The authors would like to thank Jerome Valero for editing this manuscript. Figure 1 and Figure 3I,J,L were created with BioRender.com.

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.
<ol>
	<li>Prior to the procedure, record the weight of each mouse.</li>
	<li>Euthanize the mouse using a CO2 chamber.</li>
	<li>Once death is confirmed, position the mouse in a supine position and secure the limbs with tape.</li>
	<li>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.</li>
	<li>Make a midline incision on the skin, starting from the mid-abdomen and extending to the anterior cervical region.</li>
	<li>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.</li>
	<li>Lift the xiphoid and cut the diaphragm.</li>
	<li>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.</li>
	<li>Remove the thymus and any fat or muscle obstructing the trachea to expose the tracheal bifurcation.</li>
	<li>Cut both the main bronchi and carefully separate the airway from the esophagus.</li>
	<li>Cut the larynx and remove it.</li>
	<li>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.</li>
</ol>
2. Recipient surgery
NOTE: C57BL/6 mice are used as an example of recipients for the experiment.
<ol>
	<li>Administer sustained-release buprenorphine subcutaneously at a dose of 1 mg/kg on the morning of the surgery day.</li>
	<li>Induce anesthesia in an induction chamber using 5% isoflurane.</li>
	<li>Once the mouse is lightly anesthetized, intraperitoneally inject a cocktail consisting of (0.1 mg/g) xylazine and (0.01 mg/g) ketamine.</li>
	<li>Return the mouse to the induction chamber with 2-3% isoflurane maintained.</li>
	<li>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.</li>
	<li>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 &#181;L, 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.</li>
	<li>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.</li>
	<li>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.&#160;&#160;</li>
	<li>Load the donor trachea into a 16 G intravenous catheter during this time (Figure 3C,D).</li>
	<li>Use scalpel to make an incision in the recipient&#39;s skin and cauterize the muscle and connective tissue.</li>
	<li>Open the fifth or sixth intercostal space and hold the rib cage open using two retractors.</li>
	<li>Dissect the inferior pulmonary ligament using cotton swabs and scissors.</li>
	<li>Simulate the creation of the pathway for the donor trachea (Figure 3G,H).</li>
	<li>Secure the ventilator outflow tube by partially occluding it with a three-way stopcock to facilitate inflation of the left lung.</li>
	<li>Create a pathway by puncturing the left lung with a 20&#160;G needle. Ensuring that the puncture depth is roughly equivalent to the length of the tracheal allograft. Select the puncture site at the lung&#39;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).</li>
	<li>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.</li>
	<li>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&#39;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).</li>
	<li>Fill the thoracic cavity with saline solution and absorb the saline with a gauze.</li>
	<li>Reinflate the left lung and close the ribs using a running suture technique.</li>
	<li>Close the muscle and skin with interrupted sutures.</li>
	<li>Administer meloxicam analgesic subcutaneously at a dose of 5 mg/kg at the end of the surgery.</li>
	<li>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.</li>
	<li>Administer meloxicam (5 mg/kg) once daily via subcutaneous injection, starting 24 h after the surgery and continue for 3 days postoperatively.</li>
</ol>
3. Collection of samples from recipient mice
<ol>
	<li>Induce anesthesia in an induction chamber using 5% isoflurane.</li>
	<li>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.</li>
	<li>Position the mouse in a supine position and secure the limbs.</li>
	<li>Prep the surgical area by sterilizing it with 70% isopropyl alcohol.</li>
	<li>Make a midline incision on the skin, starting from the mid-abdomen and extending to the anterior cervical region.</li>
	<li>Exsanguinate the mouse via the inferior vena cava using a 1 ml syringe connected to a 25 G needle, resulting in euthanasia.</li>
	<li>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.</li>
	<li>Remove the thymus, fat, and muscles to expose the heart.</li>
	<li>Cut the left atrium, right atrium, and inferior vena cava. Perfuse the lungs with 3 mL of sterile saline via the right ventricle.</li>
	<li>For histologic analysis, inflate the lungs with 10% formalin via an intubating tube.</li>
	<li>Extubate the ventilation tube and tie off the trachea with 7-0 silk.</li>
	<li>Divide the larynx and esophagus. Pull them in an inferior direction, and then extract the heart and lung block, placing it into 10% formalin.</li>
</ol>

