National Institute on Deafness and Other Communication Disorders of the National Institutes of Health under award numbers 1K23DC014082 and 1R21DC017225 (Alexander Hillel). This study was also financially supported by the Triological Society and American College of Surgeons (Alexander Hillel), the American Medical Association Foundation, Chicago, IL (Madhavi Duvvuri) and a T32 NIDCD training grant (Kevin Motz).

NOTE: All methods described here were approved by the Johns Hopkins University Animal Care and Use Committee (MO12M354).
1. Preparation of rapamycin in PLLA-PCL
<ol>
	<li>Prepare two glass vials (with caps) of 70:30 PLLA-PCL polymer solutions (inherent viscosity 1.3&#8722;1.8 DL/G; Table of Materials) solutions, with one vial containing 1.0% rapamycin and the other without rapamycin.
	<ol>
		<li>Make a 1.0% rapamycin containing polymer solution by adding 6 mg of rapamycin to 600 mg of 70:30 PLLA-PCL in a glass vial.</li>
		<li>For polymer solution without rapamycin (control), only add 600 mg of 70:30 PLLA-PCL to the glass vial.</li>
	</ol>
	</li>
	<li>Under a fume hood, add 6 mL of dichloromethane to each glass vial. 
	CAUTION: Dichloromethane is a corrosive material and only glass pipettes should be used. Appropriate safety precautions include personal eye protection and gloves.</li>
	<li>Add 120 &#181;L of glycerol to each glass vial (for 2% glycerol solution). 
	NOTE: The addition of glycerol allows for increased flexibility and decreased stiffness in the stent construct.</li>
	<li>Recap the glass vials and allow 70:30 PLLA-PCL with and without rapamycin to dissolve and homogenize for 6&#8722;12 h. 
	NOTE: Glass vials can also be placed on a rotating shaking platform for faster dissolution.</li>
</ol>
2. Rapamycin elution testing
<ol>
	<li>Pipette 1 mL of 70:30 PLLA-PCL solution containing 1% rapamycin into a glass Petri dish. 
	NOTE: This volume of 70:30 PLLA-PCL solution containing 1% rapamycin will contain 120 &#181;g of rapamycin and represents the total amount of rapamycin in each construct. Alternative volumes and concentrations can be used.</li>
	<li>Allow 70:30 PLLA-PCL with 1% rapamycin to harden into a flat disk in the glass Petri dish.</li>
	<li>Once hardened, place the disk in 2 mL of phosphate-buffered saline (PBS, pH 7.4) and incubate in a 37 &#176;C chamber.</li>
	<li>Collect and replace PBS (2 mL) every 24 h and use collected PBS for high-performance liquid chromatography (HPLC) analysis of rapamycin content.</li>
	<li>Use an autosampler and a C18 4.6 cm x 250 mm HPLC column to run samples through the autosampler injector along with serial dilutions of rapamycin to create a calibrated standard curve. Run each sample in triplicate. 
	NOTE: Serial dilutions of rapamycin to be tested include 10%, 1%, 0.1%, and 0.01%.</li>
	<li>Use a mobile phase of HPLC grade water and acetonitrile with 10/90 volume/volume with a flow rate of 2.0 mL/min. Set absorbance during HPLC for rapamycin to 280 nm. 
	NOTE: Figure 1 shows the elution of rapamycin over a 14 day period.</li>
</ol>
3. Creation of rapamycin-eluting PLLA-PCL murine airway stents
NOTE: Perform steps 3.2&#8722;3.9 using sterile materials and sterile technique to avoid contamination that would influence in vivo and in vitro applications.
<ol>
	<li>Prepare PLLA-PCL solutions containing rapamycin as described in section 1.</li>
	<li>Using a glass Pasteur pipette with a rubber balloon, apply 1 mL of PLLA-PCL solution containing 1% rapamycin onto the venous cannula of a 22 G fluorinated ethylene propylene based angiocatheter (Table of Materials). Hold the angiocatheter in one hand and use the glass pipette to drop the PLLA-PCL solution onto the angiocatheter. Slowly rotate the angiocatheter to ensure a homogenous coverage of the angiocatheter with the PLLA-PCL solution. 
	NOTE: At first, the PLLA-PCL solution will be thin, but as the dichloromethane evaporates during the application onto the angiocatheter, the solution will become more viscous to mold onto the angiocatheter. Continuously turning the angiocatheter during the application is beneficial. This step must be done slowly and meticulously to ensure the consistent thickness of the stent.</li>
	<li>Prop the molded angiocatheter with the tip of the angiocatheter facing downward and the hilt on the edge of a glass Petri dish for drying.</li>
	<li>Allow stents to dry for 24 h in a vacuum hood at room temperature (Figure 2A).</li>
