This research has been supported in part by the American Thoracic Society Foundation Research Program, the New Jersey Health Foundation, and the National Science Foundation (CAREER Award 2143620) to J.K.; and the National Institutes of Health (P41 EB027062) to G.V.N.

The animal tissue protocol below has been approved by the animal welfare guideline and regulations of the Institute for Animal Care and Use Committee (IACUC) at Stevens Institute of Technology, and it complies with the National Institutes of Health (NIH) guidelines for the use of experimental animals.
1. Design and construction of imaging-guided rat trachea bioreactor
<ol>
	<li>Designing and fabrication of rat trachea bioreactor
	<ol>
		<li>Create a computer-aided design (CAD) model of the bioreactor chamber with relevant design, such as inlets, outlets, and tissue culture chamber, using CAD generator software. For this study, use the geometry and dimensions presented in Figure 1A-C. The tutorial of CAD generator software can be found in16,17.</li>
		<li>Export the generated CAD model to a computer numerical control (CNC) controller software and cut the polytetrafluoroethylene (PTFE) plastic using a CNC machine to create the bioreactor chamber. The tutorial of the CNC controller software can be found in18. 
		NOTE: In addition to PTFE, other plastic materials, such as ultra-high molecular weight polyethylene (UHMWPE) and polyetherimide can be used to fabricate the culture chamber.</li>
		<li>Sterilize all bioreactor components, such as Luer adapters and screws, to avoid contamination of the tissues and cells cultured in the device. Assemble all bioreactor components to the main tissue culture chamber in a clean environment (Figure 1A).</li>
	</ol>
	</li>
	<li>Construction of gradient index (GRIN) lens-based imaging device
	<ol>
		<li>To create the in situ imaging device, insert a tube lens into a stackable lens tube and secure it using a retaining ring. Mount the lens-tube assembly onto a scientific CMOS camera via a C-mount adapter.</li>
		<li>Use software, such as Micro-Manager, to operate the camera and acquire photos and videos. Aim an object located at a long distance (e.g., 10 m from the camera) and adjust the distance between the tube lens and imaging sensor of the camera until a focused image of the object is formed on the computer screen by the imaging software being used.</li>
		<li>Mount a filter lens on a dual-edge super-resolution dichroic mirror and a laser to the device using optical cage system components, including assembly rod, threaded cage plate, and cage cube.</li>
		<li>Connect the objective lens (20x) to the device. Mount a GRIN lens (diameter = 500 &#181;m) at the distal end of the lens tube via the XY translator. Adjust the distance between the GRIN lens and the objective lens to form focused microscopic images (Figure 2A,B).</li>
	</ol>
	</li>
</ol>
2. Isolation of the rat trachea
<ol>
	<li>Sanitize the surgical area and sterilize the surgical instruments using an autoclave at 121 &#176;C for 30 min prior to the surgery.</li>
	<li>Place a rat in the induction chamber and deliver 2.5% of isoflurane for 15 min using a small vaporizer to anesthetize the animal. Assess the anesthesia depth by pedal reflex. To do this, firmly pinch the toe and confirm that the animal does not respond to the toe pinch.</li>
	<li>After anesthesia, remove the isoflurane from the chamber and administer 1 mL of 1,000 unit/mL heparin through the lateral tail vein to prevent blood clotting in the pulmonary vasculature.</li>
	<li>To euthanize the animal, expose the animal to 5% isoflurane for an additional 15-20 min. Remove the animal from the induction chamber and place it on the surgical board in a face-up supine position.</li>
	<li>Fix the rat on the surgical board by immobilizing the legs and tail using adhesive tape. Next, sterilize the rat&#39;s neck and chest regions using 70% isopropyl alcohol (IPA). Open the abdominal cavity by making a 3-4 cm incision using scissors in the skin. 
	NOTE: Be careful not to cut the skin deeply by pointing the tips of the scissors upward.</li>
	<li>Transect the inferior vena cava (IVC) using the scissors and confirm the euthanasia by exsanguination.</li>
	<li>Perform the tracheotomy by making an incision using scissors in the midline of the neck and exposing the trachea. Perform midline thoracotomy by making an incision in the chest wall and cutting between the ribs to reach the trachea end connected to the lung. Next, use a bicep and scissors to cut both ends of the trachea and isolate the trachea.</li>
	<li>After isolation, rinse the trachea using 20 mL of 1x phosphate-buffered saline (1x PBS). Transfer the trachea to the bioreactor using sterilized biceps. Secure the two ends of the trachea to the Luer connector using a 4-0 suture thread.</li>
	<li>Deliver 5 mL of culture medium to the bioreactor chamber using the programmable syringe pump through the tubing connected to the bioreactor chamber at a flow rate of 5 mL/min. 
	NOTE: The culture medium is composed of Dulbecco&#39;s modified Eagle&#39;s medium (DMEM), recombinant human FGF-basic (0.1 ng/mL), fetal bovine serum (FBS, 10%), and antibiotic-antimycotic (1%).</li>
	<li>Tightly close the bioreactor lid with the acrylic plastic lid and screws. Close the bioreactor chamber tubing connections with the male/female Luer plugs to prevent the flow of the culture medium in the tubing.</li>
</ol>
3. Imaging-guided removal of the trachea epithelium
<ol>
	<li>To visualize the trachea lumen either in bright-field or fluorescent, place the bioreactor on the imaging stage (Figure 3A).</li>
	<li>Prepare carboxyfluorescein succinimidyl ester (CFSE) solution (concentration: 100 &#956;M) in the CFSE cell labeling kit by diluting the CFSE solution with 1x PBS. 
	NOTE: The concentration of the original CFSE solution is 10 mM (in dimethyl sulfoxide (DMSO)).</li>
	<li>Infuse 500 &#181;L of the CFSE solution through the trachea via the syringe pump through the tubing connected to the trachea cannula at a flow rate of 5 mL/min. Stop the pump when the CFSE solution fills the trachea. Wait for 10 min, and then wash the trachea lumen by infusion of 10 mL of 1x PBS using the syringe pump to remove the residual unincorporated CFSE reagent.</li>
	<li>Insert the distal imaging end of the GRIN lens into the trachea via the Luer connector attached to one end of the trachea. Then, gently move the GRIN lens inside the trachea until the trachea surface is focused and take photos and videos in bright-field or fluorescence at 20x magnification.</li>
	<li>For capturing bright-field images in the Micro-Manager software follow the steps below.
	<ol>
		<li>Introduce white light through the cage cube to illuminate the trachea lumen. Click on the Live icon to show the luminal surface of the trachea in real-time. Use Imaging Setting &#62; Exposure (ms) to change exposure time to the desired value. In this study, the exposure time used was in the range of 10 ms and 50 ms.</li>
		<li>To adjust the contrast and brightness of images, use the Histogram and Intensity Scaling window to move black and white arrows at the endpoint of the interactive histogram display. Alternatively, use the Auto Stretch option that allows the software to adjust the brightness and contrast automatically to optimal levels.</li>
		<li>Click on the Snap icon to freeze the image. Then, use Setting &#62; Export Images as Displaced to save the images in the desired format. Alternatively, use Setting &#62; ImageJ to directly export the image to the ImageJ software and save it.</li>
	</ol>
	</li>
	<li>To obtain photos and videos in fluorescent mode, illuminate the trachea lumen with CFSE-specific laser light (CFSE: excitation wavelength: 488 nm, emission wavelength: 515 nm) through the cage cube. Follow the steps 3.5.2-3.5.3 to acquire the images. After taking photos and videos, gently remove the GRIN lens from the trachea.</li>
	<li>Perform de-epithelialization as described below.
	<ol>
		<li>Prepare 2% sodium dodecyl sulfate (SDS) decellularization solution in distilled (DI) water. Instill 50 &#181;L of SDS through the trachea at a flow rate of 6.3 mL/min, by using the syringe pump through the trachea cannula to generate a thin film of the detergent solution on the trachea lumen.</li>
		<li>Close the bioreactor&#39;s tubing connections using male/female Luer plugs and transfer the bioreactor to an incubator. Allow the SDS to dwell within the trachea for 10 min at 37 &#176;C. Instill another 50 &#181;L of SDS solution through the trachea and incubate for 10 min.</li>
		<li>Remove lysed epithelium and SDS by irrigating the trachea lumen with 500 &#181;L of 1x PBS via syringe pump at a flow rate of 10 mL/min for 3x. Place the bioreactor on a shaker and mechanically vibrate the bioreactor at 20 Hz frequency and displacement amplitude of approximately 0.3 mm to physically promote detachment of SDS-treated epithelial cells from the trachea lumen. 