Exploring Obliterative Airway Disease in Lung Transplants

<ol>
	<li>Chambers, D. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+International+Thoracic+Organ+Transplant+Registry+of+the+International+Society+for+Heart+and+Lung+Transplantation:+Thirty-fifth+adult+lung+and+heart-lung+transplant+report-2018;+Focus+theme:+Multiorgan+Transplantation.">The International Thoracic Organ Transplant Registry of the International Society for Heart and Lung Transplantation: Thirty-fifth adult lung and heart-lung transplant report-2018; Focus theme: Multiorgan Transplantation.</a> J Heart Lung Transplant. 37 (10), 1169-1183 (2018).</li><li>Hertz, M. I., Jessurun, J., King, M. B., Savik, S. K., Murray, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reproduction+of+the+obliterative+bronchiolitis+lesion+after+heterotopic+transplantation+of+mouse+airways.">Reproduction of the obliterative bronchiolitis lesion after heterotopic transplantation of mouse airways.</a> American J Pathol. 142 (6), 1945-1951 (1993).</li><li>Ikonen, T. S., Brazelton, T. R., Berry, G. J., Shorthouse, R. S., Morris, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epithelial+re-growth+is+associated+with+inhibition+of+obliterative+airway+disease+in+orthotopic+tracheal+allografts+in+non-immunosuppressed+rats.">Epithelial re-growth is associated with inhibition of obliterative airway disease in orthotopic tracheal allografts in non-immunosuppressed rats.</a> Transplantation. 70 (6), 857 (2000).</li><li>Dutly, A. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+model+for+post-transplant+obliterative+airway+disease+reveals+angiogenesis+from+the+pulmonary+circulation.">A novel model for post-transplant obliterative airway disease reveals angiogenesis from the pulmonary circulation.</a> Am J Transplant. 5 (2), 248-254 (2005).</li><li>Wagnetz, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rejection+of+tracheal+allograft+by+intrapulmonary+lymphoid+neogenesis+in+the+absence+of+secondary+lymphoid+organs.">Rejection of tracheal allograft by intrapulmonary lymphoid neogenesis in the absence of secondary lymphoid organs.</a> Transplantation. 93 (12), 1212-1220 (2012).</li><li>Hirayama, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Local+long-term+expression+of+lentivirally+delivered+IL-10+in+the+lung+attenuates+obliteration+of+intrapulmonary+allograft+airways.">Local long-term expression of lentivirally delivered IL-10 in the lung attenuates obliteration of intrapulmonary allograft airways.</a> Hum Gene Ther. 22 (11), 1453-1460 (2011).</li><li>Watanabe, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recipient+bone+marrow-derived+IL-17+receptor+A-positive+cells+drive+allograft+fibrosis+in+a+mouse+intrapulmonary+tracheal+transplantation+model.">Recipient bone marrow-derived IL-17 receptor A-positive cells drive allograft fibrosis in a mouse intrapulmonary tracheal transplantation model.</a> Transpl Immunol. 69, 101467 (2021).</li><li>Matsuda, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spleen+tyrosine+kinase+modulates+fibrous+airway+obliteration+and+associated+lymphoid+neogenesis+after+transplantation.">Spleen tyrosine kinase modulates fibrous airway obliteration and associated lymphoid neogenesis after transplantation.</a> Am J Transplant. 16 (1), 342-352 (2016).</li><li>Suzuki, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+CTLA4-Ig+on+Obliterative+bronchiolitis+in+a+mouse+intrapulmonary+tracheal+transplantation+model.">Effect of CTLA4-Ig on Obliterative bronchiolitis in a mouse intrapulmonary tracheal transplantation model.</a> Ann Thorac Cardiovasc Surg. 27 (6), 355-365 (2021).</li><li>Watanabe, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+potent+anti-angiogenic+factor,+vasohibin-1,+ameliorates+experimental+bronchiolitis+obliterans.">A potent anti-angiogenic factor, vasohibin-1, ameliorates experimental bronchiolitis obliterans.</a> Transplant Proc. 44 (4), 1155-1157 (2012).</li><li>Aloisi, F., Pujol-Borrell, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lymphoid+neogenesis+in+chronic+inflammatory+diseases.">Lymphoid neogenesis in chronic inflammatory diseases.</a> Nat Rev Immunol. 6 (3), 205-217 (2006).</li><li>Cupedo, T., Jansen, W., Kraal, G., Mebius, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+of+secondary+and+tertiary+lymphoid+structures+in+the+skin.">Induction of secondary and tertiary lymphoid structures in the skin.</a> Immunity. 21 (5), 655-667 (2004).</li><li>Sato, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+intrapulmonary+de+novo+lymphoid+tissue+in+obliterative+bronchiolitis+after+lung+transplantation.">The role of intrapulmonary de novo lymphoid tissue in obliterative bronchiolitis after lung transplantation.</a> J Immunol. 182 (11), 7307-7316 (2009).</li><li>Okazaki, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maintenance+of+airway+epithelium+in+acutely+rejection+orthotopic+vascularized+mouse+lung+transplants.">Maintenance of airway epithelium in acutely rejection orthotopic vascularized mouse lung transplants.</a> A J Resp Cell Mol Biol. 37 (6), 625-630 (2007).</li><li>Yamada, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+airway+fibrosis+in+orthotopic+mouse+lung+transplantation+models-an+experimental+reappraisal.">Chronic airway fibrosis in orthotopic mouse lung transplantation models-an experimental reappraisal.</a> Transplantation. 102 (2), e49-e58 (2018).</li><li>Watanabe, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+B+cell-dependent+pathway+drives+chronic+lung+allograft+rejection+after+ischemia-reperfusion+injury+in+mice.">A B cell-dependent pathway drives chronic lung allograft rejection after ischemia-reperfusion injury in mice.</a> Am J Transplant. 19 (12), 3377-3389 (2019).</li><li>Guo, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vendor-specific+microbiome+controls+both+acute+and+chronic+murine+lung+allograft+rejection+by+altering+CD4+Foxp3++regulatory+T+cell+levels.">Vendor-specific microbiome controls both acute and chronic murine lung allograft rejection by altering CD4+Foxp3+ regulatory T cell levels.</a> Am J Transplant. 19 (10), 2705-2718 (2019).</li></ol>

