Name: murine intrapulmonary tracheal transplantation: a model for investigating obliterative airway disease after lung transplantation
Uploaded: 2023-11-10T12:40:00.000+00:00
Duration: 6 min 15 s
Description: Watch this Scientific Journal Video about Author Spotlight: Investigating the Key Factors of Obliterative Bronchiolitis After Lung Transplantation at JoVE.com

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

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	<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 recipient mice may experience &#34;hiccup&#34; episodes, indicating oxygen deficiency. In such cases, it may be necessary to increase the respiratory volume, PEEP, and temporarily clamp the expiratory tube to recruit the lung. If hiccups persist, the depth of the tracheal tube should be adjusted. Lastly, the timing of tracheal tube extubation is critical, and confirmation of the recipient&#39;s respiratory rate is essential to ensure sufficient recovery. Failure to do so may result in postextubation mortality.
Regarding the surgical procedure, proper placement of the transplanted trachea is crucial. Simulating the appropriate needle direction is necessary to accurately position the tracheal graft (Figure 3G). If the path is too shallow, the tracheal graft may penetrate the pleura, while inserting it too deep may lead to lung vessel perforation and severe bleeding. Another critical step is ensuring accurate sealing of the insertion point by securely fixing the pleural incision into which the tracheal allograft was inserted with a stainless steel clip (Figure 3H). Insufficient sealing can cause air leakage, potentially resulting in recipient mortality after chest closure.
While the IPTT model offers some advantages, such as relative simplicity, high reproducibility, and an immunological environment similar to that seen in lung transplantation, it does have some limitations. Tracheal allografts in the IPTT model differ from the clinical situation, where OB occurs in the small airways. Moreover, in this major histocompatibility complex (MHC) mismatch IPTT model, the tracheal allograft is completely occluded with fibroblastic tissue, which is in contrast with clinical OAD where fibroblastic tissue occlusion is usually partial. To address this issue, another strain combination (minor or different major mismatch) might be used for IPTT (depending on the intended study), as it could result in a milder alloimmune response and milder fibroblastic tissue occlusion compared to the BALB/c-to-C57BL/6 combination. We are unaware of any published studies examining different strain combinations, so this possibility would need to be tested empirically.
Finally, it is a nonvascularized graft, similar to the HTT and OTT models. To address this limitation, orthotopic lung transplantation (OLT) can be utilized14. However, the development of OAD-like pathology has been variable in the mouse OLT model15,16,17. Given the IPTT model&#39;s relative simplicity and reproducibility compared to OLT, it remains a reasonable model to study the mechanisms of transplant-related OAD. In conclusion, the IPTT model serves as a useful research model for studying intrapulmonary immune and profibrotic pathways involved in the development of airway obliteration after lung transplantation.