	<li>Remove the stent construct from the underlying angiocatheter by gently twisting the underlying catheter free and sliding it out of the molded stent (Figure 2B).</li>
	<li>Check the stent circumferentially for any defects that may have resulted during the casting process. If a defect in the casted stent is present, discard it.</li>
	<li>Trim each end of the casted stent with fine straight scissors so that the edges are axial.</li>
	<li>Using fine straight scissors, cut 3 mm axial segments of the cast stent for use in a mouse model of LTS (Figure 2C). 
	NOTE: Approximately 8 stents can be made from one casted angiocatheter, with each 3 mm stent containing 120 &#181;g of rapamycin.</li>
	<li>Load the 3 mm stent onto a new 22 venous catheter for use in a mouse model of LTS (Figure 1).</li>
</ol>
4. Laryngotracheal stenosis induction in mice
<ol>
	<li>Prior to conducting in vivo mouse studies obtain approval from the Animal Care and Use Committee.</li>
	<li>Anesthetize a 9-week old male C57BL/6 by giving an intraperitoneal injection of ketamine (80&#8722;100 mg/kg) and xylazine (5&#8722;10 mg/kg). 
	NOTE: Other strains of mice can be substituted, but it is recommended that the mice weight be between 20&#8722;27 g.</li>
	<li>Randomize mice into an experimental group (chemomechanical injury with a placement of a 1% rapamycin-containing PLLA-PCL stent) and two control groups: 1) chemomechanical injury with the placement of a PLLA-PCL stent, 2) chemomechanical injury without the placement of a stent.</li>
	<li>Place a mouse on a surgical platform in a supine position. Use a small loop of thread affixed to the top of the platform to extend the cervical spine by looping it around the central incisors.</li>
	<li>Affix the hands and legs of the mouse to the table using 2 inch pieces of tape. Pinch mouse paw to ensure it has had adequate anesthesia.</li>
	<li>The surgical site is then prepped. The overlying fur should be removed and the skin should be cleaned (alternating iodine or chlorhexidine scrub with alcohol or diluted skin disinfectant) three times. The area should be draped and sterile gloves be worn. The sterilized instruments should be laid on a sterile surface when not in use.</li>
	<li>Make a 1.5 cm midline vertical incision in the neck of the mouse using fine curved iris scissors. Divide the overlying thymus and lateralize the two resulting lobes to visualize the trachea. Divide the overlying sternohyoid and sternothyroid (strap) muscles at the superior attachment bilaterally. 
	NOTE: The entire laryngotracheal complex should be completely exposed after this.</li>
	<li>Pass a 22 G angiocatheter transorally through the larynx into the trachea. Use small forceps to apply pressure to the anterior larynx of the mouse to assist in the correct placement of the angiocatheter. Visualize the white angiocatheter through the trachea to ensure correct placement (non-esophageal). 
	NOTE: The mouse trachea is extremely thin and the white angiocatheter will be visualized through the translucent trachea.</li>
	<li>Pass a bleomycin-coated wire brush through the inserted angiocatheter.</li>
	<li>Slowly withdraw the angiocatheter so that only the wire brush remains in the trachea.</li>
	<li>Apply counter pressure using fine forceps on the trachea to mechanically disrupt the tracheal lumen with the wire brush.</li>
	<li>Reinsert the angiocatheter into the trachea over the wire brush.</li>
	<li>Remove the wire brush and reapply bleomycin.</li>
	<li>Repeat steps 4.8&#8722;4.12 a total of 5x. Apply bleomycin to the brush between each application. 
	NOTE: Validation and description of this model can be found in Hillel et al.8.</li>
	<li>Remove the angiocatheter from the mouse trachea. 
	NOTE: If no stent is to be placed, the incision can be closed with tissue glue.</li>
</ol>
5. Transoral PLLA-PCL stent placement in mice
<ol>
	<li>Load one 3 mm casted stent onto an empty 22 G angiocatheter (Figure 3A). 
	NOTE: The stent should rest 5 mm from the tip of the angiocatheter.</li>
	<li>Draw a thin black vertical line on the 3 mm casted stent. 
	NOTE: This line allows for improved visualization of the stent in the trachea.</li>
	<li>Transorally intubate the mouse with the angiocatheter that has been preloaded with the stent.</li>
	<li>Visualize the stent on the angiocatheter in the trachea through the transcervical incision (Figure 3B). 
	NOTE: The trachea is very thin, allowing the black dye on the stent to be visualized through the trachea. Use this to confirm correct tracheal placement.</li>
