Male Sprague-Dawley rats (180-270 g body weight) were purchased commercially (e.g., Envigo) and were housed under pathogen-free conditions at The Ohio State University Animal Facility. All procedures were humanely performed according to the NIH and the National Research Council&#39;s Guide for the Humane Care and Use of Laboratory Animals and with the approval of The Ohio State University Institutional Animal Care and Use Committee (IACUC Protocol # 2012A00000135-R2).
1. Initial setup
<ol>
	<li>Set up surgical devices. 
	NOTE: This&#160;is a non-survival surgery. If survival surgery is to be performed, sterile instruments and barrier precautions would need to be undertaken.&#160;
	<ol>
		<li>Turn on the heart rate/oxygen saturation monitoring equipment and warming board to 42 &#176;C.</li>
		<li>Turn on the ventilation and anesthesia machine to pre-warm the isoflurane evaporator. 
		NOTE: Use a tidal volume (Td) of 7.2 mL/kg, a positive-end expiratory pressure (PEEP) of 2 cmH2O, and a respiratory rate of 80 bpm.</li>
		<li>Fill the anesthesia syringe with 10 mL of liquid isoflurane and mount the syringe onto the ventilation and anesthesia machine.</li>
		<li>Turn on the surgical microscope with the height and focus adjusted to the microsurgeon&#39;s preferences.</li>
		<li>Turn on the electrocautery device.</li>
	</ol>
	</li>
	<li>Prepare and lay out surgical tools (Figure 1). 
	NOTE: All surgical tools were autoclaved at 121 &#176;C for 30 min.</li>
	<li>Collect and record body weight of donor and recipient rats.</li>
	<li>Use Table 1 and the rat&#39;s body weight to determine the correct gauge of angio-catheter (20, 18, 16 14, or 12 G) to use to make cuffs.</li>
	<li>Prepare cuffs for pulmonary artery (PA), bronchus (Br), and pulmonary vein (PV) using the size guide based on body weight (Table 1 and Figure 2).
	<ol>
		<li>Place angio-catheter sized 20 G, 18 G, 16 G, 14 G, or 12 G (Figure 2A-E) on a sterile surface under the surgical microscope.</li>
		<li>Then use a rib-back surgical blade #11 (Figure 2F) to cut the angio-catheter at a 90&#176; angle to form a 2 mm in length cuff body with a 1 mm X 1 mm tab (width x height) at the top of the cuff body (Figure 2G).</li>
		<li>Store cuffs in sterile saline until ready to use.</li>
	</ol>
	</li>
	<li>Prepare solutions.
	<ol>
		<li>Prepare a mixture of ketamine and xylazine in a sterile injection vial by adding 1 mL of xylazine (100 mg/mL) to 10 mL ketamine (100 mg/mL). 
		NOTE: The expiration date for this cocktail is determined&#160;using the earliest expiration date of the components used.</li>
		<li>Draw into syringes the proper dosage for the rats (0.1 mL of the ketamine/xylazine mixture per 100 g of rat&#39;s body weight; e.g., for a 200 g rat, 0.2 mL of ketamine/xylazine mixture would be delivered). 
		NOTE: This dosage will deliver 91 mg/kg of ketamine and 9.1 mg/kg of xylazine to the rat and should keep a rat sedated for 60-80 min.</li>
		<li>Prepare heparin which will be delivered at a dose of 1,000 U/kg.</li>
		<li>Store the saline, PBS, and preservation solution on ice (Table of Materials).</li>
	</ol>
	</li>
</ol>
2. Donor rat preparation
<ol>
	<li>Induce anesthesia in the donor rat by intraperitoneally injecting the ketamine and xylazine mixture and waiting ~10 min for a surgical plane of anesthesia to develop that can be assessed by lack of response to toe-pinch.</li>
	<li>Shave the incision area using electronic clippers.</li>
	<li>Place the donor rat in a supine position on the surgical warming board and wipe the incision area with a sterile gauze soaked with betadine. Then wipe the area with a 70% isopropyl alcohol swab.&#160;Repeat 3 times.</li>
	<li>Make a 3 to 4 cm midline skin incision mid-neck using scissors and carefully dissect out subcutaneous tissues and muscles using forceps (instead of scissors to avoid bleeding).</li>
	<li>For endotracheal intubation, thread 4-0 silk suture around the trachea and insert a 16 G angio-catheter into the trachea. Tie the suture around the trachea tightly with a double knot, and then finish with a single knot to hold the angio-catheter in place.</li>
	<li>Connect the angio-catheter to the ventilator and maintain a surgical plane of anesthesia in the rat with 1-2% of isoflurane.</li>
	<li>Perform a laparo-sternotomy as a combined midline and transverse incision using scissors.</li>
	<li>Inject heparin (1,000 U/kg) with an insulin syringe via the inferior vena cava (IVC) and allow 10 min for systemic circulation. 
	NOTE: This administration of heparin prevents blood clots in the donor lung.</li>
	<li>Dissect the diaphragm carefully by cutting along the thoracic arch, and then expose the thoracic cavity by following the sternum to the neck.</li>
</ol>
3. Donor lung warm ischemia and procurement
<ol>
	<li>Euthanize the donor rat by cutting the IVC.</li>
	<li>While lungs are still ventilating, cut both right and left auricles with micro dissecting spring scissors and gravity flush lungs with 20 mL of preservation solution in a syringe hanging at 28 cmH2O by gravity connected to tubing and an 18 G angio-catheter that is introduced directly through the pulmonary artery.</li>
	<li>Disconnect the ventilator from the endotracheal tube and connect it to a 5 mL syringe filled with a proper volume of air based on body weight. 
	NOTE: Volume of air to inflate lungs can be calculated as with twice of tidal volume (Td = 7.2 mL/kg), e.g., a 200 g rat would have a Td of 1.44 mL and multiplying it by 2 would equal 2.88 mL of air needed to inflate lungs.</li>
	<li>Inflate donor&#39;s lungs.</li>
