Name: Orthotopic Left Lung Transplantation in Rats (Video) | JoVE Journal
Uploaded: 2025-07-03T12:55:00.000+00:00
Duration: 8 min 22 s
Description: Learn an improved orthotopic left lung transplantation technique in rats for studying chronic rejection and post-transplant complications. This standardized method offers higher reproducibility and preserved lung function for translational research.

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5 minutes

All animal experiments have been approved by the Animal Ethics Committee of Guangzhou Lai&#39;an Technology Co., Ltd. (Approval No. G2024129). The experiment was conducted on male Lewis rats aged 6-8 weeks and weighing 250-300 g. Rats were raised under a 12 h light/dark cycle (light: 25 lux) and had free access to food and water.
1. Prepare cuffs (Figure 1)
<ol>
	<li>Prepare 14 G and 16 G intravenous catheters and blades (Figure 1A).</li>
	<li>Trim intravenous catheters into a cuff with a tail using a sterile blade. Divide each cuff into a body (2-3 mm length) and a tail (2-3 mm length) (Figure 1B).</li>
	<li>Create superficial scratches on the cuff body to increase friction for suture fixation. 
	NOTE: Keep 16 G cuffs for pulmonary artery (PA) and pulmonary vein (PV) anastomoses, and reserve 14 G cuffs for bronchi (Br) connections (Figure 1C). The catheters and blades are disposable sterile instruments. The surgical platform on which the cannula was cut was sterilized with UV irradiation for 30 min and three times with 75% alcohol.</li>
</ol>
2. Donor lung harvest (Figure 2)
<ol>
	<li>Anesthetize the donor rat via intraperitoneal injection of pentobarbital (80 mg/kg) and administer subcutaneous injection&#160;of meloxicam (5 mg/kg) as preoperative analgesics. Position the rat supine on the operating table with the head elevated and tail lowered and hyperextend the neck.</li>
	<li>Retract the tongue outward and upward, then position a surgical lamp anterior to the neck as a localized light source to achieve clear visualization of the glottal opening (Figure 2A).</li>
	<li>Insert a 14 G intravenous cannula into the airway through the glottis.</li>
	<li>Set the ventilator to pressure-controlled mode, input the weight parameters, adjust the pressure to 15 cmH2O, and connect the ventilator.</li>
	<li>Check whether the rise and fall of the chest is consistent with the ventilator frequency to ensure that the intravenous catheter is in the airway.</li>
	<li>Pinch the rat&#39;s toes to confirm that the rat has entered deep anesthesia. Fix the limbs and head, disinfect the chest and abdomen three times with iodophor and 75% alcohol.&#160;Lift the abdominal skin with tweezers, and use scissors to cut the skin from the abdomen, chest, and front of the neck.</li>
	<li>Make a midline incision, cut the abdominal wall, and inject heparin at a dose of 1,000 IU/kg through the exposed peritoneal cavity vein for 3 min to ensure systemic heparinization.</li>
	<li>Cut the diaphragm and cut the chest cavity from the middle of the sternum. Fix the chest wall on both sides with hemostats. Remove the thymus and fully expose the anatomical structure of the chest cavity organs.</li>
	<li>Cut the superior vena cava, inferior vena cava, and the left and right auriculae in turn. Inject cold saline with a 20 mL syringe into the root of the PA slowly (~1 min) for low-pressure perfusion until the donor lung turns white completely (Figure 2B).</li>
	<li>Clamp the Br of the donor lung with hemostatic forceps. Lift the Br and cut off the tissues connected to the donor lung. Take out the donor heart and lung en bloc and soak them in precooled saline (Figure 2C). 
	NOTE: Clamping Br while the donor lung is in the inspiratory state can ensure that the alveoli of the donor lung will not collapse.</li>
</ol>
3. Donor lung modification (Figure 3)
<ol>
	<li>Position the heart-lung block on ice under the microscope. Grip the trachea with a hemostat and secure it in plasticine.</li>
	<li>Cover the left and right lungs with wet sterile lens paper, separate the left and right lungs as much as possible, and fully expose the anatomical structure of the pulmonary hilum.</li>
	<li>Separate the PA, Br, and PV under the microscope with forceps and ligate the Br close to the lung with 6-0 surgical sutures (Figure 3A-C).</li>
