Orthotopic Left Lung Transplantation in Rats

Summary

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

Abstract

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.

Introduction

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 infection^4,5. 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.

Protocol

All animal experiments have been approved by the Animal Ethics Committee of Guangzhou Lai'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)

1. Prepare 14 G and 16 G intravenous catheters and blades (Figure 1A).
2. 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).
3. 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.

2. Donor lung harvest (Figure 2)

1. Anesthetize the donor rat via intraperitoneal injection of pentobarbital (80 mg/kg) and administer subcutaneous injection 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.
2. 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).
3. Insert a 14 G intravenous cannula into the airway through the glottis.
4. Set the ventilator to pressure-controlled mode, input the weight parameters, adjust the pressure to 15 cmH₂O, and connect the ventilator.
5. 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.
6. Pinch the rat'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. Lift the abdominal skin with tweezers, and use scissors to cut the skin from the abdomen, chest, and front of the neck.
7. 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.
8. 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.
9. 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).
10. 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.

3. Donor lung modification (Figure 3)

1. Position the heart-lung block on ice under the microscope. Grip the trachea with a hemostat and secure it in plasticine.
2. 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.
3. 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).
4. 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).
5. Preserve the modified donor lungs in saline, repositioned on ice, and await implantation.

4. Donor lung implantation (Figure 4)

1. 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.
2. Depilate the recipient'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.
3. 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.
4. 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.
5. 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's side (Figure 4A).
6. Remove the tissue around the left lung, dissect the pulmonary hilum, and separate the PA, PV, and Br.
7. Clamp the proximal ends of the PA, Br, and PV using vascular clamps (Figure 4B).
8. Use 8-0 surgical suture to pre-tie the recipient's PA, Br, and PV, ensuring rapid anastomosis during donor lung implantation (Figure 4C).
9. Trim a platform on the clamped recipient's left lung to facilitate the placement of the donor lung. Make a small incision at the distal end of the recipient's PA, Br, and PV (Figure 4D).
10. Rinse the PA and PV with heparinized saline to prevent thrombus formation (Figure 4E).
11. Take the donor lung out of the ice and place it on the recipient'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.
12. 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).
13. Remove the recipient's left lung, and return the donor lung to the chest cavity (Figure 4I,J).
14. 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's condition. If the recipient exhibits gasping respirations, it suggests the presence of hypoxia, and the ventilator parameters should be promptly adjusted accordingly.
15. Wipe the chest cavity dry with a sterile cotton ball, then close it layer by layer (Figure 4K).
16. 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.
17. House the recipients in individual cages with free access to food and water, and maintain the temperature at ~22 °C.
18. Administer subcutaneous injections of meloxicam (5 mg/kg) to the recipients for 3 consecutive days to relieve pain.

Results

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.

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. Please click here to view a larger version of this figure.

Figure 2: Donor lung harvest. (A) Donor endotracheal intubation. (B) Donor lung perfusion. (C) Place the clean perfused donor lung on ice. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

procedure	Operation time (min)
Cuff preparation	3.2±0.38
Donor Lung Harvest	9.85±0.60
Cold ischemia	29.96±2.77
Warm ischemia	12.62±2.35

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.

Discussion

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 cuff^7,10. 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 position^8,11 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 tails^12,13. 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 lung^7,10,14. 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 side^7,8,12. 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 studies^7,10,16. 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.

Materials

List of materials used in this article

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