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><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&lt;br/&gt; 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&amp;middot;S&amp;middot;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&amp;middot;S&amp;middot;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&amp;middot;S&amp;middot;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&quot; 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 Figure 4B, a tracheal isograft remains patent, and the epithelial cells are preserved.
Figure 5 illustrates a lung in which the tracheal allograft has been transplanted, showing the presence of lymphoid aggregates.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65953/65953fig01.jpg" /> 
Figure 1: Diagram of murine intrapulmonary tracheal transplantation model. A tracheal allograft is extracted from a donor mouse. The tracheal allograft is loaded into a catheter. The tracheal allograft is transplanted to the lung of a recipient. <a href="https://www.jove.com/files/ftp_upload/65953/65953fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65953/65953fig02.jpg" /> 
Figure 2: Position of the recipient during surgery. The mouse is placed in a right lateral decubitus position. The recipient mouse&#39;s head is oriented away from the surgeon and the tail faces the surgeon. <a href="https://www.jove.com/files/ftp_upload/65953/65953fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65953/65953fig03.jpg" /> 
Figure 3: Catheter for tracheal insertion and illustration of the direction of the insertion of the tracheal graft. (A) The catheter for tracheal insertion. The outer catheter has a sharp tip (left), while the inner catheter has a blunt tip (right). (B) Combination of outer and inner catheters. The inner catheter protrudes slightly from the outer catheter. (C,D) Loading the donor trachea into the catheter. The red bracket indicates the tracheal graft. (E) Tracheal graft inside the catheter. The red bracket indicates the tracheal graft. (F) Extruding the inner tracheal graft using the inner catheter as a &#34;pusher.&#34; The red bracket indicates the tracheal graft. (G, H) Simulation of the direction for placement of the tracheal graft. (I,J) Creating the pathway using a 22 G needle. The depth should closely match the length of a tracheal allograft. The needle&#39;s direction should be opposite from the surgeon and parallel to the tabletop. The correct puncture site is denoted by the red point. The appropriate insertion angle is indicated by the blue circle. Incorrect angles are indicated by red crosses. (K,L) Closing the pleural injection site with a clip. Black lines represent clips. Red point represents the puncture site. The blue circle indicates the correct clipping point. Red crosses indicate incorrect clipping points. <a href="https://www.jove.com/files/ftp_upload/65953/65953fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65953/65953fig04.jpg" /> 
Figure 4: Histopathology at 28 days post lung transplantation. (A) H&#38;E (left) and Masson&#39;s Trichrome (right) stained images of a tracheal allograft (Donor: BALB/c, Recipient: C57BL/6). The lumen of the allograft is occluded with collagen and fibrous tissue stained blue by Masson&#39;s Trichrome (black arrowhead). Additionally, the epithelial cells have been lost (blue arrowheads). (B) H&#38;E (left) and Masson&#39;s Trichrome (right) stained images of a tracheal isograft. In contrast to the allograft, the lumen of the isograft (Donor, Recipient: C57BL/6) remains open, and the epithelial cells are preserved. Stained tissue is mucus. Scale bars = 500 &#181;m. Abbreviation: H&#38;E = hematoxylin and eosin. <a href="https://www.jove.com/files/ftp_upload/65953/65953fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65953/65953fig05.jpg" /> 
Figure 5: Lymphoid aggregates in the lung with the transplanted tracheal allograft. (A) H&#38;E stained image of a lung with a transplanted tracheal allograft in situ. Lymphoid aggregates (black arrows) are observed within the lung tissue. Scale bar = 500 &#181;m. (B) Immunofluorescence image of a lymphoid aggregate, highlighting the presence of B cell (B220, red), T cell (CD3, green), and nuclei (DAPI, blue). Scale bar = 100 &#181;m. Abbreviations: H&#38;E = hematoxylin and eosin; DAPI = 4&#39;,6-diamidino-2-phenylindole. <a href="https://www.jove.com/files/ftp_upload/65953/65953fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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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Nanyang Technological University

Research

JoVE Journal

Medicine

790 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 ...

Noninvasive Intratracheal Intubation to Study the Pathology and Physiology of Mouse Lung

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular disease and for developing therapeutic intervention. With the increasing availability of knockout and transgenic derivatives, together with a vast amount of genetic information, mice provide one of the best models to study the molecular mechanisms underlying the pathology and physiology of lung diseases. Inhalation, intranasal instillation, intratracheal instillation, and intratracheal intubation are the most widely used techniques by a number of investigators to administer materials of interest to mouse lungs. There are pros and cons for each technique depending on the goals of a study. Here a noninvasive intratracheal intubation method that can directly deliver exogenous materials to mouse lungs is presented. This technique was applied to administer bleomycin to mouse lungs as a model to study pulmonary fibrosis.

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular disease and for developing therapeutic intervention. Here a noninvasive intratracheal intubation method that can directly deliver exogenous materials to mouse lungs is presented.

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular ...

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 ...

Heterotopic and Orthotopic Tracheal Transplantation in Mice used as Models to Study the Development of  Obliterative Airway Disease

Obliterative airway disease (OAD) is the major complication after lung transplantations that limits long term survival (1-7).
To study the pathophysiology, treatment and prevention of OAD, different animal models of tracheal transplantation in rodents have been developed (1-7). Here, we use two established models of trachea transplantation, the heterotopic and orthotopic model and demonstrate their advantages and limitations.
For the heterotopic model, the donor trachea is wrapped into the greater omentum of the recipient, whereas the donor trachea is anastomosed by end-to-end anastomosis in the orthotopic model.
In both models, the development of obliterative lesions histological similar to clinical OAD has been demonstrated (1-7).
This video shows how to perform both, the heterotopic as well as the orthotopic tracheal transplantation technique in mice, and compares the time course of OAD development in both models using histology.