	<li>Extract the PA, Br, and PV from the cuff made of 16 G, 14 G, and 16 G venous indentation cuff, respectively. Fix the tube walls to the cuff using 8-0 surgical sutures (Figure 3D-F).</li>
	<li>Preserve the modified donor lungs in saline, repositioned on ice, and await implantation.</li>
</ol>
4. Donor lung implantation (Figure 4)
<ol>
	<li>Anesthetize the recipient rats with an intraperitoneal injection of pentobarbital (70 mg/kg). Apply the rat eyeball with eye ointment and intubate using the same method as for steps 2.1-2.3.</li>
	<li>Depilate the recipient&#39;s left thoracodorsal region, position the recipient in the right lateral decubitus posture on a thermostatic operating table, and disinfect the surgical field three times with iodophor and 75% alcohol. Administer lidocaine as local anesthetic before making the incision. 
	NOTE: Due to the narrow operating space and the entire procedure is performed in a room free of special pathogens, so the animal is not draped, but the surgical incision is covered with sterile gauze as much as possible.</li>
	<li>Make an incision through the skin and chest wall at the point where the apical impulse is most prominent (in the fourth intercostal space) to enter the thoracic cavity.</li>
	<li>Use an eyelid retractor to open the thoracic cavity. Gently push the left lung aside with a moist cotton swab to expose the inferior pulmonary ligament, and then sharply transect the ligament.</li>
	<li>Grasp the left lung with forceps and retract it outside the thoracic cavity. Secure the hilum of the lung with a hemostat and fix it to the ventral thoracic incision. Use modeling clay as a surgical tip to further stabilize the hemostat on the recipient&#39;s side (Figure 4A).</li>
	<li>Remove the tissue around the left lung, dissect the pulmonary hilum, and separate the PA, PV, and Br.</li>
	<li>Clamp the proximal ends of the PA, Br, and PV using vascular clamps (Figure 4B).</li>
	<li>Use 8-0 surgical suture to pre-tie the recipient&#39;s PA, Br, and PV, ensuring rapid anastomosis during donor lung implantation (Figure 4C).</li>
	<li>Trim a platform on the clamped recipient&#39;s left lung to facilitate the placement of the donor lung. Make a small incision at the distal end of the recipient&#39;s PA, Br, and PV (Figure 4D).</li>
	<li>Rinse the PA and PV with heparinized saline to prevent thrombus formation (Figure 4E).</li>
	<li>Take the donor lung out of the ice and place it on the recipient&#39;s left lung platform. Lift one side of the cut opening with forceps, then implant the cuffs of the PA, Br, and PV into the recipient sequentially and ligate them (Figure 4F,G). 
	NOTE: The sequence of anastomosis should be performed as follows: first anastomose the PA, followed by the Br, and finally, the PV.</li>
	<li>Open the micro hemostatic clip and observe the donor lung gradually changing from white to red. Inspect and ensure no bleeding at the anastomotic site (Figure 4H).</li>
	<li>Remove the recipient&#39;s left lung, and return the donor lung to the chest cavity (Figure 4I,J).</li>
	<li>Appropriately increase the ventilator pressure to ensure adequate expansion of the alveoli in the donor lung. 
	NOTE: In pressure-controlled ventilation mode, the pressure should not exceed 25 cmH2O. During the procedure, ventilator parameters are adjusted based on the recipient&#39;s condition. If the recipient exhibits gasping respirations, it suggests the presence of hypoxia, and the ventilator parameters should be promptly adjusted accordingly.</li>
	<li>Wipe the chest cavity dry with a sterile cotton ball, then close it layer by layer (Figure 4K).</li>
	<li>Wait for the recipient to gradually recover spontaneous breathing, then remove the ventilator and observe. When the spontaneous breathing rate reaches about 100 times per minute, remove the endotracheal tube (Figure 4L). 
	NOTE: Signs indicating that the recipient is ready for ventilator removal upon recovering spontaneous breathing include whisker movement, limb movement, and an irregular breathing curve on the ventilator display.</li>
	<li>House the recipients in individual cages with free access to food and water, and maintain the temperature at ~22 &#176;C.</li>
	<li>Administer subcutaneous injections of meloxicam (5 mg/kg) to the recipients for 3 consecutive days to relieve pain.</li>
</ol>

