We would like to acknowledge Robert Strouse and the Research Information Solutions &#38; Innovations division at Nationwide Children&#39;s Hospital for their support in graphic design. This work was supported by a grant from the NIH (NHLBI K08HL138460).

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) at Nationwide Children&#39;s Hospital.
1. Scaffold Manufacturing
<ol>
	<li>Prepare a polymer nanofiber precursor solution by: 1) dissolving 8 wt% PET in 1,1,1,3,3,3-hexafluoroisopropanol and heating the solution to 60 &#176;C and by 2) dissolving 3 wt% PU in 1,1,1,3,3,3-hexafluoroisopropanol at room temperature.</li>
	<li>Once cooled, combine the solutions to create a final polymer mixture of PET and PU (20:80).</li>
	<li>Electrospin the PET+ PU solution onto a stainless steel rod (1 mm diameter) utilizing a 20G blunt tip needle on a 60 mL syringe filled with the PET:PU solution using a high-voltage DC power supply set to +14 kV on the needle, another high-voltage DC power supply set to -3 kV, a 5 mL/h flow rate, and a 20 cm tip-to-substrate distance. Rotate the rod at 350 rpm during fiber deposition.</li>
	<li>Continue electrospinning until the desired scaffold wall thickness of 300 &#181;m is achieved. Slide the scaffold off the rod, then hold it under vacuum overnight to remove any residual solvent.</li>
	<li>Sterilize the scaffold using an ultraviolet light dosage of 350 mJ/cm2 prior to implantation.</li>
</ol>
2. Bone Marrow-Derived Mononuclear Cell (BM-MNC) Harvesting
<ol>
	<li>Euthanize a 6-8 weeks old, female, C57BL/6 mouse with a cocktail of ketamine (200 mg/kg), xylazine (20 mg/kg) and ketoprofen (10 mg/kg). Check for complete euthanasia by tail-pinch method. Be sure to follow local guidelines for euthanasia.&#160;</li>
	<li>Under sterile conditions, remove the skin over the femurs and tibias to expose bone using fine scissors and micro-Adson forceps. Use Dumont #5 forceps and Dumont #5/45 forceps to remove the fascia and the tendons. Separate the bones and cut each of them on both the ends. Using a 5 mL syringe and a 25G needle, flush the bone marrow in a Petri dish containing 30 mL of Rosewell Park Memorial Institute (RPMI) media. Filter this RPMI with bone marrow through a 70 &#181;m nylon cell strainer into a 50 mL tube.</li>
	<li>In a Biosafety Cabinet, place 5 mL of polysurcrose and sodium diatrizoate in a 15 mL tube. Gently add the RPMI containing bone marrow mononuclear cells down the side of the tube to avoid mixing of the two layers. Centrifuge at 461 x g for 30 min with &#34;Brake 1&#34; at 24 &#176;C. 
	NOTE: On our centrifuge, brake 1 stands for the longest run out time during deceleration.</li>
	<li>Out of the three layers formed, discard the top (pink) layer of plasma and collect the middle (clear) layer consisting of the bone marrow mononuclear cells. Discard the precipitate of red blood cells.</li>
	<li>Dilute the bone marrow mononuclear cells (BM-MNC) in phosphate buffered saline (PBS) in a 1:1 ratio and centrifuge at 461 x g for 10 min with &#34;Brake 9&#34; at 24 &#176;C.</li>
	<li>Remove the supernatant and dilute the pellet with 5 mL of PBS. Centrifuge at 461 x g for 10 min with &#34;Brake 9&#34; at 24 &#176;C.</li>
	<li>Remove the supernatant and dilute the pellet with ~10 mL of RPMI.</li>
	<li>Dilute 10 &#181;L of the BM-MNC solution with an equal volume of 0.4% trypan blue in a 1 mL tube. Place 10 &#181;L of the solution in a cell counting chamber slide. Count the cells using an automated cell counter. Repeat the step and calculate the average number of cells. 
	NOTE: An average cell count of 1 X 108 cells was obtained from the use of two donor mice. This would be equivalent to two femurs, and two tibias that bone marrow is isolated from.</li>
	<li>Centrifuge the BM-MNC solution at 461 x g for 10 min with &#34;Brake 9&#34; at 24 &#176;C. Remove the supernatant and dilute the cell concentration to 107 cells/graft. 
	NOTE: We have studied seeding efficiency between 1, 10 and 100 million cells and found that 10 million cells per graft yields the highest seeding efficiency.</li>
</ol>
3. Cell Seeding on the Grafts
<ol>
	<li>Measure the lengths of the scaffolds and if needed, cut them to a length of 5 mm.</li>
	<li>Pre-wet the scaffold with 5 &#181;L of RPMI for 5 min. Remove the RPMI.</li>
	<li>Add 5 &#181;L of the BM-MNC solution to the lumen of the scaffold for 10 min.</li>
	<li>Pass a 21G needle through the lumen of the graft and incubate the graft in 1000 &#181;L of RPMI overnight at 37 &#176;C in an incubator.</li>
</ol>
4. Graft Implantation
NOTE: Care should be taken to maintain aseptic technique during the graft implantation procedure.
<ol>
	<li>Use 6-8-week-old female C57BL/6 mice as recipients for the cell seeded grafts. Inject enrofloxacin (10mg/kg) subcutaneously 24 h before surgery and just before incision.</li>