	<li>Put a Yasargil clamp on the trachea to keep lungs inflated, and cover the lungs and heart with sterile cotton gauze. Moisturize the gauze with saline, wrap the donor rat with an under pad, and leave on the warming surgery board for 1 h to induce warm ischemia in the lungs (Figure 3).</li>
	<li>After 1 h of warm ischemia, excise the heart-lung block with micro dissecting spring scissors and forceps and place on sterile gauze dampened with ice-cold PBS on a sterile Petri dish on ice. 
	NOTE: All of the following steps should occur while lungs are on the Petri dish on ice.</li>
	<li>Carefully incise pulmonary ligaments with micro dissecting spring scissors to separate the left lung from the esophagus and the post-caval lobe.</li>
	<li>Carefully trim the left lung hilar area with Vannas-Tubingen spring scissor and procure the left PV, PA, and Br.</li>
	<li>Place cuffs on PV, PA, or Br (Figure 4A-C).
	<ol>
		<li>Use a mosquito hemostat to grab the cuff tab.</li>
		<li>Use fine forceps to grab the distal end of the PV, PA, or Br through the appropriate cuff body, evert the extra tissue around the cuff, and secure using 8-0 nylon suture. Use Vannas-Tubingen spring scissors to trim extra tissue and the cuff around the cuff body.</li>
	</ol>
	</li>
	<li>Keep the donor lung covered with gauze dampened with saline on the Petri dish on ice until it is ready to be transplanted into the recipient rat (Figure 4D). 
	&#8203;NOTE: Average cold ischemia time is 84 min &#177; 11 min S.D.</li>
</ol>
4. Recipient rat preparation
<ol>
	<li>Induce anesthesia in the recipient rat in the same manner as the donor rat by intraperitoneally injecting the ketamine and xylazine mixture (0.1 mL per 100 g rat) and waiting 10 min for a surgical plane of anesthesia to develop that can be assessed by lack of response to toe-pinch.</li>
	<li>Shave incision area using electronic clippers.</li>
	<li>Place the donor rat in a supine position and wipe the incision area with a sterile gauze soaked with betadine. Then wipe the area with a 70% isopropyl alcohol swab.&#160;Repeat 3 times.</li>
	<li>Before the rat is secured to the ventilation machine, draw lines on the rat&#39;s chest in preparation for finding the 4th intercostal space.
	<ol>
		<li>Measure the chest from the suprasternal notch to the xiphoid process and draw a line (Figure 5A).</li>
		<li>At the middle of this line, draw a line along the left side of the chest that measures half of the measurement from the suprasternal notch to the xiphoid process (Figure 5A and B).</li>
	</ol>
	</li>
	<li>Intubate the recipient using a 16 G angio-catheter via visualization using the fiber optic cable connected to an LED light from the endotracheal intubation kit.</li>
	<li>Connect the angio-catheter to the ventilator and maintain a surgical plane of anesthesia with 1-2% isoflurane.</li>
	<li>To find the 4th intercostal space, find the area of the chest wall where a strong palpable cardiac impulse can be felt (Figure 5C, red circle).</li>
	<li>At this location, incise the skin with scissors and the muscle with micro dissecting spring scissors, and use the retractor to open the 4th intercostal space as widely as possible (Figure 5D and E). 
	NOTE: Use electrical cautery to avoid or stop bleeding during the muscle dissection.</li>
	<li>Once the intercostal space is opened widely, carefully dissect the ligaments around the recipient&#39;s left lung using Vannas-Tubingen spring scissors and pull the lung out of chest area using sterile cotton swabs and forceps.</li>
	<li>Place sterile gauze around the left lung and hold it with a Dieffenbach bulldog clamp.</li>
	<li>Apply a Yasargil clamp on the left lung hilar area as proximally as possible.</li>
</ol>
5. Anastomoses
<ol>
	<li>Pulmonary vein (PV) anastomosis
	<ol>
		<li>Place 7-0 nylon suture around the recipient&#39;s PV.</li>
		<li>Incise the recipient&#39;s PV using the Vannas-Tubingen spring scissors by transversely cutting the upper and lower segmental veins as distally as possible and flush out blood with 0.2 mL of heparinized saline (1 U/mL) using an insulin syringe.</li>
		<li>Place the donor lung still wrapped with ice-cold wet sterile gauze into the thoracic cavity.</li>
		<li>Insert the donor&#39;s cuffed PV into the recipient&#39;s PV, and then secure with the prepositioned 7-0 nylon suture (Figure 6).</li>
	</ol>
	</li>
	<li>Bronchial (Br) anastomosis
	<ol>
		<li>Place 7-0 nylon suture around the recipient&#39;s Br.</li>
		<li>Incise the recipient&#39;s Br by cutting the upper and lower segmental airways transversely as distally as possible with Vannas-Tubingen spring scissors.</li>
		<li>Insert the donor&#39;s cuffed Br into recipient&#39;s Br and secure with the prepositioned 7-0 nylon suture (Figure 6).</li>
	</ol>
	</li>
	<li>Pulmonary artery (PA) anastomosis:
	<ol>
		<li>Place 7-0 nylon suture around the recipient&#39;s PA.</li>
		<li>Incise the recipient&#39;s PA from its adventitial sheath, incise half of the vessel&#39;s circumference with Vannas-Tubingen spring scissors, and then flush out blood in the PA with 0.2 mL heparinized saline (1 U/mL) using an insulin syringe.</li>
		<li>Insert the donor&#39;s cuffed PA into the recipient&#39;s PA and secure with the prepositioned 7-0 nylon suture (Figure 6).</li>
	</ol>
	</li>
</ol>
6. Reperfusion
<ol>
	<li>Remove the Yasargil clamp on the hilum to allow for reperfusion and ventilation of the transplanted donor lung (Figure 7).</li>
	<li>Dissect out the recipient&#39;s native left lung using micro dissecting spring scissors and forceps.</li>
	<li>Carefully reposition the transplanted left lung into the recipient&#39;s thorax.</li>