<ol><li>Perch, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+International+Thoracic+Organ+Transplant+Registry+of+the+International+Society+for+Heart+and+Lung+Transplantation%3A+Thirty-ninth+adult+lung+transplantation+report-2022%3B+focus+on+lung+transplant+recipients+with+chronic+obstructive+pulmonary+disease%5BTitle%5D)+AND+%22J+Heart+Lung+Transplant%22%5BJournal%5D)">The International Thoracic Organ Transplant Registry of the International Society for Heart and Lung Transplantation: Thirty-ninth adult lung transplantation report-2022; focus on lung transplant recipients with chronic obstructive pulmonary disease.</a> J Heart Lung Transplant. 41 (10), 1335-1347 (2022).</li>
<li>Kulkarni, H. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bronchiolitis+obliterans+syndrome-free+survival+after+lung+transplantation%3A+An+International+Society+for+Heart+and+Lung+Transplantation+Thoracic+Transplant+Registry+analysis%5BTitle%5D)+AND+%22J+Heart+Lung+Transplant%22%5BJournal%5D)">Bronchiolitis obliterans syndrome-free survival after lung transplantation: An International Society for Heart and Lung Transplantation Thoracic Transplant Registry analysis.</a> J Heart Lung Transplant. 38 (1), 5-16 (2019).</li>
<li>Bos, S., Vos, R., Van Raemdonck, D. E., Verleden, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Survival+in+adult+lung+transplantation%3A+where+are+we+in+2020%5BTitle%5D)+AND+%22Curr+Opin+Organ+Transplant%22%5BJournal%5D)">Survival in adult lung transplantation: where are we in 2020.</a> Curr Opin Organ Transplant. 25 (3), 268-273 (2020).</li>
<li>Hu, C. X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lung+transplantation+in+China+between+2015+and+2018%5BTitle%5D)+AND+%22Chin+Med+J+%28Engl%29%22%5BJournal%5D)">Lung transplantation in China between 2015 and 2018.</a> Chin Med J (Engl). 132 (23), 2783-2789 (2019).</li>
<li>Khush, K. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+International+Thoracic+Organ+Transplant+Registry+of+the+International+Society+for+Heart+and+Lung+Transplantation%3A+Thirty-sixth+adult+heart+transplantation+report+-+2019%3B+focus+theme%3A+Donor+and+recipient+size+match%5BTitle%5D)+AND+%22J+Heart+Lung+Transplant%22%5BJournal%5D)">The International Thoracic Organ Transplant Registry of the International Society for Heart and Lung Transplantation: Thirty-sixth adult heart transplantation report - 2019; focus theme: Donor and recipient size match.</a> J Heart Lung Transplant. 38 (10), 1056-1066 (2019).</li>
<li>Goto, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Simplified+rat+lung+transplantation+using+a+new+cuff+technique%5BTitle%5D)+AND+%22Ann+Thorac+Surg%22%5BJournal%5D)">Simplified rat lung transplantation using a new cuff technique.</a> Ann Thorac Surg. 93 (6), 2078-2080 (2012).</li>
<li>Li, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+modified+orthotopic+left+lung+transplantation+model+in+rats%5BTitle%5D)+AND+%22Heliyon%22%5BJournal%5D)">A modified orthotopic left lung transplantation model in rats.</a> Heliyon. 10 (10), e30728(2024).</li>
<li>Tian, D., Shiiya, H., Sato, M., Nakajima, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rat+lung+transplantation+model%3A+modifications+of+the+cuff+technique%5BTitle%5D)+AND+%22Ann+Transl+Med%22%5BJournal%5D)">Rat lung transplantation model: modifications of the cuff technique.</a> Ann Transl Med. 8 (6), 407(2020).</li>
<li>Lee, Y. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+rat+lung+transplantation+model+of+warm+ischemia%2Freperfusion+injury%3A+Optimizations+to+improve+outcomes%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">A rat lung transplantation model of warm ischemia/reperfusion injury: Optimizations to improve outcomes.</a> J Vis Exp. (176), e62455(2021).</li>
<li>Rajab, T. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Techniques+for+lung+transplantation+in+the+rat%5BTitle%5D)+AND+%22Exp+Lung+Res%22%5BJournal%5D)">Techniques for lung transplantation in the rat.</a> Exp Lung Res. 45 (9-10), 267-274 (2019).</li>
<li>Zhao, J., Tian, D., Yang, X., Liu, M., Chen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Anterior+hilum+anastomosis+versus+posterior+hilum+anastomosis+in+a+mouse+lung+transplantation+model%5BTitle%5D)+AND+%22JTCVS+Tech%22%5BJournal%5D)">Anterior hilum anastomosis versus posterior hilum anastomosis in a mouse lung transplantation model.</a> JTCVS Tech. 14, 159-165 (2022).</li>
<li>Guo, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Improvements+of+surgical+techniques+in+a+rat+model+of+an+orthotopic+single+lung+transplant%5BTitle%5D)+AND+%22Eur+J+Med+Res%22%5BJournal%5D)">Improvements of surgical techniques in a rat model of an orthotopic single lung transplant.</a> Eur J Med Res. 18 (1), 1(2013).</li>
<li>Zhai, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Simplified+rat+lung+transplantation+by+using+a+modified+cuff+technique%5BTitle%5D)+AND+%22J+Invest+Surg%22%5BJournal%5D)">Simplified rat lung transplantation by using a modified cuff technique.</a> J Invest Surg. 21 (1), 33-37 (2008).</li>
<li>Gao, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Improvement+of+surgical+techniques+for+orthotopic+single+lung+transplantation+in+rats%5BTitle%5D)+AND+%22Ann+Transl+Med%22%5BJournal%5D)">Improvement of surgical techniques for orthotopic single lung transplantation in rats.</a> Ann Transl Med. 10 (12), 673(2022).</li>
<li>Krupnick, A. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Orthotopic+mouse+lung+transplantation+as+experimental+methodology+to+study+transplant+and+tumor+biology%5BTitle%5D)+AND+%22Nat+Protoc%22%5BJournal%5D)">Orthotopic mouse lung transplantation as experimental methodology to study transplant and tumor biology.</a> Nat Protoc. 4 (1), 86-93 (2009).</li>
<li>Kubisa, B., Schmid, R. A., Grodzki, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Model+of+single+left+rat+lung+transplantation.+Relation+between+surgical+experience+and+outcomes%5BTitle%5D)+AND+%22Rocz+Akad+Med+Bialymst%22%5BJournal%5D)">Model of single left rat lung transplantation. Relation between surgical experience and outcomes.</a> Rocz Akad Med Bialymst. 48, 70-73 (2003).</li>
</ol>