	<li>Weigh the mouse and administer a 0.1 mL/10 g dose of the anesthetic cocktail of ketamine (100 mg/kg), xylazine (10 mg/kg) and ketoprofen (5 mg/kg) as analgesia intraperitoneally.</li>
	<li>Check if the plane of anesthesia has been achieved by tail-pinch method. On confirming the level of sedation, apply a sterile ophthalmic ointment to the eyes and clip the hair on the surgical site from chin to clavicles. Place the animal on a pad in a dorsal recumbent position. Disinfect the surgical site using first a povidone-iodine prep pad, followed by an alcohol prep pad (70% alcohol), and once more a povidone-iodine prep pad.</li>
	<li>Place the animal under the dissecting microscope with its head away from the surgeon. Make a midline incision ranging from the clavicles to the hyoid bone with the help of fine scissors and micro-Adson forceps. Clean the fascia with Dumont #5 and Dumont #7 fine forceps, along with a sterile cotton swab if needed, and insert a self-retaining Colibri retractor.</li>
	<li>Open the strap muscles (Figure 3A) with Dumont #5 and Dumont #7 fine forceps to expose the thyroid cartilage, cricoid cartilage and the trachea. Bluntly separate the trachea from the recurrent laryngeal nerves running parallel on either side, followed by circumferential separation of the trachea from the esophagus (Figure 3B).</li>
	<li>Using a 20G needle and surgical marker, stain the anterior portion of the trachea.</li>
	<li>Identify the trachea and make an incision below the third tracheal cartilage ring and transect the trachea using a pair of Vannas-Tubingen spring scissors and a pair of Dumont #7 fine forceps. Holding it with the fine curved forceps, secure&#160;the distal trachea to the sternum using sterile 9-0 nylon suture to create a temporary tracheostomy (Figure 3C). NOTE: An alternative suture material that can be used is the 9-0 PDS for tracheal implantation and muscle reapproximation.&#160;</li>
	<li>With a 20G needle and a surgical marker, stain the graft to represent the anterior portion.</li>
	<li>Implant the graft inserting the proximal-posterior, proximal-lateral and proximal-anterior sutures, in that order using 9-0 sterile nylon suture (Figure 3D), Dumont #7 fine forceps and a needle holder.</li>
	<li>Turn the animal 180&#176; to place its tail away from the surgeon. Release the temporary tracheostomy. The distal trachea is incompletely transected to permit the use of the resected segment as a stump for retraction. When no longer needed, the 5 MM trachea segment is removed completely.</li>
	<li>Complete the distal anastomosis by placing the sutures in the similar fashion as the proximal anastomosis.</li>
	<li>Re-approximate the graft position and the strap muscles. Close the incision using 9-0 sterile nylon sutures in a running pattern.</li>
	<li>Inject 0.1 mL of buprenorphine (0.1 mg/kg) subcutaneously and place the animal in a recovery cage placed on a heating pad without other animals in the cage.</li>
	<li>Observe the mouse until it is conscious and able to ambulate, then transfer the mouse to a new cage with soft bedding, moist chow and medicated water (ibuprofen, 30 mg/kg) for 48 h. After this time period, return the mouse to a normal cage with other mice. 
	NOTE: Mice that maintain respiratory distress or decreased activity after implantation are observed by laboratory and/or animal staff in a 32 &#176;C incubator. If this continues for more than 48 h, then the mice should be euthanized.</li>
</ol>
5. Histology and Immunohistochemistry
NOTE: Hematoxylin and eosin stains were performed using standard technique by Nationwide Children&#39;s Hospital Morphology Core. Immunohistochemistry was performed according to the below steps.
<ol>
	<li>Deparaffinize the slides using 1) xylene for 5 min, 2) xylene for 5 min, 3) 100% ethanol for 5 min, 4) 90% ethanol for 5 min, 5) 70% ethanol for 5 min, 6) distilled H2O (dH2O) for 5 min.</li>
	<li>Submerge the slides in citrate buffer and place in water-filled pressure cooker. Heat for 10 min then allow to cool to room temperature. Wash with 0.1% bovine serum albumin in PBS (BSA-PBS) for 5 min. Rinse with dH2O. 
	NOTE: Heating can also be done with a hot water bath or microwave for 15 min.</li>
	<li>Incubate the slides with 3% normal goat serum in PBS for 1 h at room temperature. Wash with 0.1% BSA-PBS for 5 min. Incubate in primary antibody with concentrations for keratin 5 (1:1000)6, keratin 14 (1:250)6 and F4/80 (1:100)7 for 18 h overnight at 4 &#176;C.</li>
	<li>Wash with 0.1% BSA-PBS twice for 10 min each wash. Incubate with appropriate secondary antibody at concentration of 1:500 for K5/K14 and 1:300 F4/80 for 1 h at room temperature. Wash with 0.1% BSA-PBS twice for 10 min each wash.</li>
	<li>Mount using 4&#39;, 6-diamidino-2-phenolindole (DAPI) containing mounting media and glass cover slips. Allow to sit for 30 min before imaging.</li>
</ol>