	<li>Close the thoracotomy incision by using 6-0 nylon suture.
	<ol>
		<li>Place three 6-0 nylon sutures with simple double knots around the ribs superior to the 4th rib and inferior to the 5th rib (Figure 8A).</li>
		<li>Use hemostats to gather the three sutures together (Figure 8B).</li>
		<li>Increase the PEEP to 6 cmH2O in the ventilation settings.</li>
		<li>Tie together all three knots at the same time by pulling away to close the wound (Figure 8C).</li>
		<li>Decrease PEEP to 2 cmH2O immediately.</li>
		<li>Close skin with 6-0 nylon suture. 
		&#8203;NOTE: Our laboratory studies the acute phase post-transplantation, so the recipient rat in this model is survived for 3 h post-transplantation under ventilation and anesthesia and then samples are collected.</li>
	</ol>
	</li>
</ol>
7. Collection of experimental specimens (plasma, lung tissue)
<ol>
	<li>For control samples, collect the donor&#39;s right lobes after the 3 h reperfusion period is initiated.
	<ol>
		<li>Snap-freeze the superior lobe and post-caval lobe for protein or RNA expression analyses, preserve the middle lobe for histology, and use the inferior lobe for wet-to-dry weight ratio (Figure 9A).</li>
	</ol>
	</li>
	<li>At 10 min before the end of the 3h reperfusion time, prepare to harvest the recipient samples by injecting heparin (1,000 U/kg) with an insulin syringe into the jugular vein. 
	NOTE: This administration of heparin prevents blood clots in the lungs and allows for a more thorough flushing at the time of procurement.</li>
	<li>Plasma collection
	<ol>
		<li>At the end of the 3 h reperfusion period, collect 1 mL of blood with a syringe via the IVC.</li>
		<li>Store on ice and then centrifuge at 2,000 x g for 10 min to harvest plasma.</li>
	</ol>
	</li>
	<li>Euthanize the recipient rat by cutting the IVC to allow for exsanguination.</li>
	<li>Dissect the diaphragm along the thoracic arch and expose the thoracic cavity by dissecting out the rib cage.</li>
	<li>Collect BAL fluid from the native or transplanted lungs if desired (optional). 
	NOTE: If wet-to-dry weight ratio or histology is being performed on the lung, BAL fluid collection should not be performed since it can affect results.
	<ol>
		<li>Thread 4-0 silk suture around trachea and tie a tight double knot around the trachea and intubation tube to prevent fluid leakage.</li>
		<li>Calculate the amount of ice-cold PBS for BAL fluid collection from the right lobes and left lobe. 
		NOTE: The volume ratio for the right lung is 63% while the volume ratio for the left lung is 37%16. Therefore, to determine the amount of PBS to instill into each side, the volume should be calculated as twice the tidal volume (Td = 7.2 mL/kg) multiplied by 63% for the right lung and 37% for left lung.</li>
		<li>Place a Yasargil clamp on the left lung hilar area (Figure 10A), and, with a syringe connected to the angio-catheter, instill the calculated amount of ice-cold PBS into the right lung (recipient&#39;s native lobes) and collect the BAL fluid by pulling up gently on the syringe plunger. Perform twice. 
		NOTE: One should expect 70-80% recovery of instilled fluid.</li>
		<li>Remove the Yasargil clamp on the left lung and place the clamp on the right lung hilar area (Figure 10B).</li>
		<li>Collect BAL fluid from transplanted left lobe in the same way as it was collected for the right lobes, and then remove the clamp on right lung hilar area.</li>
	</ol>
	</li>
	<li>Cut the right and left auricles with micro dissecting spring scissors and flush lungs by gravity through the PA using an 18 G angio-catheter attached to tubing and a syringe with 20 mL of pre-chilled preservation solution hanging at 28 cmH2O.</li>
	<li>Collect samples from the recipient&#39;s lung.
	<ol style="margin-left: 40px;">
		<li>Snap-freeze the superior lobe and post-caval lobe for protein or RNA expression analyses, preserve the middle lobe for histology, and use the inferior lobe for wet-to-dry weight ratio) (Figure 9A).</li>
		<li>Divide the left transplanted left lobe into three parts: the upper region collected for snap-frozen, the middle region for histology, and the lower region for wet-to-dry weight ratio (Figure 9B).</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Mizuta, T., Kawaguchi, A., Nakahara, K., Kawashima, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simplified+rat+lung+transplantation+using+a+cuff+technique.">Simplified rat lung transplantation using a cuff technique.</a> Journal of Thoracic and Cardiovascular Surgery. 97 (4), 578-581 (1989).</li><li>Zhai, W., 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=Simplified+rat+lung+transplantation+by+using+a+modified+cuff+technique.">Simplified rat lung transplantation by using a modified cuff technique.</a> Journal of Investigative Surgery. 21 (1), 33-37 (2008).</li><li>Goto, T., 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=Simplified+rat+lung+transplantation+using+a+new+cuff+technique.">Simplified rat lung transplantation using a new cuff technique.</a> Annals of Thoracic Surgery. 93 (6), 2078-2080 (2012).</li><li>Guo, 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=Improvements+of+surgical+techniques+in+a+rat+model+of+an+orthotopic+single+lung+transplant.">Improvements of surgical techniques in a rat model of an orthotopic single lung transplant.</a> European Journal of Medical Research. 18, 1 (2013).</li><li>Tian, D., Shiiya, H., Sato, M., Nakajima, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+lung+transplantation+model:+modifications+of+the+cuff+technique.">Rat lung transplantation model: modifications of the cuff technique.</a> Annals of Translational Medicine. 