	<li>Hold the stent in place by gripping the trachea through the transcervical incision with fine forceps.</li>
	<li>Remove the angiocatheter transorally while maintaining a grip on the stent with the fine forceps. 
	NOTE: It is extremely important not to apply too much force to the stent to not crush the lumen when the angiocatheter is removed. However, enough force is required to ensure the stent is not removed with the angiocatheter. A 0.8 mm sialendoscope can be used to visualize the location and placement of the stent in the trachea.</li>
	<li>Close the transcervical incision with tissue glue.</li>
	<li>Allow each mouse to recover to its original activity level before placing it back in its cage.</li>
</ol>
6. Histologic preparation of samples
<ol>
	<li>Anesthetize mice with inhaled anesthetic and sacrifice mice with cervical dislocation per animal protocol after 7, 14, or 21 days. 
	NOTE: Based on the chronicity of the study, different time intervals can be used (e.g., 28, 30 days, etc.).</li>
	<li>Position a mouse on the surgical platform as described in steps 4.4 and 4.5.</li>
	<li>Reopen the neck incision on the mouse using fine curved iris scissors.</li>
	<li>Expose the trachea per step 4.6.</li>
	<li>Divide the distal trachea below the level of the stent using fine curved iris scissors.</li>
	<li>Divide the proximal trachea below the larynx and above the stent using fine curved iris scissors. 
	NOTE: It is important to divide the distal trachea first to prevent the trachea from retracting into the thorax.</li>
	<li>Separate the divided trachea from the esophagus posteriorly and remove trachea from the mouse. 
	NOTE: Stents will still be retained within the mouse trachea.</li>
	<li>Remove stent from the trachea and fix in 10% formalin for 24 h.</li>
	<li>Embed formalin-fixed mouse trachea specimens in paraffin with the distal end oriented superiorly so that axial cuts of the trachea can be made in the next step.</li>
	<li>Cut 5 &#956;m sections of the paraffin-embedded mouse trachea using a microtome. 
	NOTE: Specimens should be cut axially starting from the distal trachea.</li>
	<li>Obtain a 5 &#956;m representative section every 250 &#956;m along the length of the specimen.</li>
	<li>Complete H&#38;E stain on each representative section.
	<ol>
		<li>Deparaffinize the slides by placing in xylene 2x per slide for 3 min.</li>
		<li>Place the slides in 100% ethanol for 2 min, 95% ethanol for 2 min, and 70% ethanol for 2 min to rehydrate.</li>
		<li>Wash the slides with running tap water for 2 min.</li>
		<li>Stain the slides in hematoxylin for 3&#8722;5 min.</li>
		<li>Wash the slides with running tap water for 5 min.</li>
		<li>Place the slides in 1% acid alcohol for 1 min.</li>
		<li>Wash the slides with running tap water for 1 min.</li>
		<li>Rinse in 0.2% ammonia water for 30 s.</li>
		<li>Rinse the slides in running tap water for 5 min. Dip in 95% ethanol 10x.</li>
		<li>Eosin stain by placing slide in eosin phloxine for 30 s.</li>
		<li>Dehydrate the slides by placing in 95% ethanol for 5 min, followed by absolute ethanol 2x for 5 min.</li>
		<li>Place in xylene 2x for 5 min.</li>
		<li>Mount the slides with xylene-based mounting medium.</li>
	</ol>
	</li>
	<li>Measure the thickness of the lamina propria as described in Hillel et al.8.</li>
</ol>
7. Stent biocompatibility in vivo
<ol>
	<li>Prepare tissue sections as in steps 6.1&#8722;6.4. Do not remove stent from these tracheal sections in order to visualize the changes in inflammation relative to the location of the stent within the lumen of the trachea.
	<ol>
		<li>To determine the acute foreign body response to the stent, observe macrophage and T cell activity using CD3 and F4/80 markers. 
		NOTE: Other markers can also be used to determine acute inflammatory reaction to the stent.</li>
		<li>Obtain 5 &#956;m cut sections of the paraffin-embedded trachea on commercially available hydrophilic plus slides (Table of Materials). Place two sections on each slide. 
		NOTE: These sections should be from an area of the trachea where the PLLA-PCL stent was placed.</li>
	</ol>
	</li>
	<li>Place the slides in xylene 2x for 5 min each.</li>
	<li>Place the slides in 100% ethanol 2x for 3 min each.</li>
	<li>Place the slides in 95% ethanol for 1 min.</li>
	<li>Place the slides in a histology staining rack so that they are immersed in antigen retrieval buffer (Table of Materials) and then place in a vegetable steamer with boiling water for 20 min.</li>