<ol>
	<li>Macchiarini, P., 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=Clinical+transplantation+of+a+tissue-engineered+airway.">Clinical transplantation of a tissue-engineered airway.</a> The Lancet. 372 (9655), 2023-2030 (2008).</li><li>Jungebluth, P., 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=Tracheobronchial+transplantation+with+a+stem-cell-seeded+bioartificial+nanocomposite:+A+proof-of-concept+study.">Tracheobronchial transplantation with a stem-cell-seeded bioartificial nanocomposite: A proof-of-concept study.</a> The Lancet. 378 (9808), 1997-2004 (2011).</li><li>Elliott, M. J., 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=Stem-cell-based,+tissue+engineered+tracheal+replacement+in+a+child:+A+2-year+follow-up+study.">Stem-cell-based, tissue engineered tracheal replacement in a child: A 2-year follow-up study.</a> The Lancet. 380 (9846), 994-1000 (2012).</li><li>Chiang, T., Pepper, V., Best, C., Onwuka, E., Breuer, C. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clinical+Translation+of+Tissue+Engineered+Trachea+Grafts.">Clinical Translation of Tissue Engineered Trachea Grafts.</a> Annals of Otology, Rhinology and Laryngology. 125 (11), 873-885 (2016).</li><li>Clark, E. 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=Effect+of+cell+seeding+on+neotissue+formation+in+a+tissue+engineered+trachea.">Effect of cell seeding on neotissue formation in a tissue engineered trachea.</a> Journal of Pediatric Surgery. 51 (1), 49-55 (2016).</li><li>Cole, B. B., Smith, R. W., Jenkins, K. M., Graham, B. B., Reynolds, P. R., Reynolds, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tracheal+basal+cells:+A+facultative+progenitor+cell+pool.">Tracheal basal cells: A facultative progenitor cell pool.</a> American Journal of Pathology. 177 (1), 362-376 (2010).</li><li>Onwuka, 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=The+role+of+myeloid+cell-derived+PDGF-B+in+neotissue+formation+in+a+tissue-engineered+vascular+graft.">The role of myeloid cell-derived PDGF-B in neotissue formation in a tissue-engineered vascular graft.</a> Regenerative Medicine. 12 (3), 249-261 (2017).</li><li>Grimmer, J. F., 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=Tracheal+reconstruction+using+tissue-engineered+cartilage.">Tracheal reconstruction using tissue-engineered cartilage.</a> Archives of Otolaryngology - Head and Neck Surgery. 130 (10), 1191-1196 (2004).</li><li>Wood, M. W., Murphy, S. V., Feng, X., Wright, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tracheal+reconstruction+in+a+canine+model.">Tracheal reconstruction in a canine model.</a> Otolaryngology - Head and Neck Surgery (United States). 150 (3), 428-433 (2014).</li><li>Haag, J., 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=Biomechanical+and+angiogenic+properties+of+tissue-engineered+rat+trachea+using+genipin+cross-linked+decellularized+tissue.">Biomechanical and angiogenic properties of tissue-engineered rat trachea using genipin cross-linked decellularized tissue.</a> Biomaterials. 33 (3), 780-789 (2012).</li><li>Best, C. A., 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=Designing+a+tissue-engineered+tracheal+scaffold+for+preclinical+evaluation.">Designing a tissue-engineered tracheal scaffold for preclinical evaluation.</a> International Journal of Pediatric Otorhinolaryngology. 104, 155-160 (2018).</li></ol>

The authors declare no competing financial interests.

