Murine Intrapulmonary Tracheal Transplantation: A Model for Investigating Obliterative Airway Disease After Lung Transplantation

Summary

The murine intrapulmonary tracheal transplantation (IPTT) model is valuable for studying obliterative airway disease (OAD) after lung transplantation. It offers insights into lung-specific immunological and angiogenic behavior in airway obliteration after allotransplantation with high reproducibility. Here, we describe the IPTT procedure and its expected results.

Abstract

Murine intrapulmonary tracheal transplantation (IPTT) is used as a model of obliterative airway disease (OAD) following lung transplantation. Initially reported by our team, this model has gained use in the study of OAD due to its high technical reproducibility and suitability for investigating immunological behaviors and therapeutic interventions.

In the IPTT model, a rodent tracheal graft is directly inserted into the recipient's lung through the pleura. This model is distinct from the heterotopic tracheal transplantation (HTT) model, wherein grafts are transplanted into subcutaneous or omental sites, and from the orthotopic tracheal transplantation (OTT) model in which the donor trachea replaces the recipient's trachea.

Successful implementation of the IPTT model requires advanced anesthetic and surgical skills. Anesthetic skills include endotracheal intubation of the recipient, setting appropriate ventilatory parameters, and appropriately timed extubation after recovery from anesthesia. Surgical skills are essential for precise graft placement within the lung and for ensuring effective sealing of the visceral pleura to prevent air leakage and bleeding. In general, the learning process takes approximately 2 months.

In contrast to the HTT and OTT models, in the IPTT model, the allograft airway develops airway obliteration in the relevant lung microenvironment. This allows investigators to study lung-specific immunological and angiogenic processes involved in airway obliteration after lung transplantation. Furthermore, this model is also unique in that it exhibits tertiary lymphoid organs (TLOs), which are also seen in human lung allografts. TLOs are comprised of T and B cell populations and characterized by the presence of high endothelial venules that direct immune cell recruitment; therefore, they are likely to play a crucial role in graft acceptance and rejection. We conclude that the IPTT model is a useful tool for studying intrapulmonary immune and profibrotic pathways involved in the development of airway obliteration in the lung transplant allograft.

Introduction

Lung transplantation has been established as an effective treatment for patients with end-stage respiratory diseases. However, the median survival rate for human lung transplant recipients is only approximately 6 years, with the development of obliterative bronchiolitis (OB), a type of obstructive airway disease (OAD), being a major cause of death after the first year post transplantation¹.

Several animal models have been utilized to investigate the mechanism underlying OAD. One such model is the heterotopic tracheal transplantation (HTT) model². In this model, tracheal grafts are implanted into the recipient's subcutaneous tissue or omentum. Ischemia-induced loss of tracheal graft epithelial cells occurs, followed by alloreactive lymphocyte infiltration and apoptosis of donor epithelial cells. Fibroblasts and myofibroblasts migrate around the trachea, producing an extracellular matrix. Finally, complete fibrous obliteration of the airway lumen occurs. The HTT model is technically simple, provides an in vivo environment, and offers high reproducibility.

Another model for studying OAD is the rat orthotopic tracheal transplantation (OTT) model, where tracheal grafts are interposed into the recipient's trachea to maintain physiological ventilation³. In this model, ischemia-induced depletion of donor epithelial cells results in their replacement by recipient epithelial cells within the trachea, forming an unobstructed airway accompanied by moderate fibrosis. Although these models have contributed to the understanding of airway obliteration after lung transplantation, they have limitations in terms of recapitulation of the lung parenchymal microenvironment.

Our research group introduced the rat intrapulmonary tracheal transplantation (IPTT) model, where tracheal grafts are implanted into the recipient lung⁴ (Figure 1). The IPTT model exhibits fibrous obliteration of the airway lumen occurring within the lung microenvironment. Furthermore, it has been successfully applied to mice that are technically more challenging than rat IPTT^5,6,7,8,9,10. This adaptation of the murine IPTT model enabled us to delve deeper into the intricate details of the lung immunological environment of OAD after lung transplantation using transgenic mice.

The IPTT model possesses some unique features. One is neoangiogenesis, which is facilitated by pulmonary circulation and plays a crucial role in airway obliteration^4,10. Additionally, the IPTT model exhibits lymphoid aggregates, some of which have high endothelial venules expressing peripheral node addressin, indicating that they are tertiary lymphoid organs (TLOs)^7,8. TLOs resemble lymph nodes and consist of T cells, B cells, and frequently, a germinal center accompanied by follicular dendritic cells^11,12. TLOs have been reported in various chronic inflammatory diseases, including airway obliteration, making the IPTT model suitable for investigating the role of TLOs in airway obliteration^7,8,11,12,13. This paper presents the methodology of the murine IPTT model, with the goal of familiarizing researchers with this model and facilitate further investigations into airway obliteration following lung transplantation.