	<li>Remove the staining racks from the steamer and discard the antigen retrieval buffer.</li>
	<li>Replace the buffer with 1 mL of PBS.</li>
	<li>Place the slides in a wet staining box for 1 min.</li>
	<li>Discard the PBS and incubate the slides with Dulbecco&#39;s modified eagle medium (DMEM) with 10% FBS for 30 min.</li>
	<li>Discard the DMEM and add a mixture of two primary antibodies raised in different species. Cover the slides with paraffin film and incubate overnight at 4 &#176;C in a dark room. 
	NOTE: In this protocol rabbit anti-CD3 and rat anti-F4/80 (Table of Materials) were used.</li>
	<li>Wash the slides in PBS 3x for 5 min.</li>
	<li>Incubate the slides with secondary antibodies specific to the primary antibody species (Table of Materials) diluted in DMEM with 10% FBS for 0.5&#8722;1 h at room temperature in the dark.</li>
	<li>Wash the slides in PBS 3x for 5 min in the dark.</li>
	<li>Mount the slides on coverslips with a drop of mounting medium with 4&#39;6-diamidino-2-phenylindole (DAPI). Store the slides in the dark at either 20 &#176;C or 4 &#176;C.</li>
	<li>Observe the stained slides on a laser scan confocal microscope and photograph.</li>
	<li>Quantify the total number of cells staining with DAPI and antibodies of interest for comparison to uninjured trachea.</li>
</ol>
8. Mouse trachea quantitative gene expression analysis
<ol>
	<li>Collect mouse trachea as described in steps 6.1&#8722;6.7 and remove stent. 
	NOTE: Harvested mouse tracheas can be stored in a -80 &#176;C freezer for up to 2 years.</li>
	<li>Desiccate the tracheal tissue harvested in step 8.1 using fine scissors and further homogenize using a bead mill homogenizer with 1.4 mm ceramic beads (Table of Materials) as part of a commercially available column-based RNA extraction kit (Table of Materials). 
	NOTE: Bead mill homogenizer settings = one 40 s cycle at 6 m/s.</li>
	<li>Extract RNA from tracheal lysate using a column-based extraction and purification kit (Table of Materials) following manufacturer&#39;s instructions.</li>
	<li>Quantify RNA from each sample using a spectrophotometer (Table of Materials). 
	NOTE: RNA purity is assessed using the 260/230 nm ratio with a pure RNA sample reading 2.0.</li>
	<li>Create complementary DNA from extracted RNA using reverse transcriptase (Table of Materials) and a thermocycler with the following settings: 5 min at 25 &#176;C (priming), 46 &#176;C for 30 min (reverse transcription), and then 95 &#176;C for 1 min (inactivation of reverse transcriptase).</li>
	<li>Dilute complementary DNA samples with RNase free water to a concentration of 10&#8722;25 ng/mL for use in quantitative real-time PCR.</li>
	<li>Mix 1 &#181;L of the complementary DNA (10&#8722;25 ng/mL) with 10 &#181;L of a PCR mastermix (Table of Materials), 8 &#181;L of ddH2O and 0.5 &#956;L of 10 &#956;M forward and reverse primers for the gene of interest in one well of a 96-well PCR plate (Table of Materials). 
	NOTE: The total volume in each well should be 20 &#956;L. Each well on the 96 well plate should contain only one sample and one gene of interest.</li>
	<li>Repeat steps 8.1&#8211;8.7 for all genes of interest and run each sample for each gene in triplicate. 
	NOTE: The gene-specific forward and reverse primers (Table of Materials) for collagen 1, collagen 3, and the reference gene &#946;-actin are commonly used to investigate markers of fibrosis.</li>
	<li>Place the 96 well plate on the real-time quantitative PCR machine and initiate the following protocol: denature at 95 &#176;C for 15 s and anneal and extend at 60 &#176;C for 60 s for 40 cycles.</li>
	<li>Normalize the cycle threshold value (CT) for gene product detection of each gene of interest (GOI) to the CT value for the reference gene &#946;-actin for all samples to obtain a &#916;CT for each GOI. Then, compare the &#916;CT values for each GOI for treated and untreated samples to generate a &#916;&#916;CT. 
	NOTE: The cycle threshold is the point on the amplification curve when a predetermined amount of gene product is present (&#916;Rn). The &#916;Rn value for each reaction should be determined for each gene of interest and should fall on the linear portion of the amplification curve. 
	NOTE: &#916;CT = CT (GOI) &#8211; CT (reference gene); &#916;&#916;CT = &#916;CT (treated) &#8211; &#916;CT (untreated).</li>
	<li>Calculate the relative change in gene expression between treated and untreated samples as an expression of fold change by calculating 2&#916;&#916;CT.</li>
</ol>