Development of a mouse model for tissue engineered tracheas is essential in understanding the factors that have limited clinical translation of the TETGs; namely graft collapse, stenosis and delayed epithelialization4. A few factors that contribute to these limitations include selection of graft material, the manufacturing process, scaffold design and cell seeding protocols. This model allows for faster evaluation of these factors in order to understand the cellular and molecular mechanisms affecting them.
We have shown here the presence of epithelial cells (Figure 4) as well as the ability of the model to recapitulate graft stenosis (Figure 5); however, successful use of the model depends on a few key steps during the implantation process. First, during initial incision and isolation of the mouse trachea, it is essential to avoid injury or stretching of the adjacent recurrent laryngeal nerves located in the tracheoesophageal groove. Another important factor during resection of the native trachea is to avoid any damage to the esophagus sitting posterior to the trachea.
Endotracheal intubation can be avoided by creating a temporary tracheostomy to allow airflow during the procedure. It is important, however, to keep anything (i.e., tissue) from blocking the temporary distal tracheostomy as well as maintaining the trachea outside of the body cavity to facilitate respiration. Graft telescoping is a common occurrence in graft implantations and may be seen on histological sections.
Other animal models that have been utilized in studying TETGs include New Zealand white rabbits, dogs, rats, pigs, and sheep8,9,10,11; many of which can be cost prohibitive and lack comprehensive quantification methods (i.e., ELISAs, Immunohistochemistry antibodies, PCR primers, etc.) and/or transgenic options. Thus, large animal models used are often underpowered or are unable to answer mechanistic questions. The use of transgenic mice in this model as well as the straightforward graft design and manufacturing process make it ideal for faster mechanistic studies looking at graft stenosis and re-epithelialization. The orthotopic transplantation used in this TETG model provides the benefit of the graft being exposed to the external environment and the ability to potentially measure epithelial cell differentiation. However, it is important to remember that translation of transgenic studies to large animal models and clinical trials can be difficult. Furthermore, while this mouse model can potentially be more cost effective in increasing population size, it does not currently have the same clinical diagnostic and therapeutic capabilities as in larger animal models (i.e., bronchoscopy with endoscopic interventions) making it challenging to compare between outcomes.
Future work with this model will be focused on investigating optimal cell seeding conditions (cell type, density, etc.) and its role in limiting graft stenosis and improving epithelialization. Scaffold design is also an important variable that has significant effects on graft performance in vivo and will require more investigation as to the optimal scaffold material and microstructure. These studies will assist in the translation of TETGs to large animal models and eventually human clinical trials.

Long-segment tracheal defects can present as rare congenital conditions such as complete tracheal rings and tracheal agenesis, as well as trauma, malignancy, and infection. When exceeding 6 cm in adults or 30% of the tracheal length in children, these defects cannot be treated by surgical reconstruction. Attempts to replace the airway with autologous tissue, cadaveric transplants, and artificial constructs have been plagued by chronic infection, granulation, mechanical failure, and stenosis.
Tissue engineered tracheal grafts (TETGs) can potentially address these problems while avoiding the need for life-long immunosuppression. In the last decade, TETGs have been tested in animal models and utilized clinically in rare instances of compassionate use1,2,3. In both clinical and large animal studies, post-operative recovery from tissue engineered airway replacement required numerous interventions to combat stenosis (defined as &#62;50% luminal narrowing) and maintain airway patency. Additional TETG work has sought to reduce this stenosis through evaluating the role of cell seeding choice, vascularization and scaffold design. Cell seeding choices and scaffold design aimed at restoring native trachea structure/function have mainly been focused on respiratory epithelial cells and chondrocytes seeded on various resorbable, non-resorbable and decellularized scaffolds. As vascularization likely plays an major role in the development of stenosis, other groups have focused on optimizing in vitro or heterotopic models to expedite revascularization or neoangiogenesis4. Nonetheless, achieving successful vascularization while also maintaining a mechanically competent and functional TETG remains a challenge. Despite recent advances, minimizing stenosis remains a critical obstacle to clinical translation.
To investigate this histopathological response to TETG implantation in vivo, we developed an ovine model of tissue engineered tracheal replacement. The graft was composed of a mixed polyethylene terephthalate (PET) and polyurethane (PU) electrospun scaffold seeded with bone marrow-derived mononuclear cells (BM-MNCs). In this small cohort, we demonstrated that seeded autologous BM-MNCs accelerated re-epithelialization and delayed stenosis5. Although seeding with autologous BM-MNCs improved survival, the cellular mechanism by which BM-MNCs modulate the formation of functional neotissue remains unclear.
Investigation on the cellular level required development of a murine model of tissue engineered tracheal replacement. Similar to the ovine study, we utilized a PET:PU electrospun scaffold seeded with BM-MNCs. Consistent with the ovine model, TETG stenosis developed over the course of the first two weeks following implantation1,2,3,5. This suggested that the murine model recapitulated the pathology observed previously, enabling us to further interrogate the cellular mechanisms underlying airway stenosis.
In this report, we detail our protocol for tissue engineered tracheal replacement in the mouse model including scaffold manufacturing, BM-MNC isolation, graft seeding, and implantation (Figure 1, Figure 2).