8 (6), 407 (2020).</li><li>Rajab, T. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anastomotic+techniques+for+rat+lung+transplantation.">Anastomotic techniques for rat lung transplantation.</a> World Journal of Transplantation. 8 (2), 38-43 (2018).</li><li>Reis, A., Giaid, A., Serrick, C., Shennib, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+outcome+of+rat+lung+transplantation+with+modification+of+the+nonsuture+external+cuff+technique.">Improved outcome of rat lung transplantation with modification of the nonsuture external cuff technique.</a> Journal of Heart and Lung Transplantation. 14 (2), 274-279 (1995).</li><li>Sugimoto, R., 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=Experimental+orthotopic+lung+transplantation+model+in+rats+with+cold+storage.">Experimental orthotopic lung transplantation model in rats with cold storage.</a> Surgery Today. 39 (7), 641-645 (2009).</li><li>Santana Rodriguez, 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=Technical+modifications+of+the+orthotopic+lung+transplantation+model+in+rats+with+brain-dead+donors.">Technical modifications of the orthotopic lung transplantation model in rats with brain-dead donors.</a> Archivos de Bronconeumolog&#237;a. 47 (10), 488-494 (2011).</li><li>Gielis, J. F., 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=A+murine+model+of+lung+ischemia+and+reperfusion+injury:+tricks+of+the+trade.">A murine model of lung ischemia and reperfusion injury: tricks of the trade.</a> Journal of Surgical Research. 194 (2), 659-666 (2015).</li><li>Rajab, T. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Techniques+for+lung+transplantation+in+the+rat.">Techniques for lung transplantation in the rat.</a> Experimental Lung Research. 45 (9-10), 267-274 (2019).</li><li>Iskender, I., 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=Effects+of+Warm+Versus+Cold+Ischemic+Donor+Lung+Preservation+on+the+Underlying+Mechanisms+of+Injuries+During+Ischemia+and+Reperfusion.">Effects of Warm Versus Cold Ischemic Donor Lung Preservation on the Underlying Mechanisms of Injuries During Ischemia and Reperfusion.</a> Transplantation. 102 (5), 760-768 (2018).</li><li>Lin, 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=Five-year+update+on+the+mouse+model+of+orthotopic+lung+transplantation:+Scientific+uses,+tricks+of+the+trade,+and+tips+for+success.">Five-year update on the mouse model of orthotopic lung transplantation: Scientific uses, tricks of the trade, and tips for success.</a> Journal of Thoracic Disease. 4 (3), 247-258 (2012).</li><li>Song, J. 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=Standardization+of+bronchoalveolar+lavage+method+based+on+suction+frequency+number+and+lavage+fraction+number+using+rats.">Standardization of bronchoalveolar lavage method based on suction frequency number and lavage fraction number using rats.</a> Toxicological Research. 26 (3), 203-208 (2010).</li><li>Chang, J. E., Kim, H. J., Yi, E., Jheon, S., Kim, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reduction+of+ischemia-reperfusion+injury+in+a+rat+lung+transplantation+model+by+low-concentration+GV1001.">Reduction of ischemia-reperfusion injury in a rat lung transplantation model by low-concentration GV1001.</a> European Journal of Cardio-Thoracic Surgery. 50 (5), 972-979 (2016).</li><li>De Backer, J. W., 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=Study+of+the+variability+in+upper+and+lower+airway+morphology+in+Sprague-Dawley+rats+using+modern+micro-CT+scan-based+segmentation+techniques.">Study of the variability in upper and lower airway morphology in Sprague-Dawley rats using modern micro-CT scan-based segmentation techniques.</a> Anatomical Record. 292 (5), 720-727 (2009).</li><li>Nelson, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Method+of+isolated+ex+vivo+lung+perfusion+in+a+rat+model:+lessons+learned+from+developing+a+rat+EVLP+program.">Method of isolated ex vivo lung perfusion in a rat model: lessons learned from developing a rat EVLP program.</a> Journal of Visualized Experiments. (96), e52309 (2015).</li><li>Suzuki, H., Fan, L., Wilkes, D. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+obliterative+bronchiolitis+in+a+murine+model+of+orthotopic+lung+transplantation.">Development of obliterative bronchiolitis in a murine model of orthotopic lung transplantation.</a> Journal of Visualized Experiments. (65), e3947 (2012).</li><li>Jia, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Treatment+of+acute+lung+injury+by+targeting+MG53-mediated+cell+membrane+repair.">Treatment of acute lung injury by targeting MG53-mediated cell membrane repair.</a> Nature Communications. 5, 4387 (2014).</li><li>Kim, J. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pegylated-Catalase+Is+Protective+in+Lung+Ischemic+Injury+and+Oxidative+Stress.">Pegylated-Catalase Is Protective in Lung Ischemic Injury and Oxidative Stress.</a> Annals of Thoracic Surgery. , (2020).</li><li>Beal, E. W., 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=D-Ala(2),+D-Leu(5)]+Enkephalin+Improves+Liver+Preservation+During+Normothermic+Ex+Vivo+Perfusion.">D-Ala(2), D-Leu(5)] Enkephalin Improves Liver Preservation During Normothermic Ex Vivo Perfusion.</a> Journal of Surgical Research. 241 (2), 323-335 (2019).</li><li>Akateh, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intrahepatic+Delivery+of+Pegylated+Catalase+Is+Protective+in+a+Rat+Ischemia/Reperfusion+Injury+Model.">Intrahepatic Delivery of Pegylated Catalase Is Protective in a Rat Ischemia/Reperfusion Injury Model.</a> Journal of Surgical Research. 238, 152-163 (2019).</li></ol>

