Name: culture and imaging of human nasal epithelial organoids
Uploaded: 2021-12-17T14:00:00.000+00:00
Duration: 13 min 20 s
Description: Watch this Scientific Journal Video about Culture and Imaging of Human Nasal Epithelial Organoids at JoVE.com

We gratefully acknowledge the contributions of all the participants who donated HNE brush biopsies to develop this protocol. We thank Latona Kersh and Children&#39;s Research Unit staff for coordinating study volunteer recruitment and sample collections. We thank Lily Deng, Johnathan Bailey, and Stephen Mackay, former trainees in our laboratory, for technical assistance. We thank Zhong Liu and Rui Zhao for their technical help. Steven M. Rowe, Director of the CF Research Center at UAB, provides leadership and resources, without which this work would not be possible. We also would like to thank Sarah Guadiana at Biotek for assistance with instrument training, Robert Grabski for confocal microscopy assistance at the UAB High-Resolution Imaging Facility, and Dezhi Wang for histological assistance at the UAB Histology Core. This work was supported by the National Institutes of Health (NIH.) Grant K23HL143167 (to JSG), Cystic Fibrosis Foundation (CFF) Grant GUIMBE18A0-Q (to JSG), the Gregory Fleming James Cystic Fibrosis Center [NIH Grants R35HL135816 and DK072482 and the CFF University of Alabama at Birmingham (UAB) Research and Development Program (Rowe19RO)], and the UAB Center for Clinical and Translational Science (NIH Grant UL1TR001417).

HNE samples were collected at the Children&#39;s of Alabama hospital. All procedures and methods described here have been approved by the IRB University of Alabama at Birmingham (UAB IRB #151030001). To facilitate the expansion and improve the function of human nasal epithelial cells (HNEs), the present culturing methods are adapted from the well-known air-liquid interface (ALI) culture method28,34. HNEs were initially collected by brush biopsy as previously described12,14, with the only difference being the use of a cytology brush. All sample processing steps and cell culture were performed in the biosafety cabinet.
1. Cell culture and expansion of nasal epithelial cells
<ol>
	<li>After brushing, store the nasal biopsy sample in 8 mL of RPMI media in a 15 mL conical tube on ice and transfer it to the lab within 4 h (not more than 24 h).</li>
	<li>Dissociate the nasal brush biopsy into 8 mL of RPMI media in a 15 mL conical tube by passing the cytology brush through a 1 mL large bore pipette tip (cut off the tip) several times until the brush is clear of tissue.</li>
	<li>Centrifuge the cells at 500 x g for 5 min at 4 &#176;C, remove the supernatant, and then re-suspend the cell pellet in 3 mL of cell detachment solution (see Table of Materials) and incubate at room temperature for 16 min to digest.</li>
	<li>Use 5 mL of expansion media (Table 1) to wash the cells twice. Then, add cells to a T75 flask pre-seeded with irradiated and inactivated 3T3 fibroblasts22 (50%-60% confluence) to co-culture the cells and expand in the expansion media for 7-14 days (see Figure 1 for the appropriate colony). Before use, test the irradiated 3T3 fibroblasts to ensure they cannot proliferate, which will negatively influence the epithelial cell expansion. 
	NOTE: Discard the cells if the nasal epithelial cell confluence does not reach 70% within 14 days.</li>
	<li>For the cells derived from patients with CF, introduce four antibiotics (100 &#181;g/mL of tobramycin, 2.5 &#181;g/mL of amphotericin B, 100 &#181;g/mL of ceftazidime, 100 &#181;g/mL of vancomycin, see Table of Materials) into the expansion media for disinfection of the cells for the first 3 days of culture, and then replace the media with antibiotic-free expansion media changing the media every 2 days. 
	NOTE: This limited use of antibiotics is intended to reduce culture loss due to bacterial colonization while encouraging proliferation after the additional antibiotics are removed. These antibiotics can be tailored to the patient&#39;s specific microbiology results, with sensitivities, if needed.</li>
	<li>Harvest HNEs from co-culture using the double trypsinization method22 once the cells reach approximately 80% confluence. This method ensures that the irradiated and inactivated 3T3 fibroblasts are removed from the flask, and they do not contaminate subsequent organoid seeding.
