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>

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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><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&amp;nbsp; 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 ...

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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 ...

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2.8K Views.  The Ohio State University Wexner Medical Center. Dit protocol identificeert vier belangrijke gebieden om de methodologie te helpen verbeteren en betrouwbare longtransplantatie te hebben. De eerste is een reproduceerbare manchettechniek. Ten tweede is het nauwkeurig en reproduceerbaar identificeren van de vierde intercostale ruimte, een grondige sluiting van de wond en vervolgens hoe te herstellen en monsters te verkrijgen voor analyse.Deze technieken helpen bij het ontwikkelen van een betrouwbare methode van longtransplantatie om te ...

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

Development of Obliterative Bronchiolitis in a Murine Model of Orthotopic Lung Transplantation

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and standardized not only for the study of allo-rejection but also between syngeneic strains for examining mechanisms of ischemia-reperfusion injury after lung transplantation. Although the application of the rat and other large animal model 2 contributed significantly to the elucidation of these studies, the scope of those investigations is limited by the scarcity of knockout and transgenic rats. Due to no effective therapies for obliterative bronchiolitis, the leading cause of death in lung transplant patients, there has been an intensive search for pre-clinical models that replicate obliterative bronchiolitis. The tracheal allograft model is the most widely used and may reproduce some of the histopathologic features of obliterative bronchiolitis 3. However, the lack of an intact vasculature with no connection to the recipient's conducting airways, and incomplete pathologic features of obliterative bronchiolitis limit the utility of this model 4. Unlike transplantation of other solid organs, vascularized mouse lung transplants have only recently been reported by Okazaki and colleagues for the first time in 2007 5. Applying the basic principles of the rat lung transplant, our lab initiated the obliterative bronchiolitis model using minor histoincompatible antigen murine orthotopic single-left lung transplants which allows the further study of obliterative bronchiolitis immunopathogenesis6.

Obliterative bronchiolitis is the key impediment to the long-term survival of lung transplant recipients and the lack of a robust preclinical model precludes examining obliterative bronchiolitis immunopathogenesis. Unlike other solid organ transplants, vascularized mouse lung transplantation has only recently been developed. Here we show our independently developed obliterative bronchiolitis model after murine orthotopic single-lung transplantation.

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and ...

Surgical Procedures for a Rat Model of Partial Orthotopic Liver Transplantation with Hepatic Arterial Reconstruction

Orthotopic liver transplantation (OLT) in rats using a whole or partial graft is an indispensable experimental model for transplantation research, such as studies on graft preservation and ischemia-reperfusion injury 1,2, immunological responses 3,4, hemodynamics 5,6, and small-for-size syndrome 7. The rat OLT is among the most difficult animal models in experimental surgery and demands advanced microsurgical skills that take a long time to learn. Consequently, the use of this model has been limited. Since the reliability and reproducibility of results are key components of the experiments in which such complex animal models are used, it is essential for surgeons who are involved in rat OLT to be trained in well-standardized and sophisticated procedures for this model.
While various techniques and modifications of OLT in rats have been reported 8 since the first model was described by Lee et al. 9 in 1973, the elimination of the hepatic arterial reconstruction 10 and the introduction of the cuff anastomosis technique by Kamada et al. 11 were a major advancement in this model, because they simplified the reconstruction procedures to a great degree. In the model by Kamada et al., the hepatic rearterialization was also eliminated. Since rats could survive without hepatic arterial flow after liver transplantation, there was considerable controversy over the value of hepatic arterialization. However, the physiological superiority of the arterialized model has been increasingly acknowledged, especially in terms of preserving the bile duct system 8,12 and the liver integrity 8,13,14.
In this article, we present detailed surgical procedures for a rat model of OLT with hepatic arterial reconstruction using a 50% partial graft after ex vivo liver resection. The reconstruction procedures for each vessel and the bile duct are performed by the following methods: a 7-0 polypropylene continuous suture for the supra- and infrahepatic vena cava; a cuff technique for the portal vein; and a stent technique for the hepatic artery and the bile duct.

Orthotopic liver transplantation in rats is an indispensable experimental model for biomedical research. Here we present our surgical procedures for orthotopic rat liver transplantation with hepatic arterial reconstruction using a 50% partial graft.

Orthotopic liver transplantation (OLT) in rats using a whole or partial graft is an indispensable experimental model for transplantation research, such as ...

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 ...