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The authors have nothing to disclose.

The most critical steps for successfully constructing and using a drug-eluting stent in vivo are 1) determining the optimal PLLA-PCL ratio for the desirable drug elution rate, 2) determining the appropriate concentration of drug to be eluted, 3) molding the stents around the angiocatheter for in vivo use, and 4) transorally delivering the stent into the mice after LTS induction without causing fatal airway obstruction.
While there are several methods for drug delivery using stents in animal mo...

Laryngotracheal stenosis (LTS) is a pathologic narrowing of the trachea most often due to iatrogenic post-intubation injury. The combination of bacterial colonization, foreign body response to a tracheostomy or endotracheal tube, and patient-specific factors lead to an aberrant inflammatory response. This maladaptive immune response leads to the deposition of collagen in the trachea, resulting in luminal narrowing of the trachea and subsequent stenosis1,2. As current treatment for this disease is primarily surgical, developing an alternative medically-based treatment paradigm targeting the aberrant inflammatory and profibrotic pathways that lead to excessive collagen deposition has been studied. Rapamycin, which inhibits the mTOR signaling complex, has been shown to have immunosuppressive effects as well as a robust antifibroblast effect. However, when rapamycin is systemically administered, common side effects (e.g., hyperlipidemia, anemia, thrombocytopenia) can be pronounced3. The purpose of our methodology is to develop a vehicle for local drug delivery practicable for use in the airway that would lessen these systemic effects. Our assessments focus on investigating the local immune response to the drug delivery construct as well as its capacity to inhibit fibroblast function and alter the local immune microenvironment. Disease-specific outcomes include in vivo testing that evaluate markers of fibrosis.
Biodegradable drug-eluting stents have been used in animal models of disease in multiple organ systems, including the airway4. For the management of airway stenosis or collapse, previous investigations have used drug-coated silicone and nickel-based stents5. A PLLA-PCL construct was chosen for this particular method because of its drug elution profile and mechanical strength in physiological conditions over a period of 3 weeks, which has been demonstrated in previous published studies6. PLLA-PCL is also a biocompatible and biodegradable material already approved by the FDA4. Biocompatible stents eluting cisplatin and MMC have been studied in large animal models such as rabbits and dogs. However, in these animal models, stents were not placed in an animal model of disease and were implanted transcervically. This study provides a unique method for assessing a biocompatible drug-eluting stent placed transorally in a mouse model of airway injury and laryngotracheal stenosis. A biocompatible stent that elutes an immunomodulatory drug locally and can be miniaturized for study in a murine model is valuable for translational preclinical research. Previous attempts at stent utilization with other material constructs generated robust foreign body responses worsening the underlying inflammation that distinguishes LTS7. This methodology, to our knowledge, is the first of its kind to study the immunomodulatory and antifibrotic effects of a stent-based drug delivery system in a murine model of LTS. The murine model itself offers several advantages for studying the effects of an immunomodulatory drug on the trachea. Genetically modified mice and experimental cohorts of healthy and diseased mice can be studied, which can lead to experimental reproducibility and improve cost-effectiveness. Moreover, the delivery of the stent transorally into the mouse trachea mimics clinical delivery of such a stent in humans, which further highlights the translational advantage of this method. Finally, the relative ease with which the PLLA-PCL stent with the drug can be produced allows for modifications to deliver alternate drug therapies aimed at reducing scar formation in the trachea.