This work was supported by the National Natural Science Foundation of China (Grant No. 82470103), the Wu Jieping Medical Foundation Special Research Grant (No. 320.6750.2025-01-1), the China Medical Education Association (Grant No. ZJWYH-2023-YIZHI-003), the Clinical Epidemiology Research Project of the State Key Laboratory of Respiratory Diseases (Grant No. SKLRD-L-202504), the Guangzhou Clinical Characteristic Technology Project (Grant No. 2023C-TS10), and the Major Clinical Research Project under the Research Capacity Enhancement Program of Guangzhou Medical University (Grant No. GMUCR2024-01007).

The rat lung transplant model is a commonly used model in the field of basic research on lung transplantation. It can well simulate the pathophysiological process of human lung transplantation. Rats also have the advantages of fast reproduction and low cost. Mizuta et al.6 first used cuff technology in 1989 to greatly reduce the difficulty of this model. However, due to the complexity of the operation, many researchers are still hesitant. 
In the process of cuff preparation, most researchers have made cuffs with similar shapes. Wu et al.7 made a depression in the middle of the cuff to prevent the knot from slipping during the anastomosis process, but they needed two people to perform the anastomosis, which would undoubtedly increase the manpower and material costs. Some researchers also modified the tail of the cuff to make it smoother8, but they did not remove the tail of the cuff before implanting the cuff into the recipient, which made it easy for the cuff to cause blood vessels and airways to twist or fold in the recipient. 
In terms of cuff size selection, previous studies have suggested that an 18 G cuff should be used for the PA when the recipient's weight is less than 270 g, and a 16 G cuff should be used when the recipient's weight exceeds 270 g9. Some researchers also uniformly use an 18 G cuff7,10. In terms of donor lung modification, Tian et al.8 placed the donor lung in a self-designed culture dish to fix the pulmonary blood vessels and Br so as to facilitate their fixation on the cuff. However, this method requires a special vascular clamp and has no advantage in reducing the cold ischemia time of the donor lung. In the recipient anastomosis process, recent studies have reported a pendulum anastomosis method8. However, this method has rigorous requirements for instruments and equipment. Beginners often find it difficult to obtain some key equipment and face the risk of long anastomosis time and even vascular tearing due to high tension during the anastomosis process. 
Some researchers first perform PV anastomosis during the anastomosis process. They believe that PV anastomosis is the most difficult step. Anastomosing the PV first can ensure that the donor lung has enough space to move. However, since the PV is tougher than the Br and PA, this leads to the risk of the PV cuff falling off or even venous tearing when anastomosing the Br and PA10. Some researchers have tried to reduce the tension during anastomosis by using the right semi-supine position8,11 or the supine position. However, the method of padding the right chest will change the angle of the endotracheal tube and increase the airway pressure7. The above researchers have made innovations in cuff production, donor lung modification, and donor lung implantation. Although this model has been improved by many researchers, it has not been widely promoted and applied.
In our study, we made improvements in cuff production, donor lung modification, and donor lung implantation in the hope that researchers can master this model more quickly. We chose a 16 G venous indwelling needle cannula as the PA cuff because the 16 G cuff can be easily anastomosed to arteries with better toughness, and there is enough space in the cuff to ensure good blood flow. Since the 18 G cuff is significantly smaller than the inner diameter of the recipient PA, this is not conducive to smooth blood flow, and during the anastomosis process, the cuff is too small and lacks sufficient friction between the recipient's PA wall, which increases the difficulty of the anastomosis and easily causes the cuff to slip off. 
Regarding the selection of the size of the Br and PV cuffs, most researchers have a relatively consistent opinion. It is worth noting that to minimize the occurrence of Br anastomotic obstruction, the Br cuff selected should be as large as possible. In the treatment of the cuff, Hiroaki et al.6 made some notches on the surface of the cuff to increase friction, and some researchers used cuffs without tails12,13. In our study, a cuff with a tail was used to facilitate fixation with hemostatic forceps. After the PA, Br, and PV were fixed, the tail was removed. This method can save time in fixing the PA, Br, and PV on the cuff and can prevent damage to the blood vessels and donor lungs due to the movement of the cuff tail in the recipient after anastomosis.
During the modification of the donor lung, the complex anatomical structure increases the cold ischemia time of the donor lung. In this study, the surface and bottom of the donor lung were covered with pretrimmed sterile lens paper, and the donor lung was placed on a sterile culture dish and then placed on ice for modification. The sterile lens paper covering the surface of the donor lung was slightly soaked with precooled saline. This not only ensures the moistening of the donor lung and prevents the donor lung from being damaged by forceps, but also provides sufficient friction for adjusting the position of the left and right lungs when separating the hilar structure, so as to fully expose the hilar anatomical structure and quickly separate the PA, Br, and PV from the dorsal side of the donor lung. In previous studies, gauze was placed on the surface of the donor lung7,10,14. Due to the large gap in the middle of the gauze, the donor lung was easily damaged by forceps during the separation process, eventually leading to pneumothorax. Our method can quickly free the PA, Br, PV, and other structures and shorten the cold ischemia time of the donor lung. 
During the donor lung implantation stage, securing the recipient's left lung in the anterior chest region aligns more closely with the recipient's anatomical structure. We significantly reduced the anastomotic tension between the donor lung and the recipient's blood vessels and bronchi, thereby lowering the surgical difficulty and the risk of postoperative complications. Additionally, the posterior thoracic space is relatively larger, facilitating the manipulation of surgical instruments and the implantation of the donor lung. In contrast, most previous studies fixed the recipient's left lung on the dorsal side7,8,12. This approach generates greater tension during the anastomosis process, increasing surgical difficulty and the risk of postoperative complications. Furthermore, the limited operating space in the anterior thoracic cavity may restrict the flexibility of surgical instruments. 
Lastly, this approach requires extensive retraction of the recipient's lung, which may increase the risk of mechanical injury. In addition, we retained the recipient's left lung before the anastomosis and trimmed a platform on it. This facilitated better placement of the donor lung and adjustment of its position during the anastomosis. In previous studies, the recipient's lung stump was completely removed before the anastomosis, and the PA, PV, and Br were clamped with instruments for fixation15, which could easily cause vascular tearing. Before the anastomosis, a heparinized saline solution should be used for vascular irrigation to remove blood clots and prevent thromboembolic complications. This has been reported in many previous studies7,10,16. However, many researchers have not paid attention to the fact that air bubbles can easily be introduced during the anastomosis process, leading to air embolism. In our procedure, we use a fine cotton swab to remove larger air bubbles from the recipient's blood vessels and the donor cuff, effectively preventing air embolism. Smaller bubbles can be absorbed and do not cause significant issues. 
In the anastomotic procedure, the pulmonary artery (PA) is addressed first, followed by the bronchus (Br), concluding with the pulmonary vein. This sequential approach is strategically adopted due to the PA's superior elasticity, which minimizes the risk of vascular injury or cuff displacement caused by excessive tension during subsequent venous anastomosis. Primary anastomosis of either the PV or Br may predispose to vascular or bronchial tearing, as well as cuff migration due to tension-related complications. Through systematic training protocols and repetitive practice, this optimized sequence has demonstrated significant advantages in reducing technical complexity and enhancing operative success rates. Notably, our approach has achieved a 100% anastomotic success rate, surpassing outcomes reported in comparable studies. During the implantation procedure, the operator only needs to retract the edge of the incision with one hand, insert the donor cuff into the recipient lumen, and secure it with one-handed knotting. This technique, which enables fixation and knotting with a single hand, significantly reduces personnel requirements and enhances surgical efficiency. Thus, our study improved the method of rat left lung orthotopic transplantation, reduced the time and difficulty of surgical operation, and improved the repeatability of this model.