<table><tbody><tr><td>0.9% Sodium chloride injection</td><td>APP Pharmaceuticals</td><td>NDC 63323-186-10</td><td></td></tr><tr><td>10cc serological pipet</td><td>Falcon</td><td>357551</td><td></td></tr><tr><td>18G 1.5in. Needle</td><td>BD</td><td>305190</td><td></td></tr><tr><td>1mL Syringe</td><td>BD</td><td>309659</td><td></td></tr><tr><td>24-well plate</td><td>Corning</td><td>3526</td><td></td></tr><tr><td>25cc serological pipet</td><td>Falcon</td><td>356535</td><td></td></tr><tr><td>25G 1in. Needle</td><td>BD</td><td>305125</td><td></td></tr><tr><td>50cc tube</td><td>BD</td><td>352070</td><td></td></tr><tr><td>Alcohol prep pads</td><td>Fisher Healthcare</td><td>NDC 69250-661-02</td><td></td></tr><tr><td>Baytril (enrofloxacin) solution</td><td>Bayer Healthcare, LLC</td><td>NDC 0859-2267-01</td><td></td></tr><tr><td>Black polyamide monofilament suture, 9-0</td><td>AROSurgical Instruments Corporation</td><td>T05A09N10-13</td><td></td></tr><tr><td>C57BL/6, female</td><td>Jackson laboratories</td><td>664</td><td>6-8 weeks old</td></tr><tr><td>Citrate Buffer pH 6.0 20x concentrate</td><td>ThermoFisher</td><td>5000</td><td></td></tr><tr><td>Colibri retractors</td><td>F.S.T</td><td>17000-04</td><td></td></tr><tr><td>Cotton tipped applicators</td><td>Fisher scientific</td><td>23-400-118</td><td></td></tr><tr><td>Cytokeratin 14 Monoclonal Antibody</td><td>ThermoFisher</td><td>MA5-11599</td><td></td></tr><tr><td>Dumont #5 Forceps</td><td>F.S.T</td><td>11251-20</td><td></td></tr><tr><td>Dumont #5/45 forceps</td><td>F.S.T</td><td>11251-35</td><td></td></tr><tr><td>Dumont #7 - Fine Forceps</td><td>F.S.T</td><td>11274-20</td><td></td></tr><tr><td>F4/80 Rat anti-mouse antibody</td><td>Bio-Rad</td><td>MCA497R</td><td></td></tr><tr><td>Ficoll</td><td>Sigma</td><td>10831-100mL</td><td></td></tr><tr><td>Fine scissors- Sharp-blunt</td><td>F.S.T</td><td>14028-10</td><td></td></tr><tr><td>Fisherbrand Premium Cover Glasses</td><td>ThermoFisher</td><td>12-548-5M</td><td></td></tr><tr><td>Fluoroshield Mounting Media with DAPI</td><td>Abcam</td><td>ab104139</td><td></td></tr><tr><td>Goat-anti mouse IgG Secondary Antibody Alexa Fluor 594</td><td>ThermoFisher</td><td>A-11001</td><td></td></tr><tr><td>Goat-anti Rabbit IgG Secondary Antibody Alexa Fluor 594</td><td>ThermoFisher</td><td>A-11012</td><td></td></tr><tr><td>Goat-anti Rat IgG Secondary Antibody Alexa Fluor 647</td><td>ThermoFisher</td><td>A-21247</td><td></td></tr><tr><td>Ibuprofen</td><td>Precision Dose, Inc</td><td>NDC 68094-494-59</td><td></td></tr><tr><td>Iodine prep pads</td><td>Professional disposables international, Inc.</td><td>NDC 10819-3883-1</td><td></td></tr><tr><td>Keratin 5 Polyclonal Antibody, Purified</td><td>BioLegend</td><td>905501</td><td></td></tr><tr><td>Ketamine hydrochloride injection</td><td>Hospira Inc.</td><td>NDC 0409-2053</td><td></td></tr><tr><td>Micro-Adson forceps</td><td>F.S.T</td><td>11018-12</td><td></td></tr><tr><td>Microscope</td><td>Leica</td><td>M80</td><td></td></tr><tr><td>Non-woven sponges</td><td>Covidien</td><td>441401</td><td></td></tr><tr><td>Opthalmic ointment</td><td>Dechra Veterinary products</td><td>NDC 17033-211-38</td><td></td></tr><tr><td>PBS</td><td>Gibco</td><td>10010-023</td><td></td></tr><tr><td>PET/PU (Polyethylene terephthalate &amp;amp; Polyurethane) scaffolds</td><td>Nanofiber solutions</td><td></td><td>Custom ordered</td></tr><tr><td>Petri dish</td><td>BD</td><td>353003</td><td></td></tr><tr><td>RPMI 1640 Medium</td><td>Gibco</td><td>11875-093</td><td></td></tr><tr><td>TISH Needle Holder/Forceps</td><td>Micrins</td><td>MI1540</td><td></td></tr><tr><td>Trimmer</td><td>Wahl</td><td>9854-500</td><td></td></tr><tr><td>Vannas-T&amp;uuml;bingen Spring Scissors</td><td>F.S.T</td><td>15008-08</td><td></td></tr><tr><td>Warm water recirculator</td><td>Gaymar</td><td>TP-700</td><td></td></tr><tr><td>Xylazine sterile solution</td><td>Akorn animal health</td><td>NDC 59399-110-20</td><td></td></tr></tbody></table>

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.