	<ol>
		<li>Wash the cells with 1x DPBS, and then add 1.5 mL of 0.05% trypsin-EDTA into the T75 flask for 4 min at 37 &#176;C to remove the irradiated and inactivated 3T3 fibroblast from culture.</li>
		<li>Rinse the T75 flask with 1x DPBS twice to thoroughly wash away any remaining 3T3 fibroblasts; add 1.5 mL of 0.05% trypsin-EDTA into the T75 flask for 10 min at 37 &#176;C to detach HNEs.</li>
		<li>Neutralize the trypsin with soybean trypsin inhibitor (see Table of Materials) at a 1:1 ratio. Centrifuge the cells at 500 x g for 5 min, and then remove the supernatant. After washing the cells with 5 mL of expansion media once, the cells are ready for seeding to grow organoids. 
		&#8203;NOTE: HNEs with a passage number of three are recommended to be used for further experiments.</li>
	</ol>
	</li>
</ol>
2. Growth and differentiation of organoids in slides and culture inserts
<ol>
	<li>Thaw the organoid culture extracellular matrix (ECM) overnight at 4 &#176;C. Cool pipette tips to 4 &#176;C and 15-well angiogenesis slides (see Table of Materials) overnight at 4 &#176;C. 
	NOTE: ECM should be put at 4 &#176;C the night before the cell harvesting needs to be done.</li>
	<li>Coat the slides with 5 &#181;L of cold 100% ECM on ice (pipette 5 &#181;L of cold 100% ECM with a cold pipette tip into each well of the 15-well slide), and then place them into a cell culture incubator at 37 &#176;C for at least 30 min for consolidation.</li>
	<li>Count the HNEs harvested from co-culture using a hemocytometer and dilute the cells to 500 cells/&#181;L in total number with 20% ECM diluted by differentiation media (Table 2) on ice. Then, seed 5 &#181;L of the cold cell/ECM mixture into each well of the slides coated with ECM.</li>
	<li>Immediately transfer the slides into a culture incubator at 37 &#176;C for 1 h to consolidate the cell/ECM mixture.</li>
	<li>Feed the cells in each well of the 15-well angiogenesis slides with 50 &#181;L of differentiation media. Change the media every other day until the organoids are ready for further experiments. 
	NOTE: Organoids can usually be visualized after 1-2 days. There are 20-90 organoids commonly formed in each well of the slides. The organoids can typically survive for 40-60 days in ECM with feeding every other day when kept in a humidified incubator at 37 &#176;C.</li>
	<li>Culture the organoids in the culture inserts (see Table of Materials) for greater quantity for specific applications (such as sectioning for histology or immunofluorescence) as per the steps mentioned below.
	<ol>
		<li>To grow organoids in the culture inserts, prepare the organoid culture ECM, pipette tips, and inserts as mentioned in steps 2.1-2.2. Coat the inserts with 100 &#181;L of cold 100% ECM.</li>
		<li>Seed 60 &#181;L of cell/ECM mixture (500 cells/&#181;L with 20% ECM in differentiation media) on top of the ECM coating in the insert. 