Lung transplantation is currently the only effective treatment for end-stage lung diseases, which has developed rapidly in recent years. At present, more than 100,000 lung transplantation operations have been completed worldwide1, but the 5 year and 10 year survival rates after lung transplantation are only 54% and 32%, respectively2. Compared with other solid organ transplants, the survival rate after lung transplantation needs to be improved3. Various serious complications after lung transplantation are the key factors affecting the survival rate of recipients, including primary graft dysfunction, acute rejection, chronic lung allograft dysfunction, and lung infection4,5. The rat lung transplantation model is a valuable tool for studying these issues, but its widespread adoption has been hindered by technical difficulties. Although the procedure has been reported by several laboratories, it involves a steep learning curve, high postoperative complication rates, and low survival rates, limiting its accessibility to researchers.
In this study, we present an improved orthotopic lung transplantation technique designed to overcome these challenges. Our method builds upon existing protocols but introduces key refinements, including cuff preparation, hilar structure dissection, donor lung fixation, and optimized anastomosis techniques. These improvements enable the procedure to be performed by a single surgeon with minimal equipment, reduce anastomosis difficulty, shorten cold and warm ischemia times, and ultimately enhance the success rate of the model. This approach aims to support more robust and reproducible basic research in lung transplantation.

<table><tbody><tr><td>0.9% saline</td><td>Hopebio</td><td>HBPP008-100</td><td></td></tr><tr><td>10 cm sterile Petri dish</td><td>Beckman</td><td>10 cm plastic Petri dish</td><td></td></tr><tr><td>16 G tracheal intubation guide wire</td><td>vega</td><td>-</td><td></td></tr><tr><td>Dumont fine forceps #7-curved/11.5 cm</td><td>F.S.T</td><td>11271-30</td><td></td></tr><tr><td>Eye scissors 10 cm straight sharp</td><td>Jingzhong</td><td>Y00030</td><td></td></tr><tr><td>Hemostatic (curved full-tooth) forceps</td><td>Jingzhong</td><td>J31180</td><td></td></tr><tr><td>Heparin sodium solution</td><td>Leagene</td><td>R10119</td><td></td></tr><tr><td>Introcan needle 14 G butterfly needle</td><td>B.Braun</td><td>4254210B</td><td></td></tr><tr><td>Introcan needle 16 G butterfly needle</td><td>B.Braun</td><td>4254171B</td><td></td></tr><tr><td>LED dual-fiber cold light source lamp</td><td>RWD</td><td>76312</td><td></td></tr><tr><td>Lens cleaning paper</td><td>Kimberly</td><td>34155</td><td></td></tr><tr><td>Lewis rats&amp;nbsp;</td><td>Guangzhou Dean Gene Technology Co., LTD. (Guangzhou, China)</td><td></td><td></td></tr><tr><td>Medical cotton swabs</td><td>Winner/Wenjian</td><td>10 cm x 50 pieces</td><td></td></tr><tr><td>Medical needle holder</td><td>Chenghe</td><td>-</td><td></td></tr><tr><td>Medical sterile syringe</td><td>Winner/Wenjian</td><td>20ml</td><td></td></tr><tr><td>Medical surgical scissors 12.5 cm straight sharp</td><td>Pulun</td><td>PL1251</td><td></td></tr><tr><td>Medical tape</td><td>Wenxian</td><td>1527</td><td></td></tr><tr><td>Medical tissue forceps</td><td>RWD</td><td>J30607</td><td></td></tr><tr><td>Medical titanium eyelid retractor</td><td>Youdikang</td><td>-</td><td></td></tr><tr><td>Micro curved forceps</td><td>Jingzhong</td><td>JzCrWA3050 0.3/140 mm</td><td></td></tr><tr><td>Micro vascular hemostatic forceps 12.5 cm curved</td><td>Jinzhong</td><td>W40350</td><td></td></tr><tr><td>Micro vascular hemostatic forceps 12.5 cm