Method of Direct Segmental Intra-hepatic Delivery Using a Rat Liver Hilar Clamp Model

Major hepatic surgery with inflow occlusion, and liver transplantation, necessitate a period of warm ischemia, and a period of reperfusion leading to ischemia/reperfusion (I/R) injury with myriad negative consequences. Potential I/R injury in marginal organs destined for liver transplantation contributes to the current donor shortage secondary to a decreased organ utilization rate. A significant need exists to explore hepatic I/R injury in order to mediate its impact on graft function in transplantation. Rat liver hilar clamp models are used to investigate the impact of different molecules on hepatic I/R injury. Depending on the model, these molecules have been delivered using inhalation, epidural infusion, intraperitoneal injection, intravenous administration or injection into the peripheral superior mesenteric vein. A rat liver hilar clamp model has been developed for use in studying the impact of pharmacologic molecules in ameliorating I/R injury. The described model for rat liver hilar clamp includes direct cannulation of the portal supply to the ischemic hepatic segment via a side branch of the portal vein, allowing for direct segmental hepatic delivery. Our approach is to induce ischemia in the left lateral and median lobes for 60 min, during which time the substance under study is infused. In this case, pegylated-superoxide dismutase (PEG-SOD), a free radical scavenger, is infused directly into the ischemic segment. This series of experiments demonstrates that infusion of PEG-SOD is protective against hepatic I/R injury. Advantages of this approach include direct injection of the molecule into the ischemic segment with consequent decrease in volume of distribution and reduction in systemic side effects.

A unique rat liver hilar clamp model was developed for studying the impact of pharmacologic molecules in ameliorating ischemia-reperfusion injury. This model includes direct cannulation of the portal supply to the ischemic liver segment via a branch of the portal vein, allowing for direct hepatic delivery.

Major hepatic surgery with inflow occlusion, and liver transplantation, necessitate a period of warm ischemia, and a period of reperfusion leading to ...

Reduced Complications after Arterial Reconnection in a Rat Model of Orthotopic Liver Transplantation

The rat orthotopic liver transplantation (OLT) model is a powerful tool to study acute and chronic rejection. However, it is not a complete representation of human liver transplantation due to the absence of arterial reconnection. Described here is a modified transplantation procedure that includes the incorporation of hepatic artery (HA) reconnection, leading to a marked improvement in transplant outcomes. With a mean anhepatic time of 12 min and 14 s, HA reconnection results in improved perfusion of the transplanted liver and an increase in long-term recipient survival from 37.5% to 88.2%. This protocol includes the use of 3D-printed cuffs and holders to connect the portal vein and infrahepatic inferior vena cava. It can be implemented for studying multiple aspects of liver transplantation, from immune response and infection to technical aspects of the procedure. By incorporating a simple and practical method for arterial reconnection using a microvascular technique, this modified rat OLT protocol closely mimics aspects of human liver transplantation and will serve as a valuable and clinically relevant research model.

The goal of this study is to modify the rat orthotopic liver transplant model to better represent human liver transplantation and improve recipient survival. The presented method reestablishes hepatic arterial inflow by connecting the donor liver&#39;s common hepatic artery to the recipient liver&#39;s proper hepatic artery.

The rat orthotopic liver transplantation (OLT) model is a powerful tool to study acute and chronic rejection. However, it is not a complete representation ...

Organ Ischemia-Reperfusion Injury by Simulating Hemodynamic Changes in Rat Liver Transplant Model

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, including ischemia-reperfusion injury (IRI) of extrahepatic organs. This model requires numerous experiments and devices. The duration of anhepatic phase is closely related to the time to develop IRI after transplantation. In this experiment, we used hemodynamic changes to induce extrahepatic organ damage in rats and determined the maximum tolerance time. The time until the most severe organ injury varied for different organs. This method can easily be replicated and can also be used to study IRI of the extrahepatic organs after liver transplantation.

This paper provides a detailed description of how to build an animal model of the anhepatic phase (liver ischemia) in rats to facilitate basic research into ischemia-reperfusion injury after liver transplantation.

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, ...

Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation

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

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

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

Establishment of an Ex Vivo Lung Perfusion Rat Model for Translational Insights in Lung Transplantation

Since the establishment of lung transplantation as a therapeutic strategy for advanced lung diseases, the scientific community is faced with the problem of a low number of lungs considered viable for the donation process. In recent decades, however, this scenario has been positively changed, given the development of ex vivo lung perfusion (EVLP) as a strategy for evaluating and reconditioning marginal lungs. The establishment of EVLP in large transplant centers has favored an increase in the number of lung transplants, both by increasing the diagnostic accuracy of lung function and by constituting an effective platform for the reconditioning of lung grafts. In this context, faced with ethical and logistical issues, as well as in the study of immunological factors associated with lung transplantation, the development of rodent EVLP models has become important, given their reliability, the possibility of genetic manipulation, and lower costs. This paper describes a protocol for establishing a rat EVLP model and shows the inflammatory profile associated with the perfused lungs. This will help propagate knowledge about the rat EVLP model, promoting our understanding of the biological responses associated with that revolutionary technique.