<table><tbody><tr><td>&lt;strong&gt;1. For stent&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>22-gauge angiocatheter</td><td>Jelco</td><td>4050</td><td></td></tr><tr><td>Dichloromethane</td><td>Sigma Aldrich</td><td>270997-100ML</td><td></td></tr><tr><td>Glycerol</td><td>Fisher Scientific</td><td>56-81-5</td><td>Available from other vendors as well.</td></tr><tr><td>PDLGA</td><td>Sigma Aldrich</td><td>739955-5G</td><td></td></tr><tr><td>PLLA-PCL (70 : 30)</td><td>Evonik Industries AG</td><td>65053</td><td></td></tr><tr><td>Rapamycin</td><td>LC Laboratories</td><td>R-5000</td><td></td></tr><tr><td>&lt;strong&gt;2. Animal surgery&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Wire brush</td><td>Mill-Rose Company</td><td>320101</td><td></td></tr><tr><td>&lt;strong&gt;3. For immunohistochemistry staining&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Antigen retrival buffer</td><td>Abcam</td><td>ab93678</td><td>Available from other vendors as well; acidic pH needed</td></tr><tr><td>DAPI</td><td>Cell Signaling</td><td>8961S</td><td></td></tr><tr><td>DMEM</td><td>ThermoFisher Scientific</td><td>11965-092</td><td>Available from other vendors as well.</td></tr><tr><td>FBS (Fetal Bovine Serum)</td><td>MilliporeSigma</td><td>F4135-500ML</td><td></td></tr><tr><td>Goat anti-rabbit-488 antibody</td><td>Lif technology</td><td>a11008</td><td></td></tr><tr><td>Goat anti-rat-633 antibody</td><td>Lif technology</td><td>a21094</td><td></td></tr><tr><td>Hydrophilic plus slide</td><td>BSB7028</td><td></td><td></td></tr><tr><td>PBS</td><td>ThermoFisher Scientific</td><td>100-10023</td><td>Available from other vendors as well.</td></tr><tr><td>Rabbit anti-CD3 antibody</td><td>Abcam</td><td>ab5690</td><td></td></tr><tr><td>Rat antiF4/80 antibody</td><td>Biolengend</td><td>123101</td><td></td></tr><tr><td>Zeiss LSM 510 Meta Confocal Microscope</td><td>Zeiss</td><td></td><td></td></tr><tr><td>&lt;strong&gt;4. For quantative PCR&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>0.5mm glass beads</td><td>OMNI International</td><td>19-645</td><td></td></tr><tr><td>Bead Mill Homoginizer</td><td>OMNI International</td><td></td><td></td></tr><tr><td>Gene Specific Forward/Reverse Primers</td><td>Genomic Resources Core Facility</td><td></td><td></td></tr><tr><td>Nanodrop 2000 spectrophotometer</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Power SYBR Green Mastermix</td><td>Life Technologies</td><td>4367659</td><td></td></tr><tr><td>RNeasy mini kit</td><td>Qiagen</td><td>80404</td><td></td></tr><tr><td>StepOnePlus Real Time PCR system</td><td>Life Technologies</td><td></td><td></td></tr></tbody></table>