		NOTE: All the other steps for making organoids in the inserts are the same as those conducted in the slides, steps 2.4-2.5.</li>
		<li>Add 600 &#181;L of the differentiation media into the bottom part of the insert. Change the media every alternate day until the organoids are used for experiments, typically 2-3 weeks.</li>
	</ol>
	</li>
</ol>
3. Preparation and isolation of organoids for whole-mount immunofluorescence
<ol>
	<li>Isolation and fixation of organoids
	<ol>
		<li>Pre-treat 8-well glass-bottom chamber slides with cell adhesive (see Table of Materials) for 30 min following Reference35. After discarding the solution, air-dry the wells for 30 min.</li>
		<li>To harvest the organoids, remove the media from the top of the ECM, and then add 50 &#181;L of cold 1x PBS into each well of the 15-well slides on ice (1:1 with ECM volume).</li>
		<li>Pipette up and down 3-5 times using 200 &#181;L of the large-bore pipette tip, and then dispense the solution onto the center of a well of the 8-well chamber slides.</li>
		<li>Immediately remove excess liquid from the wells by a fine-tip pipette. Then, place the chamber slide into a 37 &#176;C incubator for 40 min to enhance the organoid adhering to the glass bottom.</li>
		<li>After gently washing with 1x PBS twice, fix the organoids with 300 &#181;L of 4% paraformaldehyde in each well for 30 min at room temperature (RT).</li>
		<li>Wash twice with 1x PBS and store the organoids in 1x PBS at 4 &#176;C for immunostaining for up to 2 weeks.</li>
	</ol>
	</li>
	<li>Immunofluorescence staining
	<ol>
		<li>To reduce auto-fluorescence, add 250 &#181;L of 50 mM NH4Cl in 1x PBS into each well of the slides at RT for 30 min while gently shaking on a shaker at 20 rpm.</li>
		<li>After washing with 1x PBS twice, permeabilize the cells with 0.1% Triton X-100 for 30 min at RT while gently shaking at 20 rpm.</li>
		<li>After washing with 1x PBS twice, add 300 &#181;L of blocking solution, including 5% BSA and 0.1% Triton X-100 in 1x PBS, into each well for 1 h at RT.</li>
		<li>Following washing, add primary antibody into the appropriate wells followed by secondary antibodies (see Table of Materials). Prepare primary and secondary antibody solutions in 2% BSA and 0.3% Triton X-100 in 1x PBS. Incubate all the primary antibodies at 4 &#176;C for 2 days and all the secondary antibodies at 4 &#176;C for 1 day. 
		NOTE: The final concentrations are dependent on the protein desired for the experiment. Please check the stock concentration on the manufacturer&#39;s datasheet. Then, calculate the final concentrations based on the dilutions used in the Table of Materials.</li>
		<li>After incubation, wash the wells thoroughly with 1x PBS and add DAPI in 2% BSA and 0.3% Triton X-100 into each well for nuclear staining.</li>
		<li>Image the organoids using a confocal laser scanning microscope, with a 20-60x oil immersion objective (see Table of Materials). Use Z-stack mode to set the upper and lower bounds of the image and use the recommended optimal Z-step size determined by the confocal software. 
		&#8203;NOTE: The following four confocal laser excitation wavelengths were used: 408.7 nm, 489.1 nm, 561.7 nm, and 637.9 nm.</li>
	</ol>
	</li>
</ol>
4. Preparation and isolation of organoids for histological sectioning
<ol>
	<li>To harvest the organoids for histological studies, remove the media from the culture and add 50 &#181;L of cold 1x PBS into each well of the slides on ice.</li>
	<li>Pipette up and down three to five times using a 200 &#181;L large-bore pipette tip, combine all the solutions from the 15-well slide or culture inserts into a 15 mL conical tube on ice. Adjust the total solution volume in the tube to 10 mL by adding additional cold 1x PBS.</li>
	<li>Centrifuge the tube at 4 &#176;C, 300 x g for 5 min; aspirate out the supernatant and add 60 &#181;L of warm histogel (see Table of Materials) to mix with the organoid pellet using a 200 &#181;L of the large-bore pipette tip.</li>
	<li>Immediately transfer the suspension into a histology mold. After the consolidation of the histogel at room temperature, put the mold block into 4% paraformaldehyde for fixation overnight at 4 &#176;C.</li>
	<li>After embedding in paraffin, cut the histogel block into 5 &#181;m cross-sections (for example, with a microtome), fix the sections onto glass slides, and stain using hematoxylin and eosin (H&#38;E) or immunofluorescence-labeled antibodies. Take images using a bright-field microscope or inverted epi-fluorescence microscope (see Table of Materials).</li>
</ol>
5. Imaging of live organoids
NOTE: The following steps are carried out using an automated imaging system (see Table of Materials). Different imaging systems need to adapt these steps following their specific manufacturer&#39;s instructions. Regardless of the equipment utilized, imaging live organoids require a temperature-controlled and humidified environmental chamber with an accompanied CO2 gas controller.