		NOTE: In this study, the shaker was custom-built by assembling a subwoofer speaker, a subwoofer plate amplifier, and an accelerometer. A sinusoidal waveform was generated by a computer and fed into the shaker via the amplifier, while the response of the shaker was monitored via the accelerometer (Figure 3B). In addition, conventional shakers, such as electromagnetic and inertia shakers, can be used to promote the detachment of the cells. To do this, set the frequency and acceleration of the shaker to 20 Hz and 0.5 g, respectively (equivalent to 0.3 mm displacement amplitude).</li>
		<li>Instill 500 &#181;L of 1x PBS twice through the trachea lumen to remove residual SDS and cell debris while the trachea is mechanically vibrated. Following the epithelium removal procedure, evaluate the clearance of the epithelial layer by measuring the intensity of CFSE using the GRIN lens imaging device as in step 3.6 (Figure 3C).</li>
	</ol>
	</li>
</ol>
4. Trachea tissue preparation for further analyses
<ol>
	<li>To confirm the epithelium removal, perform further tissue analyses, such as hematoxylin and eosin (H&#38;E) staining, trichrome, pentachrome, and immunostaining. To do this, remove the trachea from the bioreactor, and fix it in 30 mL of 4% neutral buffered paraformaldehyde solution in 1x PBS (pH = 7.4) at 4 &#176;C overnight.</li>
	<li>Dehydrate and embed the fixed trachea tissue by following the steps below.
	<ol>
		<li>After fixation, wash the trachea tissue with 10 mL of 1x PBS, transfer the trachea to 30 mL of 70% alcohol, and keep it at 4 &#176;C overnight. Cut the trachea into small cylindrical sections (~5 mm) using a sharp blade and insert the tissue sections in the tissue embedding cassettes (length x width x height: 4 cm x 2.5 cm x 0.5 cm). Keep two trachea sections in each cassette.</li>
		<li>Dehydrate the sections in a series of isopropyl alcohol (IPA) solutions - 85%, 90%, 95%, 100% - for 1 h in 30 mL of each solution. Remove the previous solution before adding the next solution.</li>
		<li>Upon completion, submerge the cassettes in 30 mL of clearing agent (e.g., xylene) for 2 h to displace the IPA solution from the tissue sections completely. Perform this step in a fume hood with proper ventilation.</li>
		<li>Submerge the cassettes in paraffin for 2 h, and then embed them in paraffin at 4 &#176;C overnight. Next, cut the paraffin-embedded tissues into thin sections (5-8 &#956;m) using a microtome device for the H&#38;E, trichrome, pentachrome, and immunostaining.</li>
	</ol>
	</li>
	<li>To prepare the tissues for scanning electron microscopy (SEM) analysis, fix the trachea in 30 mL of 2.5% glutaraldehyde solution in 1x PBS (pH = 7.4) at 4 &#176;C overnight. Then, dehydrate the fixed trachea tissue for SEM by following the steps below.
	<ol>
		<li>After fixation, rinse the trachea tissue with 10 mL of 1x PBS. Cut the trachea tissue longitudinally into small semi-cylinder sections (length: ~5 mm) using scissors and insert the tissue sections in the cassettes.</li>
		<li>Dehydrate the sections in a series of IPA solutions - 35%, 50%, 70%, 85%, 95%, and 100% - for 10 min in 30 mL of each solution. Remove the previous solution before adding the next solution.</li>
		<li>Perform hexamethyldisilazane (HMDS)-based drying method by submerging the tissues in the following solutions: 100% IPA: HMDS (2:1; v/v) for 10 min, followed by 100% IPA: HMDS (1:2; v/v) for 10 min, and finally 100% HMDS for overnight. 
		NOTE: HMDS is toxic. Work under a fume hood during all drying steps.</li>
		<li>Remove the tissues from the HMDS solution and allow them to dry under the fume hood for 1 h. Mount the section on aluminum pin stubs using a carbon double-sided conductive tape or silver conductive paste for SEM imaging.</li>
	</ol>
	</li>
</ol>
5. Homogeneous distribution of exogenous cells onto the denuded tracheal lumen
<ol>
	<li>Prepare a de-epithelialized rat trachea using the protocol in step 3. Thaw frozen mesenchymal stem cells (MSCs) for 30 s in a 37 &#176;C water bath. 
	NOTE: In this study, we used mesenchymal stem cells (MSCs) as a model cell to show the distribution of exogenous cells onto the de-epithelialized trachea. Ideally, primary airway epithelial cells, basal cells, or induced-human pluripotent cells (iPSCs) can be used for epithelium regeneration purposes.</li>
	<li>Count the cells with a hemocytometer and prepare a cell solution with a concentration of 5 x 106 cells/mL. Label the cells fluorescently by incubating the cells with 2 mL of CFSE solution (concentration: 100 &#181;M) at room temperature for 15 min. Rinse the cells with 5 mL of 1x PBS for 3x and resuspend the cells in fresh culture medium at a final concentration of 3 x 107 cells/mL.</li>
	<li>Prepare hydrogel pre-gel solution as a vehicle for cell delivery. For this study, use collagen I as a delivery vehicle for MSCs cells and follow the manufacturer&#39;s instructions to prepare the pre-gel. For example, to obtain 3.6 mg/mL collagen pre-gel, mix one part of the chilled neutralization solution with nine parts of the rat tail collagen in a 1.5 mL sterile tube. Then, gently pipet the mixture up and down for adequate mixing. 
	NOTE: Other biocompatible hydrogels may be used instead of collagen I according to the study and user needs.</li>
	<li>Once the hydrogel solution is prepared, add the cells quickly to the solution with desired concentration (e.g., 5 x 106 cells/mL). To obtain a uniform cell-hydrogel mixture, mix the cells and gel solution by gently pipetting with a micropipette.</li>
	<li>Attach one end of the trachea within the bioreactor to a programmable syringe pump through a Luer connector. Deliver 5 mL of fresh culture medium at 37 &#176;C into the bioreactor chamber to cover the exterior surface of the trachea using the syringe pump at a flow rate of 5 mL/min.</li>
	<li>Administer a 10 &#181;L bolus of the cell-hydrogel mixture into the de-epithelialized trachea within the bioreactor using the syringe pump at a flow rate of 5 mL/min to generate a cell-hydrogel layer on the trachea lumen (Figure 4).</li>
	<li>After cell injection, place the bioreactor in a sterile cell culture incubator at 37 &#176;C and 5% CO2 for gelation. For collagen I, the gelation occurs in 30 min.</li>
	<li>To visualize the distribution of implanted cells, sterilize the GRIN lens by wiping with 70% IPA or ethanol and place the bioreactor on the imaging stage. Take photos and videos in both bright-field and fluorescent modes as needed.</li>
	<li>After 30 min of cell seeding, infuse 1 mL of culture medium into the trachea lumen using a syringe pump at a flow rate of 1 mL/min.</li>
	<li>Culture the cell-seeded trachea within the bioreactor in an incubator at 37 &#176;C for the desired time. For long-term culture, refresh the culture medium in the trachea lumen and the bioreactor chamber every 24 h. During the cell culture, keep the media inside the lumen static and change it every 24 h, while the media outside the trachea is continuously perfused via a unidirectional flow at 5 mL/min.</li>
	<li>After culturing the cells for a certain time period (e.g., 1 and 4 days in this study), remove the trachea from the bioreactor. Use scissors to cut the trachea longitudinally into two semi-cylinder sections (i.e., upper and lower sections) on days 1 and 4 of in vitro culture to expose the inner surfaces for monitoring cell growth and calculating cell density. Use a conventional fluorescent microscope to visualize the cells on the inner surfaces.</li>
	<li>To acquire the images using Micro-Manager software, follow steps 3.5 and 3.6. Use a fluorescein isothiocyanate (FITC) filter to visualize the CFSE-labeled cells in fluorescence mode. Analyze the images taken by the fluorescent microscope and calculate cell densities on upper and lower sections using the ImageJ software. To calculate the number of cells and the cell density, follow the steps below.
	<ol>
		<li>Open an image in ImageJ software and convert the image to a binary image (16-bit) using Image &#62; Type &#62; 16-bit. Set the image scale with the scale bar using Analyze &#62; Set Scale.</li>
		<li>Adjust the threshold of the image to highlight the structures of the cells to count. Use Image &#62; Adjust &#62; Threshold. Use Analyze &#62; Analyze Particles to count the number of the cells. Calculate the density of the cells by dividing the number of cells by the surface area of the image.</li>
	</ol>
	</li>
	<li>To assess the viability after cell seeding, use a cell viability kit. For this study, incubate the trachea tissue and the cells in the bioreactor at 37 &#176;C for 6 h and stain the cells with 4 mM calcein-AM and 2 mM ethidium homodimer-1 in 1 mL of 1x PBS at room temperature for 15 min.</li>
	<li>After rinsing with 1x PBS, visualize the cells using a fluorescent microscope. Count the live and dead cells using the steps described in step 5.12. Calculate cell viability as the percentage of live cells (stained with calcein-AM) per total cells (both live and dead cells).</li>
</ol>