BAW, YGL and JLK are supported through the National Institutes of Health (NIH) grant R01HL143000. BAW is supported through the Department of Defense (DOD) grant W81XWH1810787. SMB is supported through NIH grant R01DK123475. JM is supported through NIH grants AR061385, AR070752, DK106394, and AG056919 as well as by DOD grant W81XWH-18-1-0787.

In this report, we have intervened at several critical steps in a rat lung transplantation protocol to optimize the procedure.&#160;While various cuffing techniques for rat lung transplantation has been reported1,2,3,4,5,6,7,8,9...

For over three decades, researchers have been modifying and improving rat lung transplantation models so that the data generated are more consistent and more reflective of the actual clinical condition. In our laboratory&#39;s time performing this model, we have determined four areas of improvement: cuffing techniques for anastomoses, identification of the recipient&#39;s 4th intercostal space, lung inflation and wound closure during the recipient&#39;s procedure, and the harvesting of samples for analysis.
Cuffing technique modifications for anastomoses can improve the entire transplantation procedure by shortening handling time of the donor lung1,2,3,4,5,6 and making the anastomosis procedure faster and technically easier for the microsurgeon. While it is critical to use the proper sized cuffs to supply the necessary blood and airflow to the transplanted lung, there is limited guidance regarding how one should choose the size of cuffs for the pulmonary vein (PV), pulmonary artery (PA), or bronchus (Br)5,7,8,9. Since the diameters of the PV, PA, and Br are related to the body weight of the donor and recipient rats, we propose that the cuff size be based on body weight. This report provides a size guide for cuffs based on a rat&#39;s body weight (180 g to over 270 g) that serves to optimize blood and air supply to the transplanted lung (Table 1).
While a newer microsurgeon can successfully and easily procure a donor lung during the donor procedure, transplanting the lung during the recipient&#39;s procedure is more complicated and is dependent on the microsurgeon&#39;s experience. Attempts to find the 4th intercostal space to access the recipient&#39;s left lung is one of the more difficult steps that holds some subjectivity and can increase the procedure time. Therefore, we introduce a simple and objective method to assist in the identification of the 4th intercostal space location by using chest measurements and the palpitations of the heart to find the correct area chest wall to dissect4,5,6,10,11,12.
We also propose an improvement to donor lung inflation, which is a potential source of injury to the organ. The donor lung is deflated until reperfusion starts. While suturing the 4th intercostal space, the donor lung is commonly inflated by increasing PEEP from 2 cmH2O to 6 cmH2O. In order to minimize lung injury from overinflation, we propose a technique where three 6-0 nylon sutures are placed around the 4th rib inferior to the 5th rib with simple double knots. When it is time for wound closure, the ends of the three sutures are held with hemostats in both hands, the wound is closed all at once by pulling up on each side, and PEEP is immediately reduced to 2 cmH2O. In this way, the lung is allowed to expand for the shortest time possible10.
At the conclusion of an experiment, the researcher often wants to collect many types of samples for many types of analysis from each transplant. For example, snap frozen tissue, formalin fixed tissue, tissue for wet-to-dry weight ratio to determine pulmonary edema, and bronchoalvelolar lavage (BAL) fluid can all be used to assess how well the transplant went. The traditional method of collecting BAL fluid allows for a mixed pooled sample from both the recipient&#39;s native lobes and the donor&#39;s transplanted lobe13,14,15. To overcome this, we present a method of clamping the hilar areas that can yield more precise insight into the transplanted and native lungs&#39; condition. Additionally, the volume of PBS used to collect BAL fluid from each side of the lungs is important to consider because BAL fluid contains numerous soluble factors such as cytokines and chemokines that are measured by concentration. Normalizing the volume of the fluid instilled to the estimated volume of lung capacity can help with comparison. With four lobes on the right side and one lobe on the left side, each of the rat&#39;s five lobes has a different volume and surface area16. According to a previous study on volume measurement of lung lobes by Backer et al., of the total volume of the whole lung the volume of the right lobes is 63% (4400 mm3) and the left lobe is 37% (2500 mm3). Therefore, we recommend that the volume of PBS used to collect BAL fluid should be calculated as twice the tidal volume (7.2 mL/kg) multiplied by 63% for the right lung and 37% for the left lung. By using this approach, one can better control for variables like body weight and timing10,16.
In all, in this report we will demonstrate a few modifications to the standard experimental model of rat lung transplantation that can make the procedure more efficient and increase the capability of generating more accurate and plentiful data from each experiment.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>12 Gauge angio-catheter</td>
			<td>BD</td>
			<td>382277</td>
			<td></td>
		</tr>
		<tr>
			<td>14 Gauge angio-catheter</td>
			<td>B. Braun</td>
			<td>4251717-02</td>
			<td></td>
		</tr>
		<tr>
			<td>16 Gauge angio-catheter</td>
			<td>B. Braun</td>
			<td>4252586-02</td>
			<td></td>
		</tr>
		<tr>
			<td>18 Gauge angio-catheter</td>
			<td>B. Braun</td>
			<td>4251679-02</td>
			<td></td>
		</tr>
		<tr>
			<td>20 Gauge angio-catheter</td>
			<td>B. Braun</td>
			<td>4252527-02</td>
			<td></td>
		</tr>
		<tr>
			<td>4-0 silk suture</td>
			<td>Surgical Specialties Corp.</td>
			<td>SP116</td>
			<td></td>
		</tr>
		<tr>
			<td>6-0 nylon suture</td>
			<td>AD Surgical</td>
			<td>S-N618R13</td>
			<td></td>
		</tr>
		<tr>
			<td>7-0 nylon suture</td>
			<td>AD Surgical</td>
			<td>S-N718SP13</td>
			<td></td>
		</tr>
		<tr>
			<td>8-0 nylon suture</td>
			<td>AD Surgical</td>
			<td>XXS-N807T6</td>
			<td></td>
		</tr>
		<tr>
			<td>Betadine Spray</td>
			<td>Avrio Health L.P</td>
			<td>UPC 367618160039</td>
			<td></td>
		</tr>
		<tr>
			<td>Clippers</td>
			<td>VWR</td>
			<td>MSPP-023326</td>
			<td></td>
		</tr>
		<tr>
			<td>Castroviejo micro dissecting spring scissors</td>
			<td>Roboz Surgical Instrument Co</td>
			<td>RS-5668</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 - Fine Forceps</td>
			<td>Fine Science Tools</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrocautery</td>
			<td>Macan</td>
			<td>MV-7A</td>
			<td></td>
		</tr>
		<tr>
			<td>Endotracheal intubation kit</td>
			<td>Kent Scientific</td>
			<td>ETI-MSE</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Fine Science Tools</td>
			<td>11027-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Halsted-mosquito hemostat</td>
			<td>Roboz Surgical Instrument Co</td>
			<td>RS-7112</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Fresnius Medical Care</td>
			<td>C504701</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin syringe</td>
			<td>Life Technologies</td>
			<td>B328446</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Piramal Critical Care</td>
			<td>NDC 66794-017-25</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropyl Alcohol Swabs</td>
			<td>BD</td>
			<td>326895</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>Hikma Pharmaceuticals PLC</td>
			<td>NDC 0413-9505-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Dieffenbach Bulldog Clamp</td>
			<td>World Precision Instruments</td>
			<td>WPI14117</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder/Forceps, Curved</td>
			<td>Micrins</td>
			<td>MI1542</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder/Forceps, Straight</td>
			<td>Micrins</td>
			<td>MI1540</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfadex Plus (Organ Preservation Solution)</td>
			<td>XVIVO Perfusion AB</td>
			<td>REF# 19950</td>
			<td></td>
		</tr>
		<tr>
			<td>PhysioSuite</td>
			<td>Kent Scientific</td>
			<td>PS-MSTAT-RT</td>
			<td>Used to check SpO2 and heartbeat</td>
		</tr>
		<tr>
			<td>Retractor</td>
			<td>Roboz Surgical Instrument Co</td>
			<td>RS-6560</td>
			<td></td>
		</tr>
		<tr>
			<td>Saline</td>
			<td>PP Pharmaceuticals LLC</td>
			<td>NDC 63323-186-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>Fine Science Tools</td>
			<td>14090-11</td>
			<td></td>
		</tr>
		<tr>
			<td>SomnoSuite Small Animal Anesthesia System</td>
			<td>Kent Scientific</td>
			<td>SS-MVG-Module</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile&#160; Cotton Gauze Pad</td>
			<td>Fisherbrand</td>
			<td>22-415-469</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Microscope</td>
			<td>Leica</td>
			<td>M500-N w/ OHS</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe 5mL</td>
			<td>BD</td>
			<td>309646</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas-Tubingen Spring Scissors</td>
			<td>Fine Science Tools</td>
			<td>15008-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>Korn Pharmaceuticals Corp</td>
			<td>NDC 59399-110-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Yasargil Clamp</td>
			<td>Aesculap, Inc</td>
			<td>FT351T</td>
			<td>Used to clamp bronchus</td>
		</tr>
		<tr>
			<td>Yasargil Clamp</td>
			<td>Aesculap, Inc</td>
			<td>FT261T</td>
			<td>Used to clamp hilum</td>
		</tr>
		<tr>
			<td>Yasargil Clamp Applicator</td>
			<td>Aesculap, Inc</td>
			<td>FT484T</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