<ol>
	<li>To monitor the differentiation of organoids before performing a functional swelling assay36, capture the whole slide images manually with any bright-field microscope or with an automated imaging system, as detailed below.
	<ol>
		<li>Turn on the power to the automated imaging system and the CO2/O2 gas controller and allow the system to complete the automated calibration (~30 min).</li>
		<li>After completion, set the imaging system temperature to 37 &#176;C; add 15 mL of sterile water to the humidification reservoir, open the CO2 valves, close the lids, and let the imaging system pre-incubate for a minimum of 30 min before imaging.</li>
		<li>Open the automated imaging software to set up a protocol for imaging and choose the location to save the raw experimental data. 
		NOTE: An example protocol file (Supplementary File 1), specific to the imaging system, has been provided as a template for the automated imaging of organoids to monitor organoid differentiation.</li>
		<li>Once the environmental settings are met, immediately transfer up to two 15-well slides with lids into the environmental chamber in the automated image system&#39;s slide holder insert (see Table of Materials).</li>
		<li>Select the wells to be imaged and start imaging to complete imaging of the desired wells. 
		NOTE: The basic recommended settings are: 4x and 10x air objective, bright-field channel, 2 x 2 montage for 4x objective to cover the whole well area, 4 x 4 montage for a 10x objective. Z-stack settings: three to six Z-stack slices, Z-step size = 50-100 &#181;m, one to two slices below autofocus point, and three to five slices above.</li>
	</ol>
	</li>
	<li>To image live organoids for use in a forskolin-induced swelling (FIS) assay, use a microscope with an automated stage equipped with an environmentally controlled imaging chamber that allows temperature and CO2 control.
	<ol>
		<li>Begin with steps 5.1.1-5.1.4, substitute the protocol file in step 5.1.3 with the one provided in the example protocol file (Supplementary File 2) containing settings specific for performing a FIS assay.</li>
		<li>Before each experiment, adjust the exposure settings by evaluating at least three wells for each slide (far left, middle, and far right). Apply the X and Y coordinate offsets to ensure that the objective will be in the center of the well for all the wells. 
		NOTE: The basic recommended settings are: 4x air objective, Channel 1 = Bright Field, Channel 2 = DAPI, 2 x 2 montage (four image tiles). Z-stack settings: three to four Z-stack slices, Z-step size = 50-100 &#181;m, one slice below autofocus point and two to three slices above. Image acquisition time: 8 h with 20 min intervals (Total reads = 25; Read 1 is T = 0).</li>
		<li>Once all the settings are appropriately selected, choose the wells to image for both slides, or select all, and begin the run as per the equipment&#39;s instructions.</li>
		<li>After running, save the experiment, close the imaging software, and shut down the imaging system37.</li>
	</ol>
	</li>
</ol>
6. Baseline lumen measurements
NOTE: This is done using manual imaging analysis software (see Table of Materials). A similar methodology can be followed using an open-source software38 or any software that can measure the area of a region on an image.
<ol>
	<li>In the software, open the Automated Measurement Panel by right-clicking the bottom of the screen, select Measurement, and then Automated Measurement Results. The area of each region of interest (ROI) measured will appear there.</li>
	<li>Open the organoid image in the software and select 5-10 organoids with visible lumens. 
	NOTE: Lumens will be a circular area in the middle of the organoid that is visibly different in color than the rest of the organoid (Figure 2A).</li>
	<li>Using the polygon ROI measurement feature, hold right click on the image to open the menu and select Polygonal ROI to outline the full organoid to obtain the organoid&#39;s total surface area (TSA). Then using the same feature, outline the lumen area (LA) (Figure 2B).</li>
	<li>Repeat for the remaining organoids in the well and all the wells in the assay.</li>
	<li>Export the data to excel. Divide the LA by the TSA and average all the organoids from the sample to get the Baseline Lumen Ratio (BLR)11,13. 