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<li>Huang, S. X. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+in+vitro+generation+of+lung+and+airway+progenitor+cells+from+human+pluripotent+stem+cells%5BTitle%5D)+AND+%22Nature+Protocols%22%5BJournal%5D)">The in vitro generation of lung and airway progenitor cells from human pluripotent stem cells.</a> Nature Protocols. 10 (3), 413-425 (2015).</li>
<li>Kim, J., O'Neill, J. D., Dorrello, N. V., Bacchetta, M., Vunjak-Novakovic, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Targeted+delivery+of+liquid+microvolumes+into+the+lung%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Targeted delivery of liquid microvolumes into the lung.</a> Proceedings of the National Academy of Sciences of the United States of America. 112 (37), 11530-11535 (2015).</li>
<li>Kim, J., O'Neill, J. D., Vunjak-Novakovic, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rapid+retraction+of+microvolume+aqueous+plugs+traveling+in+a+wettable+capillary%5BTitle%5D)+AND+%22Applied+Physics+Letters%22%5BJournal%5D)">Rapid retraction of microvolume aqueous plugs traveling in a wettable capillary.</a> Applied Physics Letters. 107 (14), 144101(2015).</li>
<li>O'Neill, J. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Decellularization+of+human+and+porcine+lung+tissues+for+pulmonary+tissue+engineering%5BTitle%5D)+AND+%22The+Annals+of+Thoracic+Surgery%22%5BJournal%5D)">Decellularization of human and porcine lung tissues for pulmonary tissue engineering.</a> The Annals of Thoracic Surgery. 96 (3), 1046-1056 (2013).</li>
<li>Sengyoku, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Sodium+hydroxide+based+non-detergent+decellularizing+solution+for+rat+lung%5BTitle%5D)+AND+%22Organogenesis%22%5BJournal%5D)">Sodium hydroxide based non-detergent decellularizing solution for rat lung.</a> Organogenesis. 14 (2), 94-106 (2018).</li>
<li>Walters, M. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Generation+of+a+human+airway+epithelium+derived+basal+cell+line+with+multipotent+differentiation+capacity%5BTitle%5D)+AND+%22Respiratory+Research%22%5BJournal%5D)">Generation of a human airway epithelium derived basal cell line with multipotent differentiation capacity.</a> Respiratory Research. 14 (1), 135(2013).</li>
<li>O'Neill, J. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cross-circulation+for+extracorporeal+support+and+recovery+of+the+lung%5BTitle%5D)+AND+%22Nature+Biomedical+Engineering%22%5BJournal%5D)">Cross-circulation for extracorporeal support and recovery of the lung.</a> Nature Biomedical Engineering. 1 (3), 0037(2017).</li>
<li>Guenthart, B. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Regeneration+of+severely+damaged+lungs+using+an+interventional+cross-circulation+platform%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Regeneration of severely damaged lungs using an interventional cross-circulation platform.</a> Nature Communications. 10 (1), 1985(2019).</li>
<li>Chen, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Non-destructive+vacuum-assisted+measurement+of+lung+elastic+modulus%5BTitle%5D)+AND+%22Acta+Biomaterialia%22%5BJournal%5D)">Non-destructive vacuum-assisted measurement of lung elastic modulus.</a> Acta Biomaterialia. 131, 370-380 (2021).</li>
<li>Dorrello, N. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Functional+vascularized+lung+grafts+for+lung+bioengineering%5BTitle%5D)+AND+%22Science+Advances%22%5BJournal%5D)">Functional vascularized lung grafts for lung bioengineering.</a> Science Advances. 3 (8), 1700521(2017).</li>
</ol>

