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

1. Donor Procedure

1. All surgical procedures were performed utilizing sterile technique. No antibiotics are given to both donor and recipient mice.
2. Induction of anesthesia of the donor mouse is initiated with 5% Isoflurane.
3. The mouse is orotracheally intubated with a 20-gauge intravenous catheter and then placed on a rodent ventilator, using 100% oxygen at a rate of 125 breaths/minute and approximately 0.5 ml tidal volume (2% of its body weight).
4. Anesthesia is maintained with inhaled 1-2% Isoflurane.
5. The donor mouse is placed in a supine position, prepped with 70% alcohol.
6. Laparosternotomy is performed as a combined midline and transverse incision.
7. Heparin at 100 u/kg is injected into the Inferior vena cava (IVC), from just below the liver.
8. The diaphragm is cut along the ventral costal attachment toward the spine, and the thoracic cavity is exposed by cutting both sides of the chest wall to the neck.
9. After incising the IVC at the level of diaphragm, the right atrial appendage is cut and the lungs flushed with 2 ml of cooled (4 °C) Lactated Ringer’s injection and 0.1 ml of heparin via a transverse incision at the root of the pulmonary artery (PA) trunk.
10. Arresting ventilation at two thirds of end-tidal inflation, the heart-lung block is excised and stored on ice (4 °C).
11. The donor left lung is prepared for the recipient by the attachment of cuffs. The pulmonary ligament is incised up to the Pulmonary Vein (PV). The hilum is brought out by removing esophagus and aorta.
12. Identify the PA, located at the most cranial aspect of the hilum, as well as the attached main bronchus (Br). Carefully dissect the PA from the bronchus.
13. PA cuff is made from a 24-gauge Intravenous (I.V.) catheter and cut to 0.5 mm in length with an extension of 0.7 mm.
14. The entire surface of the cuff is then abraded to facilitate anastomosis.
15. The cuff for the bronchus is derived from a 20-gauge IV catheter, and cut in length of 1.0 mm with an extension of 0.7 mm.
16. Using the same materials, the PV cuff varies with the weight of the donor mouse. Specifically, for mice 24-27 grams the cuff size is 22-gauge, 0.7 mm in length with a 0.7 mm extension. For mice weighing 27-32 grams, the studies utilized a 20-gauge catheter that is 0.7 mm in length with a 0.7 mm extension.
17. The cuffs are inserted into the distal ends of the PA, PV and Br and secured with a 9-0 suture.
18. The donor lung is flushed and washed with sterile saline with sterile heparin before storage. The donor lung is then wrapped with sterile gauze soaked in sterile saline on ice (4 °C), which keeps the lung very clean and sterile.
19. A microvessel clamp is placed on the bronchus to prevent Lactated Ringer entry into the airway.

2. Recipient Procedure

1. Induction of anesthesia and mechanical ventilation are the same as described for the donor above.
2. The left chest wall is shaved and prepped with 70% alcohol and the surgical field is draped.
3. A thoracotomy incision is made in the left third intercostal space, extending the incision dorsally close to the spine, and a microvessel clamp placed on the left pulmonary vessels and bronchus adjacent to the heart. You can see PA at the cranial aspect, PV at the caudal end of hilum and Br between them.
4. Using gentle traction on the hemostat to cause mild tension on the PA, Br and PV, the left lung is pulled out from the thoracic cavity while leaving the central hilar structures clamped.
5. The PA, PV and Br are isolated by blunt dissection followed by placing a 9-0 suture positioned loosely around the PA, PV and Br.
6. After dissecting the PA completely from its adventitial sheath, a small transverse incision of approximately one fourth of the vessel’s circumference is made into the anterior wall, leaving the continuation of the dorsal part of the artery intact.
7. The donor lung, wrapped in cold, Lactated Ringer-soaked cotton gauze and prepared as described above, is then positioned into the thoracic cavity, and cuffs inserted into the recipient PA, PV and Br and secured with 9-0 suture.
8. The hilar cross clamp is removed allowing for reperfusion and ventilation.
9. After positioning the transplanted lung back into the recipient thorax, the thoracotomy incision is closed using a 5-0 suture.
10. The mouse is allowed to recover from anesthesia. Buprenorphine (0.05-0.15 mg/kg) is administered immediately after surgery, and for every 8 hours for 2-3 days post-surgery.

3. Representative Results

Our experience has taught us that it requires several months of repeated practice to become proficient in the mouse lung transplant model. After proficiency is attained, we achieved a 96% (96/100 consecutive surgeries) perioperative survival rate with deaths occurring within seven days post operatively. Two deaths were due to bleeding that began intraoperatively, and pneumothraces were the cause of death in the other two mice. For all procedures, the warm ischemia time was 14.32 ± 3.14 minutes, and cold ischemia time was 58.51± 18.06 minutes. Three orthotopic lung transplant groups were studied: isograft: C57BL/6→C57BL/6, allograft: C57BL/10→C57BL/6 and C57BL/6→C57BL/10. We used only male mice but our technique can be also applied to female mice, because there is no significant anatomical difference between the sexes.

Grading of rejection pathology was conducted in a blinded fashion utilizing standard criteria for clinical lung transplantation⁷(Table 1). Whereas we observed mild or no rejection in isograft (C57BL/6→C57BL/6), both allograft combinations developed comparable acute or chronic rejection (Figure 1). In contrast, OB was significantly more frequent in the C57BL/10→C57BL/6 than C57BL/6→C57BL/10 group by day28 (Table 1).

Table 1. Histological scores of acute rejection and prevalence of obliterative bronchiolitis post transplant. Scoring of acute rejection (“A” Scores) by standard criteria as described in representative results. Data represents the mean ± SD of “A” Scores at days 28 post transplantation. Data represents the quantity and percentage of mice in each group that developed OB at days 21 and 28 post transplantation.

Figure 1. Macroscopic findings and histopathology at 28 days post lung transplantation. Panel 1A represent macroscopic findings and H&E stained Isograft lung and right naïve lung. Panel 1B and 1C represents H&E and Masson’s Trichrome stained BL/10 lung allograft transplanted into BL/6 mouse recipient, which developed OB and non OB, respectively. The white arrow in 1B identifies lesions of OB. Panel 1D shows BL/6 lung allograft transplanted into BL/10 mouse recipient.

Main trouble shooting during the procedure were as follows.

1. Blood flow disorder: Flush donor lung until the color becomes white in color. Use of excessive pressure could induce pulmonary edema. But do not push too much to prevent edema after transplant.
2. Difficulty of cuff insertion into the donor: Ascertain that vasculature is of sufficient length and free of attached fat and connective tissues.
3. Bleeding at anastomotic site: Use Q tips to apply pressure at affected site for approximately 5 minutes.
4. Recipient PA blood flow obstruction: Mainly caused by torsion of the during cuff insertion. If air is observed within the PA then re-do of the anastomosis is necessary.
5. Difficult PV anastomosis: Membrane of PV is very thin and easily tears. Select proper cuff size and insert straight to the recipient’s PV very gently.
6. Recipient PV blood flow obstruction: Make sure the position and direction of the cuff. Release the compression from the bronchial cuff.
7. Pneumothorax: This may be caused by either bronchial perforation due to manipulation of the airway, or perforation of lung surface due to trauma associated with the transplant procedure. The former is treated with surgical closure of the hole within the airway wall; and the latter is treated by oversewing the leak on the lung surface using 10-0 nylon suture.