Protocol

All animals were treated in accordance with the guidelines set forth by the Canadian Council on Animal Care in the Guide to the Care and Use of Experimental Animals. The experimental protocol was approved by the Animal Care Committee of the Toronto General Hospital Research Institute, University Health Network.

1. Donor surgery

NOTE: BALB/c mice are used as an example of donors for the experiment. All procedures must be performed utilizing a sterile technique.

1. Prior to the procedure, record the weight of each mouse.
2. Euthanize the mouse using a CO₂ chamber.
3. Once death is confirmed, position the mouse in a supine position and secure the limbs with tape.
4. Prep the surgical area by sterilizing it with 70% isopropyl alcohol.
   NOTE: If needed and as recommended by the local animal ethics committee, clip the fur from the incision site.
5. Make a midline incision on the skin, starting from the mid-abdomen and extending to the anterior cervical region.
6. Access the trachea by carefully retracting the fat pads, laterally moving the strap muscles, and separating the trachea from surrounding connective tissue. Use forceps to create space between the trachea and the esophagus.
7. Lift the xiphoid and cut the diaphragm.
8. Raise the sternum, ensuring a clear path from the sternum to the neck region by inserting a hemostat. Clamp the rib cage on both sides and cut through the sternum, extending up through the neck muscles.
9. Remove the thymus and any fat or muscle obstructing the trachea to expose the tracheal bifurcation.
10. Cut both the main bronchi and carefully separate the airway from the esophagus.
11. Cut the larynx and remove it.
12. Spray the dissected trachea with sterile saline solution or preservation solution with sterile gauze soaked in sterile saline or preservation solution and place it on ice to preserve its viability.

2. Recipient surgery

NOTE: C57BL/6 mice are used as an example of recipients for the experiment.

1. Administer sustained-release buprenorphine subcutaneously at a dose of 1 mg/kg on the morning of the surgery day.
2. Induce anesthesia in an induction chamber using 5% isoflurane.
3. Once the mouse is lightly anesthetized, intraperitoneally inject a cocktail consisting of (0.1 mg/g) xylazine and (0.01 mg/g) ketamine.
4. Return the mouse to the induction chamber with 2-3% isoflurane maintained.
5. Shave the fur at the surgical site. Also, administer bupivacaine as a line block subcutaneously along the planned insicion site at a dose of 7 mg/kg.
6. Confirm the absence of reflex response to a toe pinch before orotracheal intubation. Intubate the mouse orotracheally using a 20 G intravenous catheter and connect it to a ventilator with a tidal volume of 500 µL, a respiratory rate of 120 bpm, 100% oxygen and 2% isoflurane. Use a stand with a clamp applied to the tongue, holding the animal in a vertical position with the neck extended, to facilitate this procedure.
7. Activate a heating pad and position the mouse in a right lateral position on top of the pad, with the head away from the surgeon and the tail facing the surgeon (Figure 2). Secure the limbs with tape. Put veterinary ointment on the eyes to prevent dryness while under anesthesia.
8. Scrub the surgical area with 7.5% povidone iodine, sterilize with 70% isopropyl alcohol, and re-scrub with 10% povidone iodine. Apply sterile surgical drapes to cover the surgical area.  
9. Load the donor trachea into a 16 G intravenous catheter during this time (Figure 3C,D).
10. Use scalpel to make an incision in the recipient's skin and cauterize the muscle and connective tissue.
11. Open the fifth or sixth intercostal space and hold the rib cage open using two retractors.
12. Dissect the inferior pulmonary ligament using cotton swabs and scissors.
13. Simulate the creation of the pathway for the donor trachea (Figure 3G,H).
14. Secure the ventilator outflow tube by partially occluding it with a three-way stopcock to facilitate inflation of the left lung.
15. Create a pathway by puncturing the left lung with a 20 G needle. Ensuring that the puncture depth is roughly equivalent to the length of the tracheal allograft. Select the puncture site at the lung's edge (as indicated in Figure 3I), ensuring the pathway runs parallel to the tabletop (as marked by a blue circle in Figure 3J).
    NOTE: An upward insertion angle will result in penetration of the pleural layer, while a deeper angle may lead to bleeding from major vessels (as marked by red crosses in Figure 3J).
16. Insert the 16 G intravenous catheter into the left lung and extrude the donor trachea into the left lung. After insertion of the tracheal allograft, release the three-way stopcock to allow unobstructed expiratory flow through the outflow tube.
17. Close the pleural injection site with a clip (Figure 3K,L). Position the clip precisely onto the puncture site, with its edge aligned to match the contour of the lung's edge (indicated by the blue circle in Figure 3L).
    NOTE: An incorrect location for the clipping site can result in ineffective sealing and air leakage, while insufficient clip depth may lead to the clip detaching post-surgery (as shown by the red crosses in Figure 3L).
18. Fill the thoracic cavity with saline solution and absorb the saline with a gauze.
19. Reinflate the left lung and close the ribs using a running suture technique.
20. Close the muscle and skin with interrupted sutures.
21. Administer meloxicam analgesic subcutaneously at a dose of 5 mg/kg at the end of the surgery.
22. Observe the recipient mouse until it is awake. Then, remove the tracheal tube and put the recipient mouse in a cage.
    NOTE: Recipient mice should be housed individually.
23. Administer meloxicam (5 mg/kg) once daily via subcutaneous injection, starting 24 h after the surgery and continue for 3 days postoperatively.