The authors declare no competing financial interests.

In this work, we created an imaging-guided bioreactor that can allow (i) monitoring of the trachea lumen in situ after the cell removal and exogenous cell delivery and (ii) long-term in vitro culture of the cell-seeded trachea tissue. Using this custom-built bioreactor, we demonstrated (i) selective removal of the endogenous epithelial cells from the trachea lumen using detergent and vibration-assisted airway wash and (ii) uniform distribution of exogenous cells onto the luminal surface of the denuded t...

The luminal surface of the respiratory tract is lined by a layer of epithelium that mainly consists of multi-ciliated, club, goblet, and basal stem cells1,2. The epithelial layer serves as a primary defense mechanism of the lung, acting as a biophysical barrier that protects the underlying airway tissue against inhaled pathogens, particulates, or chemical gases. It protects the airway tissue via multiple mechanisms, including intercellular tight junction formation, mucociliary clearance, and antimicrobial and antioxidant secretion3,4. The defective airway epithelium is associated with devastating respiratory diseases, such as chronic obstructive pulmonary disease (COPD)5, primary ciliary dyskinesia (PCD)6, and cystic fibrosis (CF)7.
Advances in lung-on-chip (LOC) technology represent an opportunity to study human lung development, model various lung diseases, and develop new therapeutic materials in tightly regulated in vitro environments. For example, airway epithelium and endothelium can be cultured on opposite sides of a thin, porous membrane to mimic the gas exchanging lung tissue, allowing faithful disease modeling and drug testing8. Similarly, in vitro disease models have been created to model airway diseases in vitro, such as COPD9 and cystic fibrosis10. However, a major challenge of LOC devices is recapitulating the complex three-dimensional (3D) architecture of the lung tissue and dynamic cell-tissue matrix interactions in vitro11.
Recently, innovative tissue engineering methodologies have been developed that allow manipulation of ex vivo lung tissues12. Using these methodologies, denuded allogenic or xenogeneic tissue grafts can be prepared by removing the endogenous cells from the lung tissue via chemical, physical, and mechanical treatments13. In addition, the preserved native tissue extracellular matrix (ECM) in the decellularized lung scaffolds provide the physio-mimetic structural, biochemical, and biomechanical cues for implanted cells to attach, proliferate, and differentiate14,15.
Here, an imaging-guided bioreactor system created by combining LOC and tissue engineering technologies to allow in vitro tissue manipulation and culture of explanted rat tracheal tissues is reported. Using this airway tissue bioreactor, the protocol demonstrates selective removal of the endogenous epithelial cells without disrupting the underlying subepithelial cellular and biochemical components of the airway tissue. We next show the homogenous distribution and instantaneous deposition of the newly seeded exogenous cells, such as mesenchymal stem cells (MSCs), on the denuded airway lumen by instilling the cell-loaded collagen I pre-gel solution. In addition, by using the micro-optical imaging device integrated into the bioreactor, the visualization of the trachea lumen during epithelium removal and endogenous cell delivery is also done. Further, it is shown that the trachea and newly implanted cells can be cultured in the bioreactor without noticeable cell death and tissue degradation for 4 days. We envision that the imaging-enabled bioreactor platform, the thin film-based de-epithelialization technique, and the cell delivery method used in this study can be useful for generating airway tissues for in vitro disease modeling and drug screening.
The bioreactor includes a rectangular chamber connected to a programmable syringe pump, perfusion pump, and ventilator for culturing isolated rat trachea. The bioreactor features inlets and outlets connected to the trachea or the tissue culture chamber to separately supply reagents (e.g., culture media) to the internal and external spaces of the trachea (Figure 1). A custom-built imaging system can be used to visualize the interior of the in vitro-cultured rat trachea at the cellular level (Figure 2). The endogenous epithelium of the trachea is removed via the instillation of a detergent-based decellularization solution followed by vibration-assisted airway washing (Figure 3). Hydrogel solution, such as type I collagen, is used as a delivery vehicle for seeding exogenous cells uniformly and instantaneously across the denuded trachea lumen (Figure 4). All the materials used to construct the bioreactor and conduct the experiments are provided in the Table of Materials.