The authors of this manuscript have no conflicts of interest to disclose.

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 &#181;L and a PEEP of 2 cmH2O are sufficient for mice weighing 25-30 g. However, larger re...

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&#39;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&#39;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 IPTT5,6,7,8,9,10. 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 obliteration4,10. 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 cells11,12. 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 obliteration7,8,11,12,13. 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.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>BALB/cJ</td>
			<td>The Jackson Laboratory</td>
			<td>8-10 weeks 25-30 g</td>
			<td>Male, Donor</td>
		</tr>
		<tr>
			<td>BD 1 mL Syringe</td>
			<td>Becton Dickinson</td>
			<td>309659</td>
			<td></td>
		</tr>
		<tr>
			<td>BD PrecisionGlide Needle Aiguile BD 
			PrecisionGlide</td>
			<td>Becton Dickinson</td>
			<td>305122</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovie Change-A-Tip Deluxe High-Temperture</td>
			<td>Bovie</td>
			<td>DEL1</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6J</td>
			<td>The Jackson Laboratory</td>
			<td>8-10 weeks 25-30 g</td>
			<td>Male, Recipient</td>
		</tr>
		<tr>
			<td>Dumont #5/45 Forceps</td>
			<td>F&#183;S&#183;T</td>
			<td>11251-35</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethicon Ligaclip Multiple -Clip Appliers-</td>
			<td>Ethicon</td>
			<td>LX107</td>
			<td></td>
		</tr>
		<tr>
			<td>Extra Fine Graefe Forceps</td>
			<td>F&#183;S&#183;T</td>
			<td>11150-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Glover Bulldog Clamp</td>
			<td>Integra</td>
			<td>320-127</td>
			<td></td>
		</tr>
		<tr>
			<td>Halsted-Mosquito Hemostats</td>
			<td>F&#183;S&#183;T</td>
			<td>13009-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Horizon Titanium Ligating Clips</td>
			<td>Teleflex</td>
			<td>001201</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica M651 Manual surgical microscope for microsurgical procedures</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Magnetix Fixator with spring lock</td>
			<td>CD+ LABS</td>
			<td>ACD-001</td>
			<td></td>
		</tr>
		<tr>
			<td>Microsurgical Scissor</td>
			<td>Jarit</td>
			<td>277-051</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse and Perinatal Rat Ventilator Model 687</td>
			<td>Harvard</td>
			<td>55-0001</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfadex Plus</td>
			<td>XVIVO</td>
			<td>19850</td>
			<td></td>
		</tr>
		<tr>
			<td>Retractor Tip Blunt - 2.5 mm</td>
			<td>CD+ LABS</td>
			<td>ACD-011</td>
			<td></td>
		</tr>
		<tr>
			<td>small animal table</td>
			<td>CD+ LABS</td>
			<td>ACD-003</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgipro Blue 24&#34; CV-1 Taper, Double Armed</td>
			<td>Covidien</td>
			<td>VP702X</td>
			<td></td>
		</tr>
		<tr>
			<td>Systane ointment</td>
			<td>Alconn</td>
			<td>1444062</td>
			<td></td>
		</tr>
		<tr>
			<td>System Elastomer</td>
			<td>CD+ LABS</td>
			<td>ACD-007</td>
			<td></td>
		</tr>
		<tr>
			<td>Terumo Surflo IV Catheter, 20 G x 1 in</td>
			<td>Terumo Medical Corporation</td>
			<td>SR-OX2025CA</td>
			<td></td>
		</tr>
		<tr>
			<td>VMT table Top</td>
			<td>benson</td>
			<td>91803300</td>
			<td></td>
		</tr>
	</tbody>
</table>