In order to measure pulmonary edema, the wet-to-dry weight ratio was calculated. The donor&#39;s native lobe, the transplanted lobe, and the recipient&#39;s native lobe were collected as described in the protocol and weighed immediately for wet weight, dried at 60 &#176;C for 48 h, and then weighed again for the dry weight. An increased wet-to-dry weight ratio would be indicative of pulmonary edema. Our results indicate that the transplanted lobe did have a significant increase in wet-to-dry ...

Watch this Scientific Journal Video about A Rat Lung Transplantation Model of Warm Ischemia/Reperfusion Injury: Optimizations to Improve Outcomes at JoVE.com

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

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 ...

a-rat-lung-transplantation-model-warm-ischemiareperfusion-injury

Research

JoVE Journal

Medicine

2.8K Views.  The Ohio State University Wexner Medical Center. 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.

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

Normothermic Cardiac Arrest and Cardiopulmonary Resuscitation: A Mouse Model of Ischemia-Reperfusion Injury

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused by whole-body hypoperfusion.5,6 Successful reproduction of whole-body hypoperfusion in rodent models has been fraught with 
difficulty.7-9,9,10 Models which employ focal ischemia have repeatedly demonstrated results which do not translate to the clinical setting, and larger 
animal models which allow for whole body hypoperfusion lack access to the full toolset of genetic manipulation possible in the mouse.11,12 However, in recent 
years a mouse model of cardiac arrest and cardiopulmonary resuscitation has emerged which can be adapted to model AKI.13 This model reliably reproduces 
physiologic, functional, anatomic, and histologic outcomes seen in clinical AKI, is rapidly repeatable, and offers all of the significant advantages of a murine surgical 
model, including access to genetic manipulative techniques, low cost relative to large animals, and ease of use. Our group has developed extensive experience 
with use of this model to assess a number of organ-specific outcomes in AKI.14,15

A powerful model for perioperative and critical care related acute kidney injury is presented. Using whole body hypoperfusion induced by cardiac arrest it is possible to nearly replicate the histologic and functional changes of clinical AKI.