Figure 1 illustrates a schematic of TETG seeding and implantation. Bone marrow was harvested from C57BL/6 mice and cultured in vitro. BM-MNCs were isolated by density centrifugation and seeded onto the TETG. Seeded TETGs were implanted into a syngeneic C57BL/6 recipient mouse.
Figure 2 is an overview of the PET:PU TETG scaffold manufacturing process. PET:PU solution was electrospun onto a 20G blunt needle in order to achieve a final TETG wall thickness of 300 &#181;m and lumen diameter of 1 mm to approximate the native mouse trachea. The surface of the TETG can be seen in the scanning electron microscope image with animated mononuclear cells.
The surgical implantation procedure is outlined in Figure 3 with the most important steps shown. Figure 3 shows the separation and retraction of strap muscles (Figure 3A) and subsequent circumferential isolation of the trachea from surrounding tissues (Figure 3B). Figure 3C demonstrates the distal temporary tracheostomy with attachment to the sternal notch to ensure patent airway during the procedure. Finally, Figure 3D shows the graft in place after final anastomoses.
In a small cohort study, four mice were implanted with TETGs and followed over a two-week period. A few basal epithelial cells can be appreciated in basal cell keratin 5 and 14 staining shown in Figure 4A-C. The combined image (Figure 4C) shows the presence of basal cells on the luminal surface of the graft at 7 days after implantation.
Three out of the four animals showed signs of respiratory distress and stridor, two of which required early termination on post-operative days 1 and 7. Upon explantation and histological analysis, graft stenosis was identified as the main contributing factor to the complications. Figure 5A-E gives a view of the post-operative day 7 animal&#39;s stenotic region of the TETG. Of note, telescoping of the graft and native trachea is a common finding due to surgical technique. The low (Figure 5A) and high (Figure 5B) powered photomicrograph of the stenosis demonstrates one of the main complications of TETG implantation. Figure 5C is a representative F4/80 stain revealing the presence of host macrophages in the stenotic region. Figure 5D and Figure 5E again show the basal epithelial cell markers keratin 5 and 14 at the stenotic region of the graft.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59173/59173fig1.jpg" /> 
Figure 1: TETG seeding and implantation schematic. Bone marrow-derived mononuclear cells obtained from a donor mouse were isolated by density centrifugation, seeded onto a scaffold, and implanted into a recipient mouse. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59173/59173fig1large.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/59173/59173fig2.jpg" /> 
Figure 2: Electrospinning of tissue engineered tracheal graft. (A) Overview of electrospinning process. (B) 20G blunt needle used as rotating mandrel to manufacture mouse scaffolds. (C) Animated SEM schematic of scaffold surface after mononuclear cells have been added. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59173/59173fig2large.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/59173/59173fig3.jpg" /> 
Figure 3: Overview of surgical implantation of scaffold. (A) Separation and retraction of strap muscles. (B) Circumferential separation of the trachea from surrounding tissues/organs. (C) Creation of temporary tracheostomy and attachment to the sternal notch. (D) Implanted graft after proximal and distal anastomoses. Scale bar = 2 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59173/59173fig3large.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/59173/59173fig4.jpg" /> 
Figure 4: Histological evidence of basal epithelial cells. Merged immunofluorescence image of basal cell markers keratin 5 (A) and keratin 14 (B) and the merged keratin 5 and keratin 14 (C). The arrow denotes epithelial tissue growth at the scaffold-native tissue junction. The lumen and scaffold are denoted with (*) and (<img alt="Equation" src="/files/ftp_upload/59173/59173eq1.jpg" />), respectively. Scale bar = 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59173/59173fig4large.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/59173/59173fig5.jpg" /> 
Figure 5: Histological visualization of stenotic region of graft. (A) Longitudinally sectioned H&#38;E of length of trachea including graft with distal stenosis near the anastomosis. Scale bar = 500 &#181;m. High power images of selected stenotic region using H&#38;E (B), F4/80 (C), Keratin 5(D), and Keratin 14 (E). Scale bars = 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59173/59173fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Seeding and Implantation of a Biosynthetic Tissue-engineered Tracheal Graft in a Mouse Model at JoVE.com

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

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 ...

seeding-implantation-biosynthetic-tissue-engineered-tracheal-graft

Nanyang Technological University

Research

JoVE Journal

Bioengineering

7.2K Views.  Nationwide Children's Hospital. Klinische toepassingen van weefsel-engineered luchtpijp zijn beperkt als gevolg van grafttenose en vertraagde epitheelialisatie. Een muismodel van orthotopische tracheale vervanging maakt de studie van de cellulaire mechanismen die regeneratie drijven mogelijk. Lange segment defecten van de luchtwegen kunnen groter zijn dan wat de huidige therapieën in staat zijn om te genezen.De weefsel-engineered luchtpijp biedt het potentieel om een vervangend orgaan voor deze voorwaarden te creër...