3. Collection of samples from recipient mice

1. Induce anesthesia in an induction chamber using 5% isoflurane.
2. Confirm the absence of reflex response to the toe pinch before orotracheal intubation. The intubation method and respirator setting are the same as in recipient surgery.
3. Position the mouse in a supine position and secure the limbs.
4. Prep the surgical area by sterilizing it with 70% isopropyl alcohol.
5. Make a midline incision on the skin, starting from the mid-abdomen and extending to the anterior cervical region.
6. Exsanguinate the mouse via the inferior vena cava using a 1 ml syringe connected to a 25 G needle, resulting in euthanasia.
7. Open the chest and access the trachea in the same way as a donor mouse. Tie the trachea around the intubating tube with 7-0 silk.
8. Remove the thymus, fat, and muscles to expose the heart.
9. Cut the left atrium, right atrium, and inferior vena cava. Perfuse the lungs with 3 mL of sterile saline via the right ventricle.
10. For histologic analysis, inflate the lungs with 10% formalin via an intubating tube.
11. Extubate the ventilation tube and tie off the trachea with 7-0 silk.
12. Divide the larynx and esophagus. Pull them in an inferior direction, and then extract the heart and lung block, placing it into 10% formalin.

Results

Based on our experience, proficiency in this model typically requires approximately 2 months of training. Once proficiency is achieved, the donor procedures typically require 15 min, while the recipient procedures require approximately 30 min. The expected mortality rate for a trained operator is 0%.

In Figure 4A, a tracheal allograft exhibits complete obstruction with fibroblastic tissue, and the epithelial cells are visibly destroyed. Conversely, in

Discussion

The murine IPTT procedure includes critical steps. Regarding anesthesia, the first crucial step is endotracheal intubation. It is essential to hold the mouse at an appropriate height with its legs on the table to visualize the vocal cords and facilitate immediate intubation. Additionally, careful respiratory volume and positive end-expiratory pressure (PEEP) adjustment is necessary. Typically, a respiratory volume of 500 µL and a PEEP of 2 cmH₂O are sufficient for mice weighing 25-30 g. However, larger re...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
BALB/cJ	The Jackson Laboratory	8-10 weeks 25-30 g	Male, Donor
BD 1 mL Syringe	Becton Dickinson	309659
BD PrecisionGlide Needle Aiguile BD PrecisionGlide	Becton Dickinson	305122
Bovie Change-A-Tip Deluxe High-Temperture	Bovie	DEL1
C57BL/6J	The Jackson Laboratory	8-10 weeks 25-30 g	Male, Recipient
Dumont #5/45 Forceps	F·S·T	11251-35
Ethicon Ligaclip Multiple -Clip Appliers-	Ethicon	LX107
Extra Fine Graefe Forceps	F·S·T	11150-10
Glover Bulldog Clamp	Integra	320-127
Halsted-Mosquito Hemostats	F·S·T	13009-12
Horizon Titanium Ligating Clips	Teleflex	001201
Leica M651 Manual surgical microscope for microsurgical procedures	Leica
Magnetix Fixator with spring lock	CD+ LABS	ACD-001
Microsurgical Scissor	Jarit	277-051
Mouse and Perinatal Rat Ventilator Model 687	Harvard	55-0001
Perfadex Plus	XVIVO	19850
Retractor Tip Blunt - 2.5 mm	CD+ LABS	ACD-011
small animal table	CD+ LABS	ACD-003
Surgipro Blue 24" CV-1 Taper, Double Armed	Covidien	VP702X
Systane ointment	Alconn	1444062
System Elastomer	CD+ LABS	ACD-007
Terumo Surflo IV Catheter, 20 G x 1 in	Terumo Medical Corporation	SR-OX2025CA
VMT table Top	benson	91803300