<table><tbody><tr><td>1&amp;times; PBS</td><td>Gibco, Thermo Fisher Scientific</td><td>10-010-031</td><td></td></tr><tr><td>3-port connector</td><td>World Precision Instruments</td><td>14048-20</td><td></td></tr><tr><td>4-port connector</td><td>World Precision Instruments</td><td>14047-10</td><td></td></tr><tr><td>Accelerometer</td><td>STMicroelectronics</td><td>IIS3DWBTR</td><td></td></tr><tr><td>Achromatic doublet</td><td>Thorlabs</td><td>AC254-150-A-ML</td><td></td></tr><tr><td>Aluminum pin stub</td><td>TED PELLA</td><td>16111</td><td></td></tr><tr><td>Antibiotic-antimycotic</td><td>Thermo Fisher Scientific</td><td>15240062</td><td></td></tr><tr><td>Assembly rod</td><td>Thorlabs</td><td>ER1</td><td></td></tr><tr><td>Button head screws</td><td>McMaster-Carr</td><td>91255A274</td><td></td></tr><tr><td>Cage cube</td><td>Thorlabs</td><td>C4W</td><td></td></tr><tr><td>Carbon double-sided conductive tape</td><td>TED PELLA</td><td>16073</td><td></td></tr><tr><td>CFSE labelling kit</td><td>Abcam</td><td>ab113853</td><td></td></tr><tr><td>Citrisolv (clearing agent)</td><td>Decon</td><td>1061</td><td></td></tr><tr><td>C-mount adapter</td><td>Thorlabs</td><td>SM1A9</td><td></td></tr><tr><td>Collagen I</td><td>Advanced BioMatrix</td><td>5153</td><td></td></tr><tr><td>Conductive liquid silver paint</td><td>TED PELLA</td><td>16034</td><td></td></tr><tr><td>Dichroic mirror</td><td>Semrock</td><td>DI03-R488</td><td>Reflected laser wavelengths:&amp;nbsp; 473.0 +- 2 nm 488.0 +3/-2 nm</td></tr><tr><td>Dulbecco&#039;s modified Eagle&amp;rsquo;s medium</td><td>Gibco, Thermo Fisher Scientific</td><td>11965118</td><td></td></tr><tr><td>Female luer bulkhead to hose barb adapter</td><td>Cole-Parmer</td><td>EW-45501-30</td><td></td></tr><tr><td>Female luer to tubing barb</td><td>Cole-Parmer</td><td>EW-45508-03</td><td></td></tr><tr><td>Female to male luer connector</td><td>Cole-Parmer</td><td>ZY-45508-80</td><td></td></tr><tr><td>Fetal bovine serum</td><td>Gibco, Thermo Fisher Scientific</td><td>10082147</td><td></td></tr><tr><td>Filter lens</td><td>Chroma Technology Corp</td><td>ET535/50m</td><td></td></tr><tr><td>Fluorescent microscope</td><td>Nikon</td><td>Eclipse E1000 - D</td><td></td></tr><tr><td>Fusion 360</td><td>Autodesk</td><td></td><td></td></tr><tr><td>Hex nut</td><td>McMaster-Carr</td><td>91813A160</td><td></td></tr><tr><td>Hexamethyldisilazane (HMDS)</td><td>Fisher Scientifc</td><td>AC120585000</td><td></td></tr><tr><td>Imaging fiber</td><td>SELFOC, NSG group</td><td>GRIN lens</td><td></td></tr><tr><td>Laser</td><td>Opto Engine</td><td>MDL-D-488-150mW</td><td></td></tr><tr><td>Lens tubes</td><td>Thorlabs</td><td>SM1L40</td><td></td></tr><tr><td>LIVE/DEAD Viability/Cytotoxicity Kit (Invitrogen)</td><td>Thermo Fisher Scientific</td><td>L3224</td><td></td></tr><tr><td>MACH 3 CNC Control Software</td><td>Newfangled Solutions</td><td></td><td></td></tr><tr><td>Objective lens</td><td>Olympus</td><td>UCPLFLN20X</td><td></td></tr><tr><td>Peristaltic Pump</td><td>Cole Parmer</td><td>L/S standard digital pump system</td><td></td></tr><tr><td>Recombinant human FGF-basic</td><td>PeproTech</td><td>100-18B</td><td></td></tr><tr><td>Retaining ring</td><td>Thorlabs</td><td>SM1RR</td><td></td></tr><tr><td>Scientific CMOS camera</td><td>PCO Panda</td><td>PCO Panda 4.2</td><td></td></tr><tr><td>Sodium dodecyl sulfate</td><td>VWR</td><td>97064-472</td><td></td></tr><tr><td>Solidworks (2019)</td><td>Dassault Syst&amp;egrave;mes</td><td></td><td></td></tr><tr><td>Stackable lens tube</td><td>Thorlabs</td><td>SM1L10</td><td></td></tr><tr><td>Subwoofer plate amplifier</td><td>Dayton Audio</td><td>SPA250DSP</td><td></td></tr><tr><td>Subwoofer speaker</td><td>Dayton Audio</td><td>RSS21OHO-4</td><td>Diaphragm diameter: 21 cm</td></tr><tr><td>Syringe Pump</td><td>World Precision Instruments</td><td>AL-4000</td><td></td></tr><tr><td>Threaded cage plate</td><td>Thorlabs</td><td>CP33</td><td></td></tr><tr><td>Threaded luer adapter</td><td>Cole-Parmer</td><td>EW-45513-81</td><td></td></tr><tr><td>Tube lens</td><td>Thorlabs</td><td>AC254-150-A-ML</td><td></td></tr><tr><td>Tygon Tubing</td><td>Cole-Parmer</td><td>13-200-110</td><td></td></tr><tr><td>XY Translator</td><td>Thorlabs</td><td>CXY1</td><td></td></tr></tbody></table>

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 GRIN lens-based in situ imaging modality can allow visualization of the tracheal inner lumen in situ (Figure 5A). Using this imaging method, both bright-field and fluorescent images of the native and de-epithelialized tracheas can be obtained (Figure 5B,C). No fluorescent signal was observed from the native trachea prior to CFSE labeling (Figure 5Bii). However, when the tracheal epithelium was ...