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused ...

In situ Transverse Rectus Abdominis Myocutaneous Flap: A Rat Model of Myocutaneous Ischemia Reperfusion Injury

Free tissue transfer is the gold standard of reconstructive surgery to repair complex defects not amenable to local options or those requiring composite tissue. Ischemia reperfusion injury (IRI) is a known cause of partial free flap failure and has no effective treatment. Establishing a laboratory model of this injury can prove costly both financially as larger mammals are conventionally used and in the expertise required by the technical difficulty of these procedures typically requires employing an experienced microsurgeon. This publication and video demonstrate the effective use of a model of IRI in rats which does not require microsurgical expertise. This procedure is an in situ model of a transverse abdominis myocutaneous (TRAM) flap where atraumatic clamps are utilized to reproduce the ischemia-reperfusion injury associated with this surgery. A laser Doppler Imaging (LDI) scanner is employed to assess flap perfusion and the image processing software, Image J to assess percentage area skin survival as a primary outcome measure of injury.

In situ Transverse Rectus Abdominis Myocutaneous Flap: A Rat Model of Myocutaneous Ischemia Reperfusion Injury

Free tissue transfer is widely employed in reconstructive surgery to restore form and function following oncological resection and trauma. Preconditioning this tissue prior to surgery may improve outcome. This article describes an in situ transverse rectus abdominis myocutaneous flap (TRAM) in rats as a means for testing preconditioning strategies.

Free tissue transfer is the gold standard of reconstructive surgery to repair complex defects not amenable to local options or those requiring composite ...

Ischemia-reperfusion Model of Acute Kidney Injury and Post Injury Fibrosis in Mice

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often unreported mortality rates that may confound analyses. Bilateral renal pedicle clamping is commonly used to induce IR-AKI, but differences between effective clamp pressures and/or renal responses to ischemia between kidneys often lead to more variable results. In addition, shorter clamp times are known to induce more variable tubular injury, and while mice undergoing bilateral injury with longer clamp times develop more consistent tubular injury, they often die within the first 3 days after injury due to severe renal insufficiency. To improve post-injury survival and obtain more consistent and predictable results, we have developed two models of unilateral ischemia-reperfusion injury followed by contralateral nephrectomy. Both surgeries are performed using a dorsal approach, reducing surgical stress resulting from ventral laparotomy, commonly used for mouse IR-AKI surgeries. For induction of moderate injury BALB/c mice undergo unilateral clamping of the renal pedicle for 26 min and also undergo simultaneous contralateral nephrectomy. Using this approach, 50-60% of mice develop moderate AKI 24 hr after injury but 90-100% of mice survive. To induce more severe AKI, BALB/c mice undergo renal pedicle clamping for 30 min followed by contralateral nephrectomy 8 days after injury. This allows functional assessment of renal recovery after injury with 90-100% survival. Early post-injury tubular damage as well as post injury fibrosis are highly consistent using this model.

We describe models of moderate and severe ischemia-reperfusion-induced kidney injury in which the mice undergo unilateral renal pedicle clamping followed by simultaneous or delayed contralateral nephrectomy, respectively. These models consistently give rise to renal dysfunction and post-injury fibrosis, but injury severity and survival are dependent on mouse background, age and surgical equipment.

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often ...

A Murine Model of Myocardial Ischemia-reperfusion Injury through Ligation of the Left Anterior Descending Artery

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms at work during MI and developing effective therapeutic approaches requires methodology to reproducibly simulate the clinical incidence and reflect the pathophysiological changes associated with MI. Here, we describe a surgical method to induce MI in mouse models that can be used for short-term ischemia-reperfusion (I/R) injury as well as permanent ligation. The major advantage of this method is to facilitate location of the left anterior descending artery (LAD) to allow for accurate ligation of this artery to induce ischemia in the left ventricle of the mouse heart. Accurate positioning of the ligature on the LAD increases reproducibility of infarct size and thus produces more reliable results. Greater precision in placement of the ligature will improve the standard surgical approaches to simulate MI in mice, thus reducing the number of experimental animals necessary for statistically relevant studies and improving our understanding of the mechanisms producing cardiac dysfunction following MI. This mouse model of MI is also useful for the preclinical testing of treatments targeting myocardial damage following MI.

We introduce a surgical method to induce experimental ischemia/reperfusion (I/R) injury to simulate myocardial infarction (MI) in mouse models that allows for more clarity in positioning of the ligation on the left anterior descending artery (LAD) to increase the reproducibility of MI experiments in mice.

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms ...

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 ...

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 ...