	&#8203;NOTE: Typically, ~87% of non-CF organoids will have a BLR over 0.6, and 97% will be over 0.5, while only 14% of CF organoids will have a BLR over 0.6, and 31% will be over 0.5.</li>
</ol>
7. Pre-treatment and automated imaging of HNE organoids
NOTE: All pre-treatment steps are carried out in a clean biosafety cabinet. Pre-setup the automated imaging system and the software for recording the assay before step 7.1. The incubation with DAPI is optional but is recommended as a fail-safe if the quality of bright field images is compromised. In this case, the DAPI channel (377 nm) can be analyzed instead.
<ol>
	<li>Pre-incubate the organoids in a well of 15-well slides with 50 &#181;L of differentiation media containing DAPI with or without 100 &#181;M of CFTRinh-172 (see Table of Materials) in a 37 &#176;C incubator for 1 h. While the organoids incubate, perform step 5.2.1 using a swelling assay custom protocol following Supplementary File 2.</li>
	<li>Remove the pre-incubation medium using a glass Pasteur pipette with aspiration. Add 10 &#181;M of forskolin and 100 &#181;M of IBMX (stimulation cocktails) (see Table of Materials) for a total volume of 50 &#181;L differentiation media into each well. 
	NOTE: Ensure no bubbles are introduced into the wells. Bubbles on the images will affect the automated analysis.</li>
	<li>Without delay, begin the FIS imaging protocol following step 5.2.2 to 5.2.4. Acquire images every 20 min in each well, with a total runtime of 8 h.</li>
</ol>
8. Automated analysis of forskolin-induced swelling assay on HNE organoids
<ol>
	<li>Open the automated imaging analysis software, find the experiment previously saved from step 7.3, and bring up the images of the experiment.</li>
	<li>Choose the vessel window to be evaluated and select the option containing the processed images that have been stitched and z-projected. This step should provide a picture of the entire well with all organoids within the imaging frame for masking assessment and measurements.</li>
	<li>Choose the well image to assess. Select Analyze. Repeat this process for other images to ensure appropriate masking for all images included in the automated measurements. Save the setting parameters.</li>
	<li>After completing the analyzing settings, apply the changes. The software will change the measurements based on the settings. 
	NOTE: After the initial image pre-processing is complete, quality control (QC) measures should be performed to ensure consistent masking. These include manually reviewing all wells to ensure organoids are within the imaging frame, bubbles or debris are not inappropriately masked, and checking the masking in bright field and DAPI channels.</li>
	<li>Export the data for the summary analysis (including graphing and statistical analysis).</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+keratinocytes+are+efficiently+immortalized+by+a+Rho+kinase+inhibitor.">Human keratinocytes are efficiently immortalized by a Rho kinase inhibitor.</a> Journal of Clinical Investigation. 120 (7), 2619-2626 (2010).</li><li>Liu, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ROCK+inhibitor+and+feeder+cells+induce+the+conditional+reprogramming+of+epithelial+cells.">ROCK inhibitor and feeder cells induce the conditional reprogramming of epithelial cells.</a> The American Journal of Pathology. 180 (2), 599-607 (2012).</li><li>Palechor-Ceron, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Radiation+induces+diffusible+feeder+cell+factor(s)+that+cooperate+with+ROCK+inhibitor+to+conditionally+reprogram+and+immortalize+epithelial+cells.">Radiation induces diffusible feeder cell factor(s) that cooperate with ROCK inhibitor to conditionally reprogram and immortalize epithelial cells.</a> The American Journal of Pathology. 183 (6), 1862-1870 (2013).</li><li>Scudieri, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ionocytes+and+CFTR+chloride+channel+expression+in+normal+and+cystic+fibrosis+nasal+and+bronchial+epithelial+cells.">Ionocytes and CFTR chloride channel expression in normal and cystic fibrosis nasal and bronchial epithelial cells.</a> Cells. 9 (9), (2020).</li><li>Marthin, J. K., Stevens, E. M., Larsen, L. A., Christensen, S. T., Nielsen, K. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patient-specific+three-dimensional+explant+spheroids+derived+from+human+nasal+airway+epithelium:+a+simple+methodological+approach+for+ex+vivo+studies+of+primary+ciliary+dyskinesia.">Patient-specific three-dimensional explant spheroids derived from human nasal airway epithelium: a simple methodological approach for ex vivo studies of primary ciliary dyskinesia.