murine intrapulmonary tracheal transplantation: a model for investigating obliterative airway disease after lung transplantation

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.

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 <strong clas...

Watch this Scientific Journal Video about Author Spotlight: Investigating the Key Factors of Obliterative Bronchiolitis After Lung Transplantation at JoVE.com

Author Spotlight: Investigating the Key Factors of Obliterative Bronchiolitis After Lung Transplantation

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.

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

Murine intrapulmonary tracheal transplantation (IPTT) is used as a model of obliterative airway disease (OAD) following lung transplantation. Initially ...

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768 Views.  University Health Network. 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.

Video: Author Spotlight: Investigating the Key Factors of Obliterative Bronchiolitis After Lung Transplantation

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

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

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

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

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are available to perform mechanistic in vivo studies. While heart transplantation models using a suture technique were first successfully developed in rats, the translation into an equally widespread used murine equivalent was never achieved due the technical complexity of the microsurgical procedure. In contrast, non-suture cuff techniques, also developed initially in rats, were successfully adapted for use in mice1-3. This technique for revascularization involves two major steps I) everting the recipient vessel over a polyethylene cuff; II) pulling the donor vessel over the formerly everted recipient vessel and holding it in place with a circumferential tie. This ensures a continuity of the endothelial layer, short operating time and very high patency rates4.
Using this technique for vascular anastomosis we performed more than 1,000 cervical heart transplants with an overall success rate of 95%. For arterial inflow the common carotid artery and the proximal aortic arch were anastomosed resulting in a retrograde perfusion of the transplanted heart. For venous drainage the pulmonary artery of the graft was anastomosed with the external jugular vein of the recipient5.
Herein, we provide additional details of this technique to supplement the video.

The murine cervical heart transplantation model is well suited for immunological as well as ischemia reperfusion injury studies. We modified the procedure using a non-suture cuff technique and performed more than 1,000 successful transplants with this approach.
Herein, we provide additional details of this technique to supplement the video.

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are ...

Murine Heterotopic Heart Transplant Technique

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become proficient in and utilize this technique, it is now widely used by many laboratories around the world. A significant refinement to the original technique was developed and reported in 2001 by Niimi. Described here are the techniques that have evolved over more than a decade in the hands of three surgeons (Plenter, Grazia, Pietra) in our center. These techniques are now being passed on to a younger generation of surgeons and researchers.
Based largely on the Niimi experience, the procedures used have evolved in the fine details - details which we will endeavor to relate here in such a way that others may be able to use this very useful model. Like Niimi, we have found that a video aid to learning is a priceless resource for the beginner.