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.

The biodegradable PLLA-PCL stent construct loaded with rapamycin used in this study was capable of eluting rapamycin in a consistent and predictable fashion in physiological conditions (Figure 1). Figure 2 shows the PLLA-PCL stent casted around a 22 G angiocatheter for use in a murine model of LTS. To determine if the effects of rapamycin elution in the trachea is efficacious in attenuating fibrosis, measured changes in fibrosis-related gene expression and marke...

Watch this Scientific Journal Video about Design of a Biocompatible Drug-Eluting Tracheal Stent in Mice with Laryngotracheal Stenosis at JoVE.com

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

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 ...

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Research

JoVE Journal

Bioengineering

6.7K Views.  University of California, San Francisco. 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.

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

A Murine Model of Stent Implantation in the Carotid Artery for the Study of Restenosis

Despite the considerable progress made in the stent development in the last decades, cardiovascular diseases remain the main cause of death in western countries. Beside the benefits offered by the development of different drug-eluting stents, the coronary revascularization bears also the life-threatening risks of in-stent thrombosis and restenosis. Research on new therapeutic strategies is impaired by the lack of appropriate methods to study stent implantation and restenosis processes. Here, we describe a rapid and accessible procedure of stent implantation in mouse carotid artery, which offers the possibility to study in a convenient way the molecular mechanisms of vessel remodeling and the effects of different drug coatings.

A model of stent implantation in mouse carotid artery is described. Compared to other similar methods, this procedure is very rapid, simple and accessible, offering the possibility to study in a convenient way the vascular wall reaction to different drug-eluting stents and the molecular mechanisms of restenosis.

Despite  the considerable progress made in the stent development in the last decades,  cardiovascular diseases remain the main cause of death in western ...

Medicine

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are necessary for whole animal experiments that require surgery under deep anesthesia or measurements of lung function. Tracheostomy has been the standard for intubating the airway in these mice to allow mechanical ventilation. Orotracheal intubation has been reported but has not been successfully used in many studies because of the substantial technical difficulty or a requirement for highly specialized and expensive equipment. Here we report a technique of direct laryngoscopy using an otoscope fitted with a 2.0 mm speculum and using a 20 G&#160;intravenous catheter as an endotracheal tube. We have used this technique extensively and reliably to intubate and conduct accurate assessments of lung function in mice. This technique has proven safe, with essentially no animal loss in experienced hands. Moreover, this technique can be used for repeated studies of mice in chronic models.

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

We have developed a simple, reliable, and relatively inexpensive method for endotracheal intubation in mice via direct laryngoscopy using an otoscope with a 2.0 mm speculum. This technique is atraumatic and can be used for repeated measurements in chronic experiments. We find it superior to tracheostomy or previously reported nonsurgical techniques.

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation 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 ...