</a> Cilia. 6, 3 (2017).</li><li>Blouquit, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ion+and+fluid+transport+properties+of+small+airways+in+cystic+fibrosis.">Ion and fluid transport properties of small airways in cystic fibrosis.</a> American Journal of Respiratory Cell and Molecular Biology. 174 (3), 299-305 (2006).</li><li>Birket, S. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Combination+therapy+with+cystic+fibrosis+transmembrane+conductance+regulator+modulators+augment+the+airway+functional+microanatomy.">Combination therapy with cystic fibrosis transmembrane conductance regulator modulators augment the airway functional microanatomy.</a> The American Journal of Physiology - Lung Cellular and Molecular Physiology. 310 (10), 928-939 (2016).</li><li>Birket, S. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+functional+anatomic+defect+of+the+cystic+fibrosis+airway.">A functional anatomic defect of the cystic fibrosis airway.</a> American Journal of Respiratory and Critical Care Medicine. 190 (4), 421-432 (2014).</li><li>Chu, K. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Particle-tracking+microrheology+using+micro-optical+coherence+tomography.">Particle-tracking microrheology using micro-optical coherence tomography.</a> Biophysical Journal. 111 (5), 1053-1063 (2016).</li><li>Chu, K. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=et+al.+In+vivo+imaging+of+airway+cilia+and+mucus+clearance+with+micro-optical+coherence+tomography.">et al. In vivo imaging of airway cilia and mucus clearance with micro-optical coherence tomography.</a> Biomedical Optics Express. 7 (7), 2494-2505 (2016).</li><li>Liu, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Method+for+quantitative+study+of+airway+functional+microanatomy+using+micro-optical+coherence+tomography.">Method for quantitative study of airway functional microanatomy using micro-optical coherence tomography.</a> PLoS One. 8 (1), 54473 (2013).</li><li>Tuggle, K. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+defects+in+ion+transport+and+tissue+development+in+cystic+fibrosis+transmembrane+conductance+regulator+(CFTR)-knockout+rats.">Characterization of defects in ion transport and tissue development in cystic fibrosis transmembrane conductance regulator (CFTR)-knockout rats.</a> PLoS One. 9 (3), 91253 (2014).</li><li>McCravy, M. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Personalised+medicine+for+non-classic+cystic+fibrosis+resulting+from+rare+CFTR+mutations.">Personalised medicine for non-classic cystic fibrosis resulting from rare CFTR mutations.</a> European Respiratory Journal. 56 (1), 2000062 (2020).</li><li>Mutyam, V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Therapeutic+benefit+observed+with+the+CFTR+potentiator,+ivacaftor,+in+a+CF+patient+homozygous+for+the+W1282X+CFTR+nonsense+mutation.">Therapeutic benefit observed with the CFTR potentiator, ivacaftor, in a CF patient homozygous for the W1282X CFTR nonsense mutation.</a> Journal of Cystic Fibrosis. 16 (1), 24-29 (2017).</li><li>Corning Inc. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> CORNING CELL-TAK CELL AND TISSUE ADHESIVE. , (2013).</li><li>Anderson, J. D., Liu, Z., Odom, L. V., Kersh, L., Guimbellot, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CFTR+function+and+clinical+response+to+modulators+parallel+nasal+epithelial+organoid+swelling.">CFTR function and clinical response to modulators parallel nasal epithelial organoid swelling.</a> The American Journal of Physiology - Lung Cellular and Molecular Physiology. 321 (1), 119-129 (2021).</li><li>Biotek Instruments, Incorporated. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Lionheart FX Live Cell Imager Operator's Manual. , (2016).</li><li>Schneider, C. A., Rasband, W. S., Eliceiri, K. 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E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+and+in+vitro+ivacaftor+response+in+cystic+fibrosis+patients+with+residual+CFTR+function:+N-of-1+studies.">In vivo and in vitro ivacaftor response in cystic fibrosis patients with residual CFTR function: N-of-1 studies.</a> Pediatric Pulmonology. 52 (4), 472-479 (2017).</li><li>Garratt, L. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determinants+of+culture+success+in+an+airway+epithelium+sampling+program+of+young+children+with+cystic+fibrosis.">Determinants of culture success in an airway epithelium sampling program of young children with cystic fibrosis.</a> Experimental Lung Research. 40 (9), 447-459 (2014).</li></ol>