Watch this Scientific Journal Video about Imaging-Guided Bioreactor for Generating Bioengineered Airway Tissue at JoVE.com

Imaging-Guided Bioreactor for Generating Bioengineered Airway Tissue

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 ...

imaging-guided-bioreactor-for-generating-bioengineered-airway-tissue

Research

JoVE Journal

Bioengineering

2.6K Views.  Stevens Institute of Technology. Dit protocol maakt in vitro kweek en directe microscopische visualisatie van de geïsoleerde luchtwegweefsels mogelijk tijdens verschillende weefselmanipulatieprocedures, zoals de-epithelisatie en regeneratie van luchtwegweefsel. De in situ beeldvorming voorgesteld in deze studie maakt snelle en niet-destructieve monitoring van het tracheale lumen mogelijk tijdens beeldvormingsgeleide gecontroleerde verwijdering van het endogene epitheel en levering van de exogene cellen. Het beeldvormingsges...

Video: Imaging-Guided Bioreactor for Generating Bioengineered Airway Tissue

Engineering Biological-Based Vascular Grafts Using a Pulsatile Bioreactor

Much effort has been devoted to develop and advance the methodology to regenerate functional small-diameter arterial bypasses. In the physiological environment, both mechanical and chemical stimulation are required to maintain the proper development and functionality of arterial vessels1,2.
Bioreactor culture systems developed by our group are designed to support vessel regeneration within a precisely controlled chemo-mechanical environment mimicking that of native vessels. Our bioreactor assembly and maintenance procedures are fairly simple and highly repeatable3,4. Smooth muscle cells (SMCs) are seeded onto a tubular polyglycolic acid (PGA) mesh that is threaded over compliant silicone tubing and cultured in the bioreactor with or without pulsatile stimulation for up to 12 weeks. There are four main attributes that distinguish our bioreactor from some predecessors. 1) Unlike other culture systems that simulate only the biochemical surrounding of native blood vessels, our bioreactor also creates a physiological pulsatile environment by applying cyclic radial strain to the vessels in culture. 2) Multiple engineered vessels can be cultured simultaneously under different mechanical conditions within a controlled chemical environment. 3) The bioreactor allows a mono layer of endothelial cells (EC) to be easily coated onto the luminal side of engineered vessels for animal implantation models. 4) Our bioreactor can also culture engineered vessels with different diameter size ranged from 1 mm to 3 mm, saving the effort to tailor each individual bioreactor to fit a specific diameter size. 
The engineered vessels cultured in our bioreactor resemble native blood vessels histologically to some degree. Cells in the vessel walls express mature SMC contractile markers such as smooth muscle myosin heavy chain (SMMHC)3. A substantial amount of collagen is deposited within the extracellular matrix, which is responsible for ultimate mechanical strength of the engineered vessels5. Biochemical analysis also indicates that collagen content of engineered vessels is comparable to that of native arteries6. Importantly, the pulsatile bioreactor has consistently regenerated vessels that exhibit mechanical properties that permit successful implantation experiments in animal models3,7. Additionally, this bioreactor can be further modified to allow real-time assessment and tracking of collagen remodeling over time, non-invasively, using a non-linear optical microscopy (NLOM)8. To conclude, this bioreactor should serve as an excellent platform to study the fundamental mechanisms that regulate the regeneration of functional small-diameter vascular grafts.

Our group has developed a bioreactor culture system that mimics the physiological pulsatile stresses of the cardiovascular system to regenerate implantable small-diameter vascular grafts.

Much effort has been devoted to develop and advance the methodology to regenerate functional small-diameter arterial bypasses. In the physiological ...

Procedure for Lung Engineering

Lung tissue, including lung cancer and chronic lung diseases such as chronic obstructive pulmonary disease, cumulatively account for some 280,000 deaths annually; chronic obstructive pulmonary disease is currently the fourth leading cause of death in the United States1. Contributing to this mortality is the fact that lungs do not generally repair or regenerate beyond the microscopic, cellular level. Therefore, lung tissue that is damaged by degeneration or infection, or lung tissue that is surgically resected is not functionally replaced in vivo. To explore whether lung tissue can be generated in vitro, we treated lungs from adult rats using a procedure that removes cellular components to produce an acellular lung extracellular matrix scaffold. This scaffold retains the hierarchical branching structures of airways and vasculature, as well as a largely intact basement membrane, which comprises collagen IV, laminin, and fibronectin. The scaffold is mounted in a bioreactor designed to mimic critical aspects of lung physiology, such as negative pressure ventilation and pulsatile vascular perfusion. By culturing pulmonary epithelium and vascular endothelium within the bioreactor-mounted scaffold, we are able to generate lung tissue that is phenotypically comparable to native lung tissue and that is able to participate in gas exchange for short time intervals (45-120 minutes). These results are encouraging, and suggest that repopulation of lung matrix is a viable strategy for lung regeneration. This possibility presents an opportunity not only to work toward increasing the supply of lung tissue for transplantation, but also to study respiratory cell and molecular biology in vitro for longer time periods and in a more accurate microenvironment than has previously been possible.

We have developed a decellularized lung extracellular matrix and novel biomimetic bioreactor that can be used to generate functional lung tissue. By seeding cells into the matrix and culturing in the bioreactor, we generate tissue that demonstrates effective gas exchange when transplanted in vivo for short periods of time.

Lung tissue, including lung cancer and chronic lung diseases such as chronic obstructive pulmonary disease, cumulatively account for some 280,000 deaths ...

Nonhuman Primate Lung Decellularization and Recellularization Using a Specialized Large-organ Bioreactor

There are an insufficient number of lungs available to meet current and future organ transplantation needs. Bioartificial tissue regeneration is an attractive alternative to classic organ transplantation. This technology utilizes an organ&#39;s natural biological extracellular matrix (ECM) as a scaffold onto which autologous or stem/progenitor cells may be seeded and cultured in such a way that facilitates regeneration of the original tissue. The natural ECM is isolated by a process called decellularization. Decellularization is accomplished by treating tissues with a series of detergents, salts, and enzymes to achieve effective removal of cellular material while leaving the ECM intact. Studies conducted utilizing decellularization and subsequent recellularization of rodent lungs demonstrated marginal success in generating pulmonary-like tissue which is capable of gas exchange in vivo. While offering essential proof-of-concept, rodent models are not directly translatable to human use. Nonhuman primates (NHP) offer a more suitable model in which to investigate the use of bioartificial organ production for eventual clinical use.
The protocols for achieving complete decellularization of lungs acquired from the NHP rhesus macaque are presented. The resulting acellular lungs can be seeded with a variety of cells including mesenchymal stem cells and endothelial cells. The manuscript also describes the development of a bioreactor system in which cell-seeded macaque lungs can be cultured under conditions of mechanical stretch and strain provided by negative pressure ventilation as well as pulsatile perfusion through the vasculature; these forces are known to direct differentiation along pulmonary and endothelial lineages, respectively. Representative results of decellularization and cell seeding are provided.