This video shows and compares two experimental models to study the development of obliterative airway disease (OAD) in mice, the heterotopic and orthotopic tracheal transplantation model.

Obliterative airway disease (OAD) is the major complication after lung transplantations that limits long term survival (1-7).
To study the ...

Biology

An Improved Method for Rapid Intubation of the Trachea in Mice

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity is a commonly used model to study both acute lung injury and fibrosis, and multiple methods have been developed for administering bleomycin (and other toxic agents) into the lungs. However, many of these approaches, such as transtracheal instillation, have inherent drawbacks, including the need for strong anesthetics and survival surgery. This paper reports a quick, reproducible method of intratracheal intubation that involves mild inhaled anesthesia, visualization of the trachea, and the use of a surrogate spirometer to confirm exposure. As a proof of concept, 8-12 week old C57BL/6 mice were administered either 2.0 U/kg of bleomycin or an equivalent volume of PBS, and both damage and fibrotic endpoints were measured post-exposure. This procedure allows researchers to treat a large cohort of mice in a relatively short period with little expense and minimal post-procedure care.

This article presents a rapid and simple method for administering bleomycin directly into the mouse trachea via intubation. Key advantages of this method are that it is highly reproducible, easy to master, and does not require specialized equipment or lengthy recovery times.

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity ...

Seeding and Implantation of a Biosynthetic Tissue-engineered Tracheal Graft in a Mouse Model

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional tissue. Tissue engineering holds great promise as a potential solution with its ability to integrate cells and signaling molecules into a 3-dimensional scaffold. Recent work with tissue engineered tracheal grafts (TETGs) has seen some success but their translation has been limited by graft stenosis, graft collapse, and delayed epithelialization. In order to investigate the mechanisms driving these issues, we have developed a mouse model for tissue engineered tracheal graft implantation. TETGs were constructed using electrospun polymers polyethylene terephthalate (PET) and polyurethane (PU) in a mixture of PET and PU (20:80 percent weight). Scaffolds were then seeded using bone marrow mononuclear cells isolated from 6-8 week-old C57BL/6 mice by gradient centrifugation. Ten million cells per graft were seeded onto the lumen of the scaffold and allowed to incubate overnight before implantation between the third and seventh tracheal rings. These grafts were able to recapitulate the findings of stenosis and delayed epithelialization as demonstrated by histological analysis and lack of Keratin 5 and Keratin 14 basal epithelial cells on immunofluorescence. This model will serve as a tool for investigating cellular and molecular mechanisms involved in host remodeling.

Graft stenosis poses a critical obstacle in tissue engineered airway replacement. To investigate cellular mechanisms underlying stenosis, we utilize a murine model of tissue engineered tracheal replacement with seeded bone marrow mononuclear cells (BM-MNC). Here, we detail our protocol, including scaffold manufacturing, BM-MNC isolation, graft seeding, and implantation.

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional ...

Bioengineering

A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR permits the experimental study of this specific process with continued alveolar ventilation, thereby avoiding the compounding injurious processes of hypoxia and atelectasis. In the clinical context, lung ischemia reperfusion injury (also known as lung IRI or LIRI) is caused by numerous processes, including but not limited to pulmonary embolism, resuscitated hemorrhagic trauma, and lung transplantation. There are currently limited effective treatment options for LIRI. Here, we present a reversible surgical model of lung IR involving first orotracheal intubation followed by unilateral left lung ischemia and reperfusion with preserved alveolar ventilation or gas exchange. Mice undergo a left thoracotomy, through which the left pulmonary artery is exposed, visualized, isolated, and compressed using a reversible slipknot. The surgical incision is then closed during the ischemic period, and the animal is awakened and extubated. With the mouse spontaneously breathing, reperfusion is established by releasing the slipknot around the pulmonary artery. This clinically relevant survival model permits the evaluation of lung IR injury, the resolution phase, downstream effects on lung function, as well as two-hit models involving experimental pneumonia. While technically challenging, this model can be mastered over the course of a few weeks to months with an eventual survival or success rate of 80%-90%.

A mouse surgical model to create left lung ischemia reperfusion (IR) injury while maintaining ventilation and avoiding hypoxia.

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR ...