Transplantation of a 3D Bioprinted Patch in a Murine Model of Myocardial Infarction

Testing regenerative properties of 3D bioprinted cardiac patches in vivo using murine models of heart failure via permanent left anterior descending (LAD) ligation is a challenging procedure and has a high mortality rate due to its nature. We developed a method to consistently transplant bioprinted patches of cells and hydrogels onto the epicardium of an infarcted mouse heart to test their regenerative properties in a robust and feasible way. First, a deeply anesthetized mouse is carefully intubated and ventilated. Following left lateral thoracotomy (surgical opening of the chest), the exposed LAD is permanently ligated and the bioprinted patch transplanted onto the epicardium. The mouse quickly recovers from the procedure after chest closure. The advantages of this robust and quick approach include a predicted 28-day mortality rate of up to 30% (lower than the 44% reported by other studies using a similar model of permanent LAD ligation in mice). Moreover, the approach described in this protocol is versatile and could be adapted to test bioprinted patches using different cell types or hydrogels where high numbers of animals are needed to optimally power studies. Overall, we present this as an advantageous approach which may change preclinical testing in future studies for the field of cardiac regeneration and tissue engineering.

This protocol aims to transplant a 3D bioprinted patch onto the epicardium of infarcted mice modeling heart failure. It includes details regarding anesthesia, the surgical chest opening, permanent ligation of the left anterior descending (LAD) coronary artery and application of a bioprinted patch onto the infarcted area of the heart.

Testing regenerative properties of 3D bioprinted cardiac patches in vivo using murine models of heart failure via permanent left anterior descending (LAD) ...

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 ...

Low-Dose Gamma Radiation Sterilization for Decellularized Tracheal Grafts

One of the main key aspects in ensuring that a transplant evolves correctly is the sterility of the medium. Decellularized tracheal transplantation involves implanting an organ that was originally in contact with the environment, thus not being sterile from the outset. While the decellularization protocol (through detergent exposition [2% sodium dodecyl sulfate], continuous stirring, and osmotic shocks) is conducted in line with aseptic measures, it does not provide sterilization. Therefore, one of the main challenges is ensuring sterility prior to in vivo implantation. Although there are established gamma radiation sterilization protocols for inorganic materials, there are no such measures for organic materials. Additionally, the protocols in place for inorganic materials cannot be applied to organic materials, as the established radiation dose (25 kGy) would completely destroy the implant. This paper studies the effect of an escalated radiation dose in a decellularized rabbit trachea. We maintained the dose range (kGy) and tested escalated doses until finding the minimal dose at which sterilization is achieved. After determining the dose, we studied effects of it on the organ, both histologically and biomechanically. We determined that while 0.5 kGy did not achieve sterility, doses of both 1 kGy and 2 kGy did, with 1 kGy, therefore, being the minimal dose necessary to achieve sterilization. Microscopic studies showed no relevant changes compared to non-sterilized organs. Axial biomechanical characteristics were not altered at all, and only a slight reduction in the force per unit of length that the organ can radially tolerate was observed. We can therefore conclude that 1 kGy achieves complete sterilization of decellularized rabbit trachea with a minimal, if any, effects on the organ.

Obtaining sterilization is essential for tracheal tissue transplant. Herein, we present a sterilization protocol using low-dose gamma irradiation that is fully tolerated by organs.

One of the main key aspects in ensuring that a transplant evolves correctly is the sterility of the medium. Decellularized tracheal transplantation ...

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 ...

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

Summary

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Chapters in this video

A Murine Model of Stent Implantation in the Carotid Artery for the Study of Restenosis

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

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

An Improved Method for Rapid Intubation of the Trachea in Mice

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

Transplantation of a 3D Bioprinted Patch in a Murine Model of Myocardial Infarction

Imaging-Guided Bioreactor for Generating Bioengineered Airway Tissue

Low-Dose Gamma Radiation Sterilization for Decellularized Tracheal Grafts

A Heterotopic Mouse Model for Studying Laryngeal Transplantation

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