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

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 ...

Medicine

Generation and Grafting of Tissue-engineered Vessels in a Mouse Model

The construction of vascular conduits is a fundamental strategy for surgical repair of damaged and injured vessels resulting from cardiovascular diseases. The current protocol presents an efficient and reproducible strategy in which functional tissue engineered vessel grafts can be generated using partially induced pluripotent stem cell (PiPSC) from human fibroblasts. We designed a decellularized vessel scaffold bioreactor, which closely mimics the matrix protein structure and blood flow that exists within a native vessel, for seeding of PiPSC-endothelial cells or smooth muscle cells prior to grafting into mice. This approach was demonstrated to be advantageous because immune-deficient mice engrafted with the PiPSC-derived grafts presented with markedly increased survival rate 3 weeks after surgery. This protocol represents a valuable tool for regenerative medicine, tissue engineering and potentially patient-specific cell-therapy in the near future.

Here, we present a protocol to generate tissue engineered vessel grafts that are functional for grafting into mice by double seeding partially induced pluripotent stem cell (PiPSC) - derived smooth muscle cells and PiPSC - derived endothelial cells on a decellularized vessel scaffold bioreactor.

The construction of vascular conduits is a fundamental strategy for surgical repair of damaged and injured vessels resulting from cardiovascular diseases. ...

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

Implantation of Inferior Vena Cava Interposition Graft in Mouse Model

Biodegradable scaffolds seeded with bone marrow mononuclear cells (BMCs) are often used for reconstructive surgery to treat congenital cardiac anomalies. The long-term clinical results showed excellent patency rates, however, with significant incidence of stenosis. To investigate the cellular and molecular mechanisms of vascular neotissue formation and prevent stenosis development in tissue engineered vascular grafts (TEVGs), we developed a mouse model of the graft with approximately 1 mm internal diameter. First, the TEVGs were assembled from biodegradable tubular scaffolds fabricated from a polyglycolic acid nonwoven felt mesh coated with &#949;-caprolactone and L-lactide copolymer. The scaffolds were then placed in a lyophilizer, vacuumed for 24 hr, and stored in a desiccator until cell seeding. Second, bone marrow was collected from donor mice and mononuclear cells were isolated by density gradient centrifugation. Third, approximately one million cells were seeded on a scaffold and incubated O/N. Finally, the seeded scaffolds were then implanted as infrarenal vena cava interposition grafts in C57BL/6 mice. The implanted grafts demonstrated excellent patency (&#62;90%) without evidence of thromboembolic complications or aneurysmal formation. This murine model will aid us in understanding and quantifying the cellular and molecular mechanisms of neotissue formation in the TEVG.

To improve our knowledge of cellular and molecular neotissue formation, a murine model of the TEVG was recently developed. The grafts were implanted as infrarenal vena cava interposition grafts in C57BL/6 mice. This model achieves similar results to those achieved in our clinical investigation, but over a far shortened time-course.

Biodegradable scaffolds seeded with bone marrow mononuclear cells (BMCs) are often used for reconstructive surgery to treat congenital cardiac anomalies. ...

Design of a Biocompatible Drug-Eluting Tracheal Stent in Mice with Laryngotracheal Stenosis

Laryngotracheal stenosis (LTS) is a pathologic narrowing of the subglottis and trachea leading to extrathoracic obstruction and significant shortness of breath. LTS results from mucosal injury from a foreign body in the trachea, leading to tissue damage and a local inflammatory response that goes awry, leading to the deposition of pathologic scar tissue. Treatment for LTS is surgical due to the lack of effective medical therapies. The purpose of this method is to construct a biocompatible stent that can be miniaturized to place into mice with LTS. We demonstrated that a PLLA-PCL (70% poly-L-lactide and 30% polycaprolactone) construct had optimal biomechanical strength, was biocompatible, practicable for an in vivo placement stent, and capable of eluting drug. This method provides a drug delivery system for testing various immunomodulatory agents to locally inhibit inflammation and reduce airway fibrosis. Manufacturing the stents takes 28&#8722;30 h and can be reproduced easily, allowing for experiments with large cohorts. Here we incorporated the drug rapamycin within the stent to test its effectiveness in reducing fibrosis and collagen deposition. Results revealed that PLLA-PCL tents showed reliable rapamycin release, were mechanically stable in physiological conditions, and were biocompatible, inducing little inflammatory response in the trachea. Further, the rapamycin-eluting PLLA-PCL stents reduced scar formation in the trachea in vivo.

Laryngotracheal stenosis results from pathologic scar deposition that critically narrows the tracheal airway and lacks effective medical therapies. Using a PLLA-PCL (70% poly-L-lactide and 30% polycaprolactone) stent as a local drug delivery system, potential therapies aimed at decreasing scar proliferation in the trachea can be studied.

