a reproducible injury with wide differences between different injurious and non-injurious insults is important.
The murine gastric acid aspiration model presented here using hydrochloric acid employs an open tracheostomy and recreates a pathogenic scenario that reproduces the low pH pneumonitis injury in humans. Additionally, this model can be used to examine interaction of a low pH insult with other pulmonary injurious entities (e.g., food particles, pathogenic bacteria).

This protocol induces acute lung injury in a mouse that has close fidelity to the pathogenesis of acid pneumonitis observed in humans. We generate a maximal acute nonlethal low pH lung injury and account for differences in rodent-human anatomic respiratory structure using an open tracheostomy coupled with circumferential pressure release.

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute ...

Transthoracic Echocardiographic Examination in the Rabbit Model

Large animal models such as the rabbit are valuable for translational preclinical research. Rabbits have a similar cardiac electrophysiology compared to that of humans and that of other large animal models such as dogs and pigs. However, the rabbit model has the additional advantage of lower maintenance costs compared to other large animal models. The longitudinal evaluation of cardiac function using echocardiography, when appropriately implemented, is a useful methodology for preclinical assessment of novel therapies for heart failure with reduced ejection fraction (e.g. cardiac regeneration). The correct use of this non-invasive tool requires the implementation of a standardized examination protocol following international guidelines. Here we describe, step by step, a detailed protocol supervised by veterinary cardiologists for performing echocardiography in the rabbit model, and demonstrate how to correctly obtain the different echocardiographic views and imaging planes, as well as the different imaging modes available in a clinical echocardiography system routinely used in human and veterinary patients.

Here we describe, step by step, a detailed protocol for performing echocardiography in the rabbit model. We show how to correctly obtain the different echocardiographic views and imaging planes, as well as the different imaging modes available in a clinical echocardiography system routinely used in human and veterinary patients.

Large animal models such as the rabbit are valuable for translational preclinical research. Rabbits have a similar cardiac electrophysiology compared to ...

Laser Doppler Perfusion Imaging in the Mouse Hindlimb

Blood flow recovery is a critical outcome measure after experimental hindlimb ischemia or ischemia-reperfusion. Laser Doppler perfusion imaging (LDPI) is a common, noninvasive, repeatable method for assessing blood flow recovery. The technique calculates overall blood flow in the sampled tissue from the Doppler shift in frequency caused when a laser hits moving red blood cells. Measurements are expressed in arbitrary perfusion units, so the contralateral non-intervened upon leg is usually used to help control measurements. Measurement depth is in the range of 0.3-1 mm; for hindlimb ischemia, this means that dermal perfusion is assessed. Dermal perfusion is dependent on several factors&#8212;most importantly skin temperature and anesthetic agent, which must be carefully controlled to result in reliable readings. Furthermore, hair and skin pigmentation can alter the ability of the laser to either reach or penetrate to the dermis. This article demonstrates the technique of LDPI in the mouse hindlimb.

Here, we present a protocol that demonstrates the technique and necessary controls for Laser Doppler perfusion imaging to measure blood flow in the mouse hindlimb.

Blood flow recovery is a critical outcome measure after experimental hindlimb ischemia or ischemia-reperfusion. Laser Doppler perfusion imaging (LDPI) is ...

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

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

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

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

Establishment of the KOA Model in Rabbits Using the Modified Videman Method

Application of Acupotomy in a Knee Osteoarthritis Model in Rabbit

Knee osteoarthritis (KOA) is one of the most frequently encountered diseases in the orthopedic department, which seriously reduces the quality of life of people with KOA. Among several pathogenic factors, the biomechanical imbalance of the knee joint is one of the main causes of KOA. Acupotomology believes that restoring the mechanical balance of the knee joint is the key to treating KOA. Clinical studies have shown that acupotomy can effectively reduce pain and improve knee mobility by reducing adhesion, contracture of soft tissues, and stress concentration points in muscles and tendons around the knee joint. 
In this protocol, we used the modified Videman method to establish a KOA model by immobilizing the left hindlimb in a straight position. We have outlined the method of operation and the precautions related to acupotomy in detail and evaluated the efficacy of acupotomy in conjunction with the theory of "Modulating Muscles and Tendons to Treat Bone Disorders" through the detection of the mechanical properties of quadriceps femoris and tendon, as well as cartilage mechanics and morphology. The results show that acupotomy has a protective effect on cartilage by adjusting the mechanical properties of the soft tissues around the knee joint, improving the cartilage stress environment, and delaying cartilage degeneration.

Author Spotlight: Investigating the Mechanism of Action of Acupotomy in Treating Knee Osteoarthritis

In this protocol, a knee osteoarthritis model was prepared using the modified Videman method, and the operation procedures and precautions of acupotomy are detailed. The effectiveness of acupotomy has been demonstrated by testing the mechanical properties of quadriceps femoris and tendon and the mechanical and morphological properties of cartilage.

Knee osteoarthritis (KOA) is one of the most frequently encountered diseases in the orthopedic department, which seriously reduces the quality of life of ...

Advancing Lung Transplantation with PolyhHb-Based Perfusion in Rat EVLP Models

Exploring Alternative Perfusion Solutions Using Next-Generation Polymerized Hemoglobin-Based Oxygen Carriers in a Model of Rat Ex Vivo Lung Perfusion

Lung transplantation is hampered by the lack of suitable donors. Previously, donors that were thought to be marginal or inadequate were discarded. However, new and exciting technology, such as ex vivo lung perfusion (EVLP), offers lung transplant providers extended assessment for marginal donor allografts. This dynamic assessment platform has led to an increase in lung transplantation and has allowed providers to use donors that were previously discarded, thus expanding the donor pool. Current perfusion techniques use cellular or acellular perfusates, and both have distinct advantages and disadvantages. Perfusion composition is critical to maintaining a homeostatic environment, providing adequate metabolic support, decreasing inflammation and cellular death, and ultimately improving organ function. Perfusion solutions must contain sufficient protein concentration to maintain appropriate oncotic pressure. However, current perfusion solutions often lead to fluid extravasation through the pulmonary endothelium, resulting in inadvertent pulmonary edema and damage. Thus, it is necessary to develop novel perfusion solutions that prevent excessive damage while maintaining proper cellular homeostasis. Here, we describe the application of a polymerized human hemoglobin (PolyhHb)-based oxygen carrier as a perfusate and the protocol in which this perfusion solution can be tested in a model of rat EVLP. The goal of this study is to provide the lung transplant community with key information in designing and developing novel perfusion solutions, as well as the proper protocols to test them in clinically relevant translational transplant models.

Author Spotlight: Utilizing Next-Generation Polymerized Human Hemoglobin for Improved Donor Lung Evaluation and Preservation in Rats

Here, we describe the application of a polymerized human hemoglobin (PolyhHb)-based oxygen carrier as a perfusate and the protocol in which this perfusion solution can be tested in a model of rat ex vivo lung perfusion.