straight</td><td>Jinzhong</td><td>W40340</td><td></td></tr><tr><td>Micro-CT</td><td>Pinseng</td><td>SNC-100</td><td></td></tr><tr><td>Miniature vascular clips</td><td>F.S.T</td><td>18055-06</td><td></td></tr><tr><td>Multi-channel small animal anesthesia machine</td><td>RWD</td><td>R550</td><td></td></tr><tr><td>Neurosurgical coagulation kit, coagulation kit</td><td>F.S.T</td><td>18010-00</td><td></td></tr><tr><td>Pentobarbital sodium</td><td>Sigma</td><td>P3761</td><td></td></tr><tr><td>Pointed cotton swabs</td><td>Other</td><td>1000 pieces</td><td></td></tr><tr><td>Rat/Mouse intubation platform</td><td>Puxin</td><td>-</td><td></td></tr><tr><td>Rat/Mouse water bath system (intraoperative) 10-65 &amp;deg;C temperature control, 310 x 290 MM, 220 V</td><td>Huayang</td><td>36-0041</td><td></td></tr><tr><td>Shanghai Golden Bell medical microscopic forceps Microscopic fine forceps straight bending microhand surgical instruments Microscopic instruments</td><td>Jingzhong</td><td>WA1020</td><td></td></tr><tr><td>Shanghai Pudong Jinhuan Medical absorbable surgical sutures Sterile sutures with needle implantation lifts (4-0)</td><td>Shanghai Pudong Jinhuan</td><td>473445</td><td></td></tr><tr><td>Small animal ventilators</td><td>Kent Scientific</td><td>RoVent Jr</td><td></td></tr><tr><td>Small animal water bath insulation system</td><td>HUAYON</td><td>TG-TP-BL</td><td></td></tr><tr><td>Spring scissors (large)</td><td>Jing Zhong</td><td>JzCrWA1020</td><td></td></tr><tr><td>Stainless steel medical mosquito forceps</td><td>Jiate</td><td>LZ1021</td><td></td></tr><tr><td>Steel flat handle curved non-toothed 0.3 teeth</td><td>Zhenbang (Medical)</td><td>Toothed with hook/platform for tying</td><td></td></tr><tr><td>Steel flat handle straight non-toothed 0.3 teeth</td><td>Zhenbang (Medical)</td><td>Toothed with hook/platform for tying</td><td></td></tr><tr><td>Sterile cotton balls</td><td>Boshengtianai</td><td>Small size</td><td></td></tr><tr><td>Sterile gauze</td><td>Hisern</td><td>6 cm x 8 cm</td><td></td></tr><tr><td>Suture with needle</td><td>Lingqiao</td><td>8-0</td><td></td></tr><tr><td>Suture with needle</td><td>Lingqiao</td><td>6-0</td><td></td></tr><tr><td>Tweezers Dumont #5 Forceps, Straight</td><td>F.S.T</td><td>11254-20</td><td></td></tr><tr><td>Ultra-light clay</td><td>Bokee</td><td>CQNT-24</td><td></td></tr><tr><td>Vascular clamp holder</td><td>RWD</td><td>R34001-14</td><td></td></tr></tbody></table>

orthotopic left lung transplantation in rats

Orthotopic Left Lung Transplantation in Rats

The rat left lung orthotopic transplantation model is an essential tool in lung transplant research. Although lung transplantation is a complex procedure, its complexity is significantly reduced with the introduction of the cuff technique. Most surgeons can master this technique after training, but its application has been limited due to the long learning curve and low model survival rate. To address these issues, we propose an improved technique with high repeatability, which includes donor lung harvest, modification, and implantation. Compared to previous methods, our improvements primarily focus on the dissection of the donor lung harvest, donor lung modification, and donor lung implantation. This modification simplifies the procedure, reduces operational difficulty, and increases repeatability. Additionally, the cold ischemia and warm ischemia times during the procedure have been shortened, further improving the success rate of transplantation. Our research provides a more efficient and reproducible technical foundation for the rat lung transplantation model, supporting future research in lung transplantation.