JSG is listed as an inventor on a patent application 20170242033 from the University of North Carolina that describes a similar model. When licensed technology from UNC produces royalties, the inventors receive a share of the revenue. Otherwise, the authors declare no conflicts of interest. The funders had no role in the study's design, in the collection, analyses, or interpretation of data, in the writing of the manuscript, or in the decision to publish the results.

This manuscript provides detailed methodologies for comprehensive live and fixed imaging of the airway epithelial organoids derived from HNE brush biopsy. It describes functional assays that can determine CFTR activity in an individual. HNEs provide a minimally invasive, primary tissue for a variety of applications. The expansion techniques offered here can be used for modeling airways disease, including organoids. Organoids can be used for precision therapeutic approaches and to monitor the stability of gene or mRNA-bas...

Introduction to the technique 
Ex vivo culture-based assays are an increasingly utilized tool for precision medicine and the study of disease pathophysiology. Primary human nasal epithelial (HNE) cell culture has been used in numerous studies of cystic fibrosis1,2,3,4,5,6,7,8,9,10,11,12,13, an autosomal recessive disease that affects epithelial cell function in multiple organs. HNE culture provides a renewable source of airway epithelia that may be obtained prospectively and recapitulates electrophysiological and biochemical qualities to test Cystic Fibrosis Transmembrane conductance Regulator (CFTR) activity. HNE cells can be sampled with minimal side effects14, similar to common viral respiratory swabs. Research work describing a model for cystic fibrosis study derived from HNE brush biopsies has been recently published11,13. While similar to other models using primary HNE2,3 and intestinal tissue15,16,17,18,19, detailed characterization of the differentiation and imaging of this model are described here for use in CF research and for aiding in the studies of other airways diseases13. The organoid model is not unlimited like immortalized cell lines but can be expanded by conditional reprogramming (using irradiated and inactivated feeder fibroblasts and Rho-kinase inhibitors) to a more stem cell-like state20,21,22,23. The processing of HNE brush biopsies using this method yields large numbers of epithelial cells for use in multiple applications at higher throughput while still retaining the ability to differentiate fully. While this protocol was developed using feeder cells, other methodologies may be used by investigators wishing to avoid feeder cell technology14,24.
Importance of the technique to pulmonary biology 
A significant study has been devoted to understanding how the absence of regular, functioning CFTR in the cell membrane of epithelial cells results in dysfunction in the lungs, pancreas, liver, intestine, or other tissues. Dysfunctional epithelial ion transport, particularly that of chloride and bicarbonate, results in a decreased volume of the epithelial lining fluids and changes in mucous secretions, leading to mucous stasis and obstruction. In other airway diseases, such as primary ciliary dyskinesia, altered ciliary motion impairs mucociliary clearance and leads to mucous stasis and obstruction25. Therefore, the current HNE organoid model has been developed for various applications, depending on the investigator&#39;s experimental design and resources. This includes live-cell imaging using live-cell stains; fixation and sectioning to characterize the morphology; immunofluorescence staining with antibodies and whole-mount confocal imaging to avoid disrupting intraluminal structures; and bright-field imaging and micro-optical coherence tomography for quantitative measurements of ciliary beat frequency and mucociliary transport13. To facilitate expansion to other investigators, commercially available reagents and supplies were used for culturing. A functional assay was developed that used common microscope techniques and more specialized equipment. Overall, while the present model was designed to assess CFTR activity at baseline or in response to therapeutics, the techniques described in this protocol can be applied to other diseases involving epithelial cell function, especially epithelial cell fluid transport.