Whole-organ decellularization produces natural biological scaffolds that may be used for regenerative medicine. The description of a nonhuman primate model of lung regeneration in which whole lungs are decellularized and then seeded with adult stem cells and endothelial cells in a bioreactor that facilitates vascular circulation and liquid media ventilation is presented.

There are an insufficient number of lungs available to meet current and future organ transplantation needs. Bioartificial tissue regeneration is an ...

Adapting the Electrospinning Process to Provide Three Unique Environments for a Tri-layered In Vitro Model of the Airway Wall

Electrospinning is a highly adaptable method producing porous 3D fibrous scaffolds that can be exploited in in vitro cell culture. Alterations to intrinsic parameters within the process allow a high degree of control over scaffold characteristics including fiber diameter, alignment and porosity. By developing scaffolds with similar dimensions and topographies to organ- or tissue-specific extracellular matrices (ECM), micro-environments representative to those that cells are exposed to in situ can be created. 
The airway bronchiole wall, comprised of three main micro-environments, was selected as a model tissue. Using decellularized airway ECM as a guide, we electrospun the non-degradable polymer, polyethylene terephthalate (PET), by three different protocols to produce three individual electrospun scaffolds optimized for epithelial, fibroblast or smooth muscle cell-culture. Using a commercially available bioreactor system, we stably co-cultured the three cell-types to provide an in vitro model of the airway wall over an extended time period.
This model highlights the potential for such methods being employed in in vitro diagnostic studies investigating important inter-cellular cross-talk mechanisms or assessing novel pharmaceutical targets, by providing a relevant platform to allow the culture of fully differentiated adult cells within 3D, tissue-specific environments.

Adapting the Electrospinning Process to Provide Three Unique Environments for a Tri-layered In Vitro Model of the Airway Wall

Advancements in biomaterial technologies enable the development of three-dimensional multi-cell-type constructs. We have developed electrospinning protocols to produce three individual scaffolds to culture the main structural cells of the airway to provide a 3D in vitro model of the airway bronchiole wall.

Electrospinning is a highly adaptable method producing porous 3D fibrous scaffolds that can be exploited in in vitro cell culture. Alterations to ...

Construction of Defined Human Engineered Cardiac Tissues to Study Mechanisms of Cardiac Cell Therapy

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that replicate the biofidelity of three-dimensional native human myocardium, while also enabling a controlled level of biological complexity, and allowing non-destructive longitudinal monitoring of tissue contractile function. Initially, human engineered cardiac tissues (hECT) were created using the entire cell population obtained from directed differentiation of human pluripotent stem cells, which typically yielded less than 50% cardiomyocytes. However, to create reliable predictive models of human myocardium, and to elucidate mechanisms of heterocellular interaction, it is essential to accurately control the biological composition in engineered tissues. 
To address this limitation, we utilize live cell sorting for the cardiac surface marker SIRP&#945; and the fibroblast marker CD90 to create tissues containing a 3:1 ratio of these cell types, respectively, that are then mixed together and added to a collagen-based matrix solution. Resulting hECTs are, thus, completely defined in both their cellular and extracellular matrix composition.
Here we describe the construction of defined hECTs as a model system to understand mechanisms of cell-cell interactions in cell therapies, using an example of human bone marrow-derived mesenchymal stem cells (hMSC) that are currently being used in human clinical trials. The defined tissue composition is imperative to understand how the hMSCs may be interacting with the endogenous cardiac cell types to enhance tissue function. A bioreactor system is also described that simultaneously cultures six hECTs in parallel, permitting more efficient use of the cells after sorting.

This manuscript describes the creation of defined engineered cardiac tissues using surface marker expression and cell sorting. The defined tissues can then be used in a multi-tissue bioreactor to investigate mechanisms of cardiac cell therapy in order to provide a functional, yet controlled, model system of the human heart.

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that ...

A Multi-Cue Bioreactor to Evaluate the Inflammatory and Regenerative Capacity of Biomaterials under Flow and Stretch

The use of resorbable biomaterials to induce regeneration directly in the body is an attractive strategy from a translational perspective. Such materials induce an inflammatory response upon implantation, which is the driver of subsequent resorption of the material and the regeneration of new tissue. This strategy, also known as in situ tissue engineering, is pursued to obtain cardiovascular replacements such as tissue-engineered vascular grafts. Both the inflammatory and the regenerative processes are determined by the local biomechanical cues on the scaffold (i.e., stretch and shear stress). Here, we describe in detail the use of a custom-developed bioreactor that uniquely enables the decoupling of stretch and shear stress on a tubular scaffold. This allows for the systematic and standardized evaluation of the inflammatory and regenerative capacity of tubular scaffolds under the influence of well-controlled mechanical loads, which we demonstrate on the basis of a dynamic co-culture experiment using human macrophages and myofibroblasts. The key practical steps in this approach&#8212;the construction and setting up of the bioreactor, preparation of the scaffolds and cell seeding, application and maintenance of stretch and shear flow, and sample harvesting for analysis&#8212;are discussed in detail.

The goal of this protocol is to execute a dynamic co-culture of human macrophages and myofibroblasts in tubular electrospun scaffolds to investigate material-driven tissue regeneration, using a bioreactor which enables the decoupling of shear stress and cyclic stretch.

The use of resorbable biomaterials to induce regeneration directly in the body is an attractive strategy from a translational perspective. Such materials ...