Immunology and Infection

A Heterotopic Mouse Model for Studying Laryngeal Transplantation

Laryngeal heterotopic transplantation, although a technically challenging procedure, offers more scientific analysis and cost benefits compared to other animal models. Although first described by Shipchandler et al. in 2009, this technique is not widely used, possibly due to the difficulties in learning the microsurgical technique and time required to master it. This paper describes the surgical steps in detail, as well as potential pitfalls to avoid, in order to encourage effective use of this technique.
In this model, the bilateral carotid arteries of the donor larynx are anastomosed to the recipient carotid artery and external jugular vein, allowing for blood flow through the graft. Blood flow can be confirmed intraoperatively by the visualization of blood filling in the graft bilateral carotid arteries, reddening of the thyroid glands of the graft, and bleeding from micro vessels in the graft. The crucial elements for success include delicate preservation of the graft vessels, making the correct size arteriotomy and venotomy, and using the appropriate number of sutures on the arterial-arterial and arterial-venous anastomoses to secure vessels without leakage and prevent occlusion. 
Anyone can become proficient in this model with sufficient training and perform the procedure in approximately 3 h. If performed successfully, this model allows for immunologic studies to be performed with ease and at low cost.

The aim of this manuscript is to describe the microsurgical steps required to perform a heterotopic laryngeal transplant in mice. The advantages of this mouse model compared to other animal models of laryngeal transplantation are its cost-effectiveness and the availability of immunologic assays and data.

Laryngeal heterotopic transplantation, although a technically challenging procedure, offers more scientific analysis and cost benefits compared to other ...

A Simple Cuff Technique for Murine Left Lung Transplantation

Over the past decade, our laboratory has made significant progress in developing and refining vascularized mouse lung transplantation models using an efficient and highly reliable "cuff technique" of transplantation. This article describes a sophisticated and comprehensive method for orthotopic lung transplantation in a vascularized orthotopic lung model, representing the most physiologic and clinically relevant model of mouse lung transplantation to date. The transplantation process consists of two distinct stages: donor harvest and subsequent implantation into the recipient. The method has been successfully mastered, and with several months of sufficient training, a skilled practitioner can perform the procedure in approximately 90 min from skin-to-skin. Surprisingly, once individuals overcome the initial learning curve, the survival rate during the perioperative period approaches nearly 100%. The mouse model allows for the use of multiple commercially available transgenic and mutant strains of mice, enabling the study of tolerance and rejection. Additionally, the unique features of this model make it a valuable tool for investigating tumor biology and immunology.

The present protocol describes a cuff technique for a mouse left lung transplantation model. This technique has been developed over several years and has performed well, serving effectively in immunological research.

Over the past decade, our laboratory has made significant progress in developing and refining vascularized mouse lung transplantation models using an ...

Real-Time Monitoring of Cellular Dynamics in Mouse Lung Transplantation

Intravital Two-Photon Microscopy of the Transplanted Mouse Lung

Complications after lung transplantation are largely related to the host immune system responding to the graft. Such immune responses are regulated by crosstalk between donor and recipient cells. A better understanding of these processes relies on the use of preclinical animal models and is aided by an ability to study intra-graft immune cell trafficking in real-time. Intravital two-photon microscopy can be used to image tissues and organs for depths up to several hundred microns with minimal photodamage, which affords a great advantage over single-photon confocal microscopy. Selective use of transgenic mice with promoter-specific fluorescent protein expression and/or adoptive transfer of fluorescent dye-labeled cells during intravital two-photon microscopy allows for the dynamic study of single cells within their physiologic environment. Our group has developed a technique to stabilize mouse lungs, which has enabled us to image cellular dynamics in na&#239;ve lungs and orthotopically transplanted pulmonary grafts. This technique allows for detailed assessment of cellular behavior within the vasculature and in the interstitium, as well as for examination of interactions between various cell populations. This procedure can be readily learned and adapted to study immune mechanisms that regulate inflammatory and tolerogenic responses after lung transplantation. It can also be expanded to the study of other pathogenic pulmonary conditions.

Author Spotlight: Advancing Lung Transplant Immunology Through Intravital Imaging

Here, we present a protocol to intravitally image the transplanted mouse left lung using two-photon microscopy. This represents a valuable tool for studying cellular dynamics and interactions in real-time following murine lung transplantation.

Complications after lung transplantation are largely related to the host immune system responding to the graft. Such immune responses are regulated by ...