We recorded the operation time of each step in the process of lung transplantation in 30 rats. The time of cuff preparation, donor lung harvest, cold ischemia, and warm ischemia are shown in Table 1. Using our improved method, after preliminary training, the success rate of transplantation surgery reached 100%. This technology can provide support for subsequent research on complications after lung transplantation. We have used this technology for more than 2 years. The following figures show our process from cuff making (Figure 1), donor lung extraction (Figure 2C), donor lung modification (Figure 3), to donor lung implantation (Figure 4) and evaluation of surgical results (Figure 5). Six months after the syngeneic transplantation, we performed Computed Tomography (CT) imaging and pathological evaluation on the recipients. We found that the ventilation of the transplanted lungs was good in CT imaging, similar to that of the sham operation group (only thoracotomy without lung transplantation); there was no significant difference in the macroscopic appearance of the ex vivo lungs between the sham operation group and the autologous right lung, and no obvious pathological changes were found in the Hematoxylin-Eosin (HE) staining results. In summary, our rat left lung orthotopic lung transplantation was successful, and the maturity of this technology laid the foundation for the construction of other disease models after lung transplantation.
<img alt="Catheter gauge comparison, 14G vs 16G, includes size measurement using a razor blade and ruler setup." class="xfigimg" src="/files/ftp_upload/68106/68106fig1.jpg" data-altupdatedat="1783396841000" data-alt="Figure 1"> 
Figure 1: Cuff preparation. (A) Different sizes of IV cannulae and cuff cutting blades. (B) Magnification of 5x. Cutting the cuff under a microscope. Scale bar = 1 mm. (C) Cuffs for the PA (16 G), PV (16 G), and Br (14 G). Abbreviations: IV = intravenous; PA = pulmonary artery; PV = pulmonary vein; Br = bronchi. <a href="https://www.jove.com/files/ftp_upload/68106/68106fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Organ harvesting procedure; mouse dissection; surgical tools; biological research method." class="xfigimg" src="/files/ftp_upload/68106/68106fig2.jpg" data-altupdatedat="1783396841000" data-alt="Figure 2"> 
Figure 2: Donor lung harvest. (A) Donor endotracheal intubation. (B) Donor lung perfusion. (C) Place the clean perfused donor lung on ice. <a href="https://www.jove.com/files/ftp_upload/68106/68106fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Surgical anatomy, tissue dissection, labeled PA, Br, PV, close-up, experimental setup." class="xfigimg" src="/files/ftp_upload/68106/68106fig3.jpg" data-altupdatedat="1783396841000" data-alt="Figure 3"> 
Figure 3: Donor lung modification. (A) Free the PA. (B) Free and ligate the Br. (C) Free the PV (D-F) Fix the PA, Br, and PV to the cuff successively. All microscopic images were captured at a magnification of 5x. Scale bar = 1 mm. Abbreviations: PA = pulmonary artery; PV = pulmonary vein; Br = bronchi. <a href="https://www.jove.com/files/ftp_upload/68106/68106fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Surgical procedure on rat heart; diagram; PV, Br, PA; cardiovascular research; experimental setup." class="xfigimg" src="/files/ftp_upload/68106/68106fig4.jpg" data-altupdatedat="1783396841000" data-alt="Figure 4"> 
Figure 4: Donor lung implantation. (A) The recipient was fixed on the thermostatic operating table; the chest wall was opened; and the left lung was retracted and fixed on the incision. (B) Free and block the hilar structures. (C) Pretied surgical knot. (D) Make a small incision at the distal end of the PA, Br, and PV. (E) Flush with heparinized saline. (F,G) Anastomosis of PA, Br, and PV in sequence. (H) Open the hemostatic clip to allow ventilation and blood circulation. (I) Resection of the recipient's left lung. (J) Return the donor lung to the chest cavity. (K) Closing the chest. (L) Waiting for the recipient to wake up. All microscopic images were captured at a magnification of 5x. Scale bar = 1 mm. Abbreviations: PA = pulmonary artery; PV = pulmonary vein; Br = bronchi. <a href="https://www.jove.com/files/ftp_upload/68106/68106fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="CT scan, macroscopic, and HE staining in lung transplantation study; donor and sham comparison." class="xfigimg" src="/files/ftp_upload/68106/68106fig5.jpg" data-altupdatedat="1783396841000" data-alt="Figure 5"> 
Figure 5: Posttransplantation evaluation. The sham operation group only underwent thoracotomy without transplantation. CT imaging and pathological evaluation of the samples were performed 6 months later. The transplant group received syngeneic transplantation and underwent CT imaging and pathological evaluation 6 months later. <a href="https://www.jove.com/files/ftp_upload/68106/68106fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>procedure</td>
			<td>Operation time (min)</td>
		</tr>
		<tr>
			<td>Cuff preparation</td>
			<td>3.2±0.38</td>
		</tr>
		<tr>
			<td>Donor Lung Harvest</td>
			<td>9.85±0.60</td>
		</tr>
		<tr>
			<td>Cold ischemia</td>
			<td>29.96±2.77</td>
		</tr>
		<tr>
			<td>Warm ischemia</td>
			<td>12.62±2.35</td>
		</tr>
	</tbody>
</table>
Table 1: Operation time of each procedure. Cuff preparation: the total time taken to make three cuffs (including cuffs for PA, PV, and Br), Donor lung harvest: the time taken to remove the donor lung from the donor. Cold ischemia: the duration for which the donor lung is kept on ice. Warm ischemia: the time taken from the removal of the donor lung from ice to reperfusion. Abbreviations: PA = pulmonary artery; PV = pulmonary vein; Br = bronchi.