Establishment of an Ex Vivo Lung Perfusion Rat Model for Translational Insights in Lung Transplantation

Here, we describe the necessary steps for establishing a rat EVLP model and show the inflammatory profile associated with the perfused lungs. The aim is to propagate knowledge and experiences about the rat EVLP model, enabling the integral understanding of the biological responses associated with that revolutionary technique.

Since the establishment of lung transplantation as a therapeutic strategy for advanced lung diseases, the scientific community is faced with the problem ...

Mouse Model for Studying Lung Ischemia-Reperfusion Injury

Murine Left Pulmonary Hilar Clamp Model of Lung Ischemia Reperfusion Injury

Ischemia reperfusion injury (IRI) during lung transplantation is a major risk factor for post-transplant complications, including primary graft dysfunction, acute and chronic rejection, and mortality. Efforts to study the underpinnings of IRI led to the development of a reliable and reproducible mouse model of left lung hilar clamping. This model involves a surgical procedure performed in an anesthetized and intubated mouse. A left thoracotomy is performed, followed by careful lung mobilization and dissection of the left pulmonary hilum. The hilar clamp involves reversible suture ligation of the pulmonary hilum with a slipknot, which stops the arterial inflow, venous outflow, and airflow through the left mainstem bronchus. Reperfusion is initiated by careful removal of the suture. Our laboratory uses 30 min of ischemia and 1 h of reperfusion for the experimental model in the current investigations. However, these time periods can be modified depending on the specific experimental question. Immediately prior to sacrifice, arterial blood gas can be obtained from the left ventricle after a 4 min period of right hilar clamping to ensure that the PaO2 values obtained are attributed to the injured left lung alone. We also describe a method to measure cell extravasation with flow cytometry, which involves intravenous injection of a fluorochrome-labeled antibody specific for the cell(s) to be studied prior to sacrifice. The left lung can then be harvested for flow cytometry, frozen or fixed, paraffin-embedded immunohistochemistry, and quantitative polymerase chain reaction. This hilar clamp technique allows for&#160;detailed study of the cellular and molecular mechanisms underlying IRI. Representative results reveal decreased left lung oxygenation and histologic evidence of lung injury following hilar clamping. This technique can be readily learned and reproduced by personnel with and without microsurgical experience, leading to reliable and consistent results and serving as a widely adoptable model for studying lung IRI.

Author Spotlight: Insights into the Effect of Ischemia Reperfusion on Lung Transplantation

The protocol outlines a feasible, reliable, and reproducible method of left pulmonary hilar clamping that can be used to study lung ischemia-reperfusion injury in mouse models.

Ischemia reperfusion injury (IRI) during lung transplantation is a major risk factor for post-transplant complications, including primary graft ...

Evaluation of Apoptosis and Lung Injury in DCD Donor Lungs Preserved by EVLP

Establishment of a Novel Ex Vivo Lung Perfusion System for Rat Lungs After Circulatory Death

The increasing scarcity of donor lungs for transplantation has spurred significant interest in the utilization of organs procured from donors after cardiac death (DCD). However, a critical challenge remains in maximizing the number of viable lungs that meet stringent criteria for clinical use. Ex vivo lung perfusion (EVLP) emerges as a promising technology to evaluate and potentially improve the function of donated lungs. This paper describes the development of a cost-effective and reproducible small animal experimental platform specifically designed for EVLP research. This platform offers a high degree of customization, enabling researchers to tailor experimental parameters to address specific research questions within the field of EVLP. The team established a rat model to simulate DCD and implemented a two-stage preservation approach. Lungs were subjected to static cold storage using a perfusate solution at 4 &#176;C for 4 h, followed by normothermic EVLP for another 4 h. Researchers evaluated various lung function parameters, including pulmonary vascular resistance, pulmonary dynamic compliance, blood gas measurements of the perfusate, and apoptotic cell death in lung tissues. The study demonstrates that EVLP could significantly improve the functional performance of DCD lungs compared to simple cold storage at 4 &#176;C, suggesting that EVLP has the potential to mitigate ischemia-reperfusion injury and enhance graft function following circulatory death, thereby increasing the pool of transplantable lungs.

The objective is to develop a new ex vivo lung perfusion (EVLP) model in rats that can sustain consistent lung function for 4 h throughout EVLP and furnish the industry with implementation specifics that will offer a sturdy method for applying EVLP to rodent models.

The increasing scarcity of donor lungs for transplantation has spurred significant interest in the utilization of organs procured from donors after ...

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

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