Murine Model of Intestinal Ischemia-reperfusion Injury

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, hypotension, necrotizing enterocolitis, bowel transplantation, trauma and chronic inflammation. Intestinal ischemia-reperfusion (IR) injury is a consequence of acute mesenteric ischemia, caused by inadequate blood flow through the mesenteric vessels, resulting in intestinal damage. Reperfusion following ischemia can further exacerbate damage of the intestine. The mechanisms of IR injury are complex and poorly understood. Therefore, experimental small animal models are critical for understanding the pathophysiology of IR injury and the development of novel therapies.
Here we describe a mouse model of acute intestinal IR injury that provides reproducible injury of the small intestine without mortality. This is achieved by inducing ischemia in the region of the distal ileum by temporally occluding the peripheral and terminal collateral branches of the superior mesenteric artery for 60 min using microvascular clips. Reperfusion for 1 hr, or 2 hr after injury results in reproducible injury of the intestine examined by histological analysis. Proper position of the microvascular clips is critical for the procedure. Therefore the video clip provides a detailed visual step-by-step description of this technique. This model of intestinal IR injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Here we describe the detailed procedure of intestinal ischemia-reperfusion in mice which results in reproducible injury without mortality to encourage the standardization of this technique across the field. This model of intestinal ischemia-reperfusion injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, ...

A Mouse Model of Retinal Ischemia-Reperfusion Injury Through Elevation of Intraocular Pressure

Retinal ischemia-reperfusion (I/R) is a pathophysiological process contributing to cellular damage in multiple ocular conditions, including glaucoma, diabetic retinopathy, and retinal vascular occlusions. Rodent models of I/R injury are providing significant insights into mechanisms and treatment strategies for human I/R injury, especially with regard to neurodegenerative damage in the retinal neurovascular unit. Presented here is a protocol for inducing retinal I/R injury in mice through elevation of intraocular pressure (IOP). In this protocol, the ocular anterior chamber is cannulated with a needle, through which flows the drip of an elevated saline reservoir. Using this drip to raise IOP above systolic arterial blood pressure, a practitioner temporarily halts inner retinal blood flow (ischemia). When circulation is reinstated (reperfusion) by removal of the cannula, severe cellular damage ensues, resulting ultimately in retinal neurodegeneration. Recent studies demonstrate inflammation, vascular permeability, and capillary degeneration as additional elements of this model. Compared to alternative retinal I/R methodologies, such as retinal arterial ligation, retinal I/R injury by elevated IOP offers advantages in its&#160;anatomical specificity, experimental tractability, and technical accessibility, presenting itself as a valuable tool for examining neuronal pathogenesis and therapy in the retinal neurovascular unit.

This article describes a procedure for inducing retinal ischemia-reperfusion injury by elevated intraocular pressure in mice. Retinal ischemia-reperfusion injury by elevated intraocular pressure serves to model human pathologies characterized by compromised oxygen and nutrient delivery in the retina, enabling researchers to examine potential cellular mechanisms and treatments for human diseases of the retinal neurovascular unit.

Retinal ischemia-reperfusion (I/R) is a pathophysiological process contributing to cellular damage in multiple ocular conditions, including glaucoma, ...

A Pre-clinical Rat Model for the Study of Ischemia-reperfusion Injury in Reconstructive Microsurgery

Ischemia-reperfusion injury is the main cause of flap failure in reconstructive microsurgery. The rat is the preferred preclinical animal model in many areas of biomedical research due to its cost-effectiveness and its translation to humans. This protocol describes a method to create a preclinical free skin flap model in rats with ischemia-reperfusion injury. The described 3 cm x 6 cm rat free skin flap model is easily obtained after the placement of several vascular ligatures and the section of the vascular pedicle. Then, 8 h after the ischemic insult and completion of the microsurgical anastomosis, the free skin flap develops the tissue damage. These ischemia-reperfusion injury-related damages can be studied in this model, making it a suitable model for evaluating therapeutic agents to address this pathophysiological process. Furthermore, two main monitoring techniques are described in the protocol for the assessment of this animal model: transit-time ultrasound technology and laser speckle contrast analysis.

Here, we describe a pre-clinical animal model for studying the pathophysiology of ischemia-reperfusion injury in reconstructive microsurgery. This free skin flap model based on the superficial caudal epigastric vessels in the rat may also allow for the evaluation of different therapies and compounds to counteract ischemia-reperfusion injury-related damage.

Ischemia-reperfusion injury is the main cause of flap failure in reconstructive microsurgery. The rat is the preferred preclinical animal model in many ...

Delayed Intramyocardial Delivery of Stem Cells after Ischemia Reperfusion Injury in a Murine Model

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, cardiac stem cell therapy is studied by delivering SCs concurrently with the induction of myocardial injury. However, this approach presents two significant limitations: the early hostile pro-inflammatory ischemic environment may affect the survival of transplanted SCs, and it does not represent the subacute infarction scenario where SCs will likely be used. Here we describe a two-part series of surgical procedures for the induction of ischemia-reperfusion injury and delivery of mesenchymal stem cells (MSCs). This method of stem cell administration may allow for the longer viability and retention around damaged tissue by circumventing the initial immune response. A model of ischemia reperfusion injury was induced in mice accompanied by the delivery of mesenchymal stem cells (3.0 x 105), stably expressing the reporter gene firefly luciferase under the constitutively expressed CMV promoter, intramyocardially 7 days later. The animals were imaged via ultrasound and bioluminescent imaging for confirmation of injury and injection of cells, respectively. Importantly, there was no added complication rate when performing this two-procedure approach for SC delivery. This method of stem cell administration, collectively with the utilization of state-of-the-art reporter genes, may allow for the in vivo study of viability and retention of transplanted SCs in a situation of chronic ischemia commonly seen clinically, while also circumventing the initial pro-inflammatory response. In summary, we established a protocol for the delayed delivery of stem cells into the myocardium, which can be used as a potential new approach in promoting regeneration of the damaged tissue.

Stems cells are continuously investigated as potential treatments for individuals with myocardial damage, however, their decreased viability and retention within injured tissue can impact their long-term efficacy. In this manuscript we describe an alternative method for stem cell delivery in a murine model of ischemia reperfusion injury.

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, ...