This protocol describes an improved technique for orthotopic left lung transplantation in rats, focusing on reducing operational difficulty and improving survival rates.

The rat left lung orthotopic transplantation model is an essential tool in lung transplant research. Although lung transplantation is a complex procedure, ...

My research focused on developing standardized, reproducible left lung transplantation model to investigate post-transplant complications, particularly chronic rejection, and to better understand the mechanism driving long-term graft failure. Our protocol offers a shorter learning curve, single operational capability, and a higher reproducibility compared to existing techniques. Our standardized direct lung transplantation model provides a reliable platform for studying complications after lung transplantation and can accelerate translational research to improve graft survival.In the future, our laboratory will focus on the study of chronic rejection after lung transplantation in order to improve the long-term survival rate of lung transplant recipients. To begin, obtain 14-gauge and 16-gauge intravenous catheters and blades. Trim the catheters into a cuff with a tail with a blade.Then divide each cuff into a body and a tail, each approximately two to three millimeters in length. Using a scalpel, create superficial scratches on the cuff body to increase friction for suture fixation. To harvest the donor lung, retract the tongue of an anesthetized rat outward and upward using forceps.Position a surgical lamp anterior to the neck to illuminate the glottal opening clearly. Now, insert a 14-gauge intravenous cannula into the airway through the glottis. Set the ventilator to pressure controlled mode.Then, input the weight parameters, adjust the pressure to 15 centimeters of water, and connect the ventilator. Observe the rise and fall of the chest to ensure synchronization with the ventilator frequency. Next, fix the limbs and head using tape or restraints.Make a midline incision along the abdominal wall. Inject heparin through the exposed peritoneal vein, then allow systemic circulation for three minutes to ensure complete heparinization. Cut the diaphragm and open the chest cavity from the midline of the sternum.Then, fix the chest wall on both sides with hemostats. Remove the thymus to expose the chest cavity organs. Sequentially, cut the superior vena cava, inferior vena cava, and the left and right auriculi.Now, inject cold saline into the root of the pulmonary artery using a 20-milliliter syringe over one minute for low pressure perfusion until the donor lung turns completely white. Cut tissues connecting the donor lung and extract the donor heart and lung in block. Soak the donor heart-lung block in pre-cooled saline.Position the heart-lung block on ice under a microscope. Grip the trachea with a hemostat and secure it in plasticine. Cover the lungs with wet sterile lens paper.Now, use forceps to separate the pulmonary artery, bronchus, and pulmonary vein under the microscope. Ligate the bronchus close to the lung using 6-0 surgical sutures. Use spring scissors to cut the pulmonary artery, bronchus, and pulmonary vein.Extract the pulmonary artery, bronchus, and pulmonary vein from respective cuffs made from 16-gauge, 14-gauge, and 16-gauge indentation catheters. Fix the tube walls to the cuff with 8-0 surgical sutures. Preserve the donor lungs in saline, and place them back on ice until implantation.Depilate the left thoracodorsal region of the recipient animal. Position the animal in the right lateral decubitus posture on a thermostatic operating table. Now, make an incision through the skin and chest wall at the point of the apical impulse in the fourth intercostal space to access the thoracic cavity.Use an eyelid retractor to open the thoracic cavity. Then, gently push the left lung aside using a moist cotton swab to expose and sharply transect the inferior pulmonary ligament. Next, grasp the left lung with forceps and retract it outside the thoracic cavity.Secure the hilum with a hemostat, then stabilize it with modeling clay. Now, dissect the pulmonary hilum to separate the pulmonary artery, pulmonary vein, and bronchus. Clamp the proximal ends of the pulmonary artery, pulmonary vein, and bronchus using vascular clamps.Pre-tie the recipient's pulmonary artery, bronchus, and pulmonary vein with 8-0 surgical sutures for rapid anastomosis. Trim a platform on the recipient's clamped left lung to facilitate the placement of the donor lung. Make small incisions at the distal ends of the pulmonary artery, pulmonary vein, and bronchus.Rinse the pulmonary artery and pulmonary vein with heparinized saline to prevent clot formation. Then, take the donor lung from ice and position it on the prepared platform of the recipient's left lung. Use forceps to lift one side of the cut opening, then sequentially implant and ligate the cuffs of the pulmonary artery, pulmonary vein, and bronchus into the recipient.Open the micro hemostatic clip and observe the donor lung turning from white to red. Inspect the anastomotic sites for any signs of bleeding. Now, remove the recipient's left lung and return the donor lung into the thoracic cavity.Wipe the chest cavity dry with sterile cotton. Then, close the thoracic cavity layer by layer. Wait for spontaneous breathing recovery before removing the ventilator.After one hour, the recipients were free to eat. Six months after transplantation, CT imaging showed that the ventilation of the transplanted lungs was comparable to the sham group, indicating preserved lung function. The macroscopic appearance of the lungs did not differ visibly between the sham and transplanted groups, showing no signs of deterioration or abnormality.Hematoxylin and eosin staining of the lung tissues revealed no significant pathological changes in the transplanted lungs compared to the sham group, with preserved alveolar structures and absence of inflammatory infiltration.

orthotopic-left-lung-transplantation-in-rats

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JoVE Journal

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Orthotopic Left Lung Transplantation in Rats

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