Laryngotracheal stenosis (LTS) is a pathologic narrowing of the subglottis and trachea leading to extrathoracic obstruction and significant shortness of ...

Imaging-Guided Bioreactor for Generating Bioengineered Airway Tissue

Repeated injury to airway tissue can impair lung function and cause chronic lung disease, such as chronic obstructive pulmonary disease. Advances in regenerative medicine and bioreactor technologies offer opportunities to produce lab-grown functional tissue and organ constructs that can be used to screen drugs, model disease, and engineer tissue replacements. Here, a miniaturized bioreactor coupled with an imaging modality that allows in situ visualization of the inner lumen of explanted rat trachea during in vitro tissue manipulation and&#160;culture is described. Using this bioreactor, the protocol demonstrates imaging-guided selective removal of endogenous cellular components while preserving the intrinsic biochemical features and ultrastructure of the airway tissue matrix. Furthermore, the delivery, uniform distribution, and subsequent prolonged culture of exogenous cells on the decellularized airway lumen with optical monitoring in situ are shown. The results highlight that the imaging-guided bioreactor can potentially be used to facilitate the generation of functional in vitro airway tissues.

The protocol describes an imaging-enabled bioreactor that allows the selective removal of the endogenous epithelium from the rat trachea and homogenous distribution of exogenous cells on the lumen surface, followed by long-term in vitro culture of the cell-tissue construct.

Repeated injury to airway tissue can impair lung function and cause chronic lung disease, such as chronic obstructive pulmonary disease. Advances in ...

Transplantation of Bioengineered Lung Using Decellularized Mouse Lungs and Primary Human Endothelial Cells

Lung transplantation is a critical treatment for patients with end-stage lung diseases like idiopathic pulmonary fibrosis, but challenges such as donor shortages and posttransplant complications persist. Bioengineered lungs, integrating patient-specific cells into decellularized animal scaffolds, present a promising alternative. Despite progress in using bioengineered lungs in animal models, functionality and structure remain immature. This protocol addresses a critical barrier in organ bioengineering: the need for a cost-effective experimental platform. By using mouse models instead of larger animals like rats or swine, researchers can significantly reduce the resources required for each experiment, accelerating research progress. 
The protocol outlines a detailed procedure for lung bioengineering using mouse heart-lung blocks and human primary cells, focusing on isolation strategy for the mouse heart-lung block, decellularization, bioreactor setup, perfusion-based organ culture, and orthotopic transplantation of bioengineered lungs. This mouse-scale platform not only reduces experimental costs but also provides a viable framework for optimizing cell types and numbers for recellularization, testing different cell types using histological and molecular methods, and ensuring blood flow post-transplantation. The method holds potential for broad applications, including studying cell interactions in three-dimensional culture conditions, cell-matrix interactions, and ex vivo cancer modeling, thereby advancing the field of organ bioengineering.

This paper describes how to create bioengineered mouse lungs using decellularization and recellularization methods. It also details subsequent orthotopic lung transplantation.

Lung transplantation is a critical treatment for patients with end-stage lung diseases like idiopathic pulmonary fibrosis, but challenges such as donor ...

Implantation of Tissue-Engineered Vascular Graft in Mouse Carotid Artery via Cuff Technique

The development of small-diameter vascular grafts has been a global endeavor, with numerous research groups contributing to this field. Animal experimentation plays a pivotal role in assessing the efficacy and safety of vascular grafts, particularly in the absence of clinical applications. Compared to alternative animal models, the mouse implantation model offers several advantages, including a well-defined genetic background, a mature method for disease model construction, and a straightforward surgical procedure. Based on these advantages, the present study devised a simple cuff technique for the implantation of tissue-engineered vascular grafts in the mouse carotid artery. This technique began with the fabrication of polycaprolactone (PCL) small-diameter vascular grafts via electrostatic spinning, followed by the seeding of macrophages onto the grafts through perfusion adsorption. Subsequently,&#160;the cellularized tissue-engineered vascular grafts were transplanted into the mouse carotid artery using the cuff technique to evaluate patency and regenerative capability. After 30 days of in vivo implantation, vascular patency was found to be satisfactory, with evidence of neo-tissue regeneration and the formation of an endothelial layer within the lumen of the grafts. All data were analyzed using statistical and graphing software. This study successfully established a mouse carotid artery implantation model that can be used to explore the cellular sources of vascular regeneration and the mechanisms of action of active substances. Furthermore, it provides theoretical support for the development of novel small-diameter vascular grafts.

Here, a protocol is presented for the implantation of a tissue-engineered vascular graft into the mouse carotid artery using the cuff technique, providing a suitable animal model for investigating vascular tissue regeneration mechanisms.

The development of small-diameter vascular grafts has been a global endeavor, with numerous research groups contributing to this field. Animal ...

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 ...

Exploring Obliterative Airway Disease in Lung Transplants

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.

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

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

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