Engineering

Generating 3D Spheres and 2D Air-Liquid Interface Cultures of Human Induced Pluripotent Stem Cell-Derived Type 2 Alveolar Epithelial Cells

In the lung, the alveolar epithelium is a physical barrier from environmental stimuli and plays an essential role in homeostasis and disease. Type 2 alveolar epithelial cells (AT2s) are the facultative progenitors of the distal lung epithelium. Dysfunction and injury of AT2s can result from and contribute to various lung diseases. Improved understanding of AT2 biology is, thus, critical for understanding lung biology and disease; however, primary human AT2s are generally difficult to isolate and limited in supply. To overcome these limitations, human induced pluripotent stem cell (iPSC)-derived type 2 alveolar epithelial cells (iAT2s) can be generated through a directed differentiation protocol that recapitulates in vivo lung development. iAT2s grow in feeder-free conditions, share a transcriptomic program with human adult primary AT2s, and execute key functions of AT2s such as production, packaging, and secretion of surfactant. This protocol details the methods for maintaining self-renewing iAT2s through serial passaging in three-dimensional (3D) culture or adapting iAT2s to air-liquid interface (ALI) culture. A single-cell suspension of iAT2s is generated before plating in 3D solubilized basement membrane matrix (hereafter referred to as &#34;matrix&#34;), where they self-assemble into monolayered epithelial spheres. iAT2s in 3D culture can be serially dissociated into single-cell suspensions to be passaged or plated in 2D ALI culture. In ALI culture, iAT2s form a polarized monolayer with the apical surface exposed to air, making this platform readily amenable to environmental exposures. Hence, this protocol generates an inexhaustible supply of iAT2s, producing upwards of 1 x 1030 cells per input cell over 15 passages while maintaining the AT2 program indicated by SFTPCtdTomato expression. The resulting cells represent a reproducible and relevant platform that can be applied to study genetic mutations, model environmental exposures, or screen drugs.

The present protocol describes human induced pluripotent stem cell-derived type 2 alveolar epithelial-like cells (iAT2s). These cells can be cultured as self-renewing spheres in 3D culture or adapted to air-liquid interface (ALI) culture.

In the lung, the alveolar epithelium is a physical barrier from environmental stimuli and plays an essential role in homeostasis and disease. Type 2 ...

Developmental Biology

Generation of Airway Epithelial Cell Air-Liquid Interface Cultures from Human Pluripotent Stem Cells

Diseases of the conducting airway such as asthma, cystic fibrosis (CF), primary ciliary dyskinesia (PCD), and viral respiratory infections are major causes of morbidity and mortality worldwide. In vitro platforms using human bronchial epithelial cells (HBECs) have been instrumental to our understanding of the airway epithelium in health and disease. Access to HBECs from individuals with rare genetic diseases or rare mutations is a bottleneck in lung research.
Induced pluripotent stem cells (iPSCs) are readily generated by "reprogramming" somatic cells and retain the unique genetic background of the individual donor. Recent advances allow for the directed differentiation of iPSCs to lung epithelial progenitor cells, alveolar type 2 cells, as well as the cells of the conducting airway epithelium via basal cells, the major airway stem cells.
Here we outline a protocol for the maintenance and expansion of iPSC-derived airway basal cells (hereafter iBCs) as well as their trilineage differentiation in air-liquid interface (ALI) cultures. iBCs are maintained and expanded as epithelial spheres suspended in droplets of extracellular matrix cultured in a primary basal cell medium supplemented with inhibitors of TGF-&#223; and BMP signaling pathways. iBCs within these epithelial spheres express key basal markers TP63 and NGFR, can be purified by fluorescence activated cell sorting (FACS), and when plated on porous membranes in standard ALI culture conditions, differentiate into a functional airway epithelium. ALI cultures derived from healthy donors are composed of basal, secretory and multiciliated cells and demonstrate epithelial barrier integrity, motile cilia, and mucus secretion. Cultures derived from individuals with CF or PCD recapitulate the dysfunctional CFTR-mediated chloride transport or immotile cilia, the respective disease-causing epithelial defects.
Here, we present a protocol for the generation of human cells that can be applied for modeling and understanding airway diseases.

Recent advances in human induced pluripotent stem cell differentiation protocols allow for the stepwise derivation of organ-specific cell types. Here, we provide detailed steps for the maintenance and expansion of iPSC-derived airway basal cells and their differentiation into a mucociliary epithelium in air-liquid interface cultures.

Diseases of the conducting airway such as asthma, cystic fibrosis (CF), primary ciliary dyskinesia (PCD), and viral respiratory infections are major ...

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 ...

Airway Model for Studying Lung Barrier Integrity, Disease, and Drug Discovery

iPSC-Derived Epithelial, Mesenchymal, Endothelial, and Immune Cell Co-Culture to Model Airway Barrier Integrity in Lung Health and Disease

Human lung tissue is composed of an interconnected network of epithelium, mesenchyme, endothelium, and immune cells from the upper airway of the nasopharynx to the smallest alveolar sac. Interactions between these cells are crucial in lung development and disease, acting as a barrier against harmful chemicals and pathogens. Current in vitro co-culture models utilize immortalized cell lines with different biological backgrounds, which may not accurately represent the cellular milieu or interactions of the lung. We differentiated human iPSCs into 3D lung organoids (containing both epithelium and mesenchyme), endothelial cells, and macrophages. These were co-cultured in an air-liquid interface (ALI) format to form an epithelial/mesenchymal apical barrier invested with macrophages and a basolateral endothelial barrier (iAirway). iPSC-derived iAirways showed a reduction in barrier integrity in response to infection with respiratory viruses and cigarette toxins. This multi-lineage lung co-culture system provides a platform for studying cellular interactions, signaling pathways, and molecular mechanisms underlying lung development, homeostasis, and disease progression. iAirways closely mimic human physiology and cellular interactions, can be generated from patient-derived iPSC&#39;s, and can be customized to include different cell types of the airway. Overall, iPSC-derived iAirway models offer a versatile and powerful tool for studying barrier integrity to better understand genetic drivers for disease, pathogen response, immune regulation, and drug discovery or repurposing in vitro, with the potential to advance our understanding and treatment of airway diseases.

Author Spotlight: Modeling Human Airway Remodeling and Viral Responses Using Isogenic Epithelial, Endothelial, and Immune Cells

This article describes the generation of a complex, multi-cellular airway barrier model composed of induced pluripotent stem cell (iPSC)-derived lung epithelium, mesenchyme, endothelial cells, and macrophages in an air-liquid interface culture.

Imaging-Guided Bioreactor for Generating Bioengineered Airway Tissue

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