The manuscript describes the steps required to perform the heterotopic heart transplant in the mouse.

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become ...

Murine Corneal Transplantation: A Model to Study the Most Common Form of Solid Organ Transplantation

Corneal transplantation is the most common form of organ transplantation in the United States with between 45,000 and 55,000 procedures performed each year. While several animal models exist for this procedure and mice are the species that is most commonly used. The reasons for using mice are the relative cost of using this species, the existence of many genetically defined strains that allow for the study of immune responses, and the existence of an extensive array of reagents that can be used to further define responses in this species. This model has been used to define factors in the cornea that are responsible for the relative immune privilege status of this tissue that enables corneal allografts to survive acute rejection in the absence of immunosuppressive therapy. It has also been used to define those factors that are most important in rejection of such allografts. Consequently, much of what we know concerning mechanisms of both corneal allograft acceptance and rejection are due to studies using a murine model of corneal transplantation. In addition to describing a model for acute corneal allograft rejection, we also present for the first time a model of late-term corneal allograft rejection.

Mice have been used as a model for studying many forms of transplantation, including corneal transplantation. We describe in this report a murine model for both acute and late-term corneal transplantation.

Corneal transplantation is the most common form of organ transplantation in the United States with between 45,000 and 55,000 procedures performed each ...

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. As this technology expands and improves, the ability to potentially evaluate and improve the quality of substandard lungs prior to transplant is a critical need. In order to more rigorously evaluate these approaches, a reproducible animal model needs to be established that would allow for testing of improved techniques and management of the donated lungs as well as to the lung-transplant recipient. In addition, an EVLP animal model of associated pathologies, e.g., ventilation induced lung injury (VILI), would provide a novel method to evaluate treatments for these pathologies. Here, we describe the development of a rat EVLP lung program and refinements to this method that allow for a reproducible model for future expansion. We also describe the application of this EVLP system to model VILI in rat lungs. The goal is to provide the research community with key information and &#8220;pearls of wisdom&#8221;/techniques that arose from trial and error and are critical to establishing an EVLP system that is robust and reproducible.

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. Here, we describe the development of a rat EVLP program and refinements that allow for a reproducible model for future expansion.

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed ...

Cardiac Catheterization in Mice to Measure the Pressure Volume Relationship: Investigating the Bowditch Effect

Animal models that mimic human cardiac disorders have been created to test potential therapeutic strategies. A key component to evaluating these strategies is to examine their effects on heart function. There are several techniques to measure in vivo cardiac mechanics (e.g., echocardiography, pressure/volume relations, etc.). Compared to echocardiography, real-time left ventricular (LV) pressure/volume analysis via catheterization is more precise and insightful in assessing LV function. Additionally, LV pressure/volume analysis provides the ability to instantaneously record changes during manipulations of contractility (e.g., &#946;-adrenergic stimulation) and pathological insults (e.g., ischemia/reperfusion injury). In addition to the maximum (+dP/dt) and minimum (-dP/dt) rate of pressure change in the LV, an accurate assessment of LV function via several load-independent indexes (e.g., end systolic pressure volume relationship and preload recruitable stroke work) can be attained. Heart rate has a significant effect on LV contractility such that an increase in the heart rate is the primary mechanism to increase cardiac output (i.e., Bowditch effect). Thus, when comparing hemodynamics between experimental groups, it is necessary to have similar heart rates. Furthermore, a hallmark of many cardiomyopathy models is a decrease in contractile reserve (i.e., decreased Bowditch effect). Consequently, vital information can be obtained by determining the effects of increasing heart rate on contractility. Our and others data has demonstrated that the neuronal nitric oxide synthase (NOS1) knockout mouse has decreased contractility. Here we describe the procedure of measuring LV pressure/volume with increasing heart rates using the NOS1 knockout mouse model.

This article describes the measurement of murine left ventricular function via pressure/volume analysis at different heart rates.

Animal models that mimic human cardiac disorders have been created to test potential therapeutic strategies. A key component to evaluating these ...

Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. However, only minor improvement in the long-term results of transplanted solid organs could be observed over the past decades. In this context, chronic allograft vasculopathy (CAV) still represents the leading cause of late organ failure in cardiac, renal and pulmonary transplantation.
Thus far, the underlying pathogenesis of CAV development remains unclear, explaining why effective treatment strategies are presently missing and emphasizing a need for relevant experimental models in order to study the underlying pathophysiology leading to CAV formation. The following protocol describes a murine heterotopic cervical aortic transplantation model using a modified non-suture cuff technique. In this technique, a segment of the thoracic aorta is interpositioned in the right common carotid artery. With the use of the non-suture cuff technique, an easy to learn and reproducible model can be established, minimizing the possible heterogeneity of sutured vascular micro anastomoses.

Here, we present a protocol of heterotopic aortic transplantation in mice using the non-suture cuff technique in a cervical murine model. This model can be used to study the underlying pathology of chronic allograft vasculopathy (CAV) and can help evaluate new therapeutic agents in order to prevent its formation.

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. ...

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

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

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

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

Inguinal Subcutaneous White Adipose Tissue (ISWAT) Transplantation Model of Murine Islets

Pancreatic islet transplantation is a well-established therapeutic treatment for type 1 diabetes. The kidney capsule is the most commonly used site for islet transplantation in rodent models. However, the tight kidney capsule limits the transplantation of sufficient islets in large animals and humans. The inguinal subcutaneous white adipose tissue (ISWAT), a new subcutaneous space, was found to be a potentially valuable site for islet transplantation. This site has better blood supply than other subcutaneous spaces. Moreover, the ISWAT accommodates a larger islet mass than the kidney capsule, and transplantation into it is simple. This manuscript describes the procedure of mouse islet isolation and transplantation in the ISWAT site of syngeneic diabetic mouse recipients. Using this protocol, murine pancreatic islets were isolated by standard collagenase digestion and a basement membrane matrix hydrogel was used for fixing the purified islets in the ISWAT site. The blood glucose levels of the recipient mice were monitored for more than 100 days. Islet grafts were retrieved at day 100 after transplantation for histological analysis. The protocol for islet transplantation in the ISWAT site described in this manuscript is simple and effective.

Inguinal Subcutaneous White Adipose Tissue ISWAT Transplantation Model of Murine Islets

In this protocol, a method of murine islet isolation and transplantation into the inguinal subcutaneous white adipose tissue is described. Isolated syngeneic murine islets are transplanted into a murine recipient using a basement membrane hydrogel. The blood glucose level of the recipients is monitored, and histology analysis of the islet grafts is performed.

Pancreatic islet transplantation is a well-established therapeutic treatment for type 1 diabetes. The kidney capsule is the most commonly used site for ...

Intramyocardial Transplantation of MSC-Loading Injectable Hydrogels after Myocardial Infarction in a Murine Model

One of the major issues facing current cardiac stem cell therapies for preventing postinfarct heart failure is the low retention and survival rates of transplanted cells within the injured myocardium, limiting their therapeutic efficacy. Recently, the use of scaffolding biomaterials has gained attention for improving and maximizing stem cell therapy. The objective of this protocol is to introduce a simple and straightforward technique to transplant bone marrow-derived mesenchymal stem cells (MSCs) using injectable hydroxyphenyl propionic acid (GH) hydrogels; the hydrogels are favorable as a cell delivery platform for cardiac tissue engineering applications due to their ability to be cross-linked in situ and high biocompatibility. We present a simple method to fabricate MSC-loading GH hydrogels (MSC/hydrogels) and evaluate their survival and proliferation in three-dimensional (3D) in vitro culture. In addition, we demonstrate a technique for intramyocardial transplantation of MSC/hydrogels in mice, describing a surgical procedure to induce myocardial infarction (MI) via left anterior descending (LAD) coronary artery ligation and subsequent MSC/hydrogels transplantation.

Stem cell-based therapy has emerged as an efficient strategy to repair injured cardiac tissues after myocardial infarction. We provide an optimal in vivo application for stem cell transplantation using gelatin hydrogels that are able to be enzymatically cross-linked.

One of the major issues facing current cardiac stem cell therapies for preventing postinfarct heart failure is the low retention and survival rates of ...