Comparison to other methodologies 
Recently the utility of this organoid model was developed by correlating in vitro CFTR modulator responses of patients&#39; organoids with their clinical response11. Notably, it is also demonstrated that the present model paralleled short-circuit current responses, the current gold standard for assessing CFTR function, in the same patients. Short-circuit current differs from the swelling assay because the former measures CFTR function via ion transport26. In contrast, this assay measures a more downstream effect with fluid transport, providing additional information about the overall function of CFTR27,28,29,30,31,32. Short-circuit current measurements have continued to be a common and reliable method for determining CFTR chloride channel activity1,33. These electrophysiological assays require specialized, expensive equipment, require many times more cells for each experimental replicate than the organoid assay, cannot be easily automated, and are not amenable to scaling up for higher throughput applications. Another organoid model derived from intestinal epithelia has additional advantages15,16,17,18, such as more excellent replicative capability, but is neither derived from an airway tissue nor is universally available. HNE brushings are obtained with inexpensive cytology brushes without the need for sedation and at minimal risk. Getting the brushing does not require a clinician and can be performed by trained research coordinators and other research staff14. The HNE organoid model can be cultured by any laboratory with primary cell culture capabilities, and some of the applications can be performed with standard microscopy techniques. Altogether, these advantages provide additional access to technology for assessing airway epithelial function that might otherwise be unavailable to some laboratories. Furthermore, HNE organoids can be utilized to study other disease states that affect the airway, such as primary ciliary dyskinesia25 or viral infection, which intestinal organoids cannot.

<table><tbody><tr><td>Nasal brush</td><td>Medical Packaging CYB1</td><td>CYB-1</td><td>Length: 8 inches, width approximately 7 mm</td></tr><tr><td>Large-Orifice Pipette Tips</td><td>ThermoFisher Scientific</td><td>02-707-141</td><td>Large bore pipette tips</td></tr><tr><td>Accutase</td><td>ThermoFisher Scientific</td><td>A1110501</td><td>Cell detachment solution</td></tr><tr><td>0.05% trypsin -EDTA</td><td>Gibco</td><td>25300-054</td><td></td></tr><tr><td>Trypsin inhibitor from soybean</td><td>Sigma</td><td>T6522</td><td>Working solution: 1mg/mL in 1XDPBS</td></tr><tr><td>Matrigel matrix</td><td>Corning</td><td>356255</td><td>Extracellular matrix (EM)</td></tr><tr><td>&amp;micro;-Slide Angiogenesis</td><td>Ibidi</td><td>81506</td><td>15-well slide</td></tr><tr><td>24-Well Transwell</td><td>Corning</td><td>7200154</td><td>Culture insert</td></tr><tr><td>Chambered Coverglass</td><td>ThermoFisher Scientific</td><td>155409</td><td>8-well glass-bottom chamber slides</td></tr><tr><td>Cell-Tak Cell and Tissue Adhesive</td><td>ThermoFisher Scientific</td><td>354240</td><td>Cell adhesive</td></tr><tr><td>Paraformaldehyde</td><td>Electron Microscopy Sciences</td><td>50980487</td><td></td></tr><tr><td>Triton X-100</td><td>Alfa Aesar</td><td>A16046</td><td></td></tr><tr><td>BSA</td><td>ThermoFisher Scientific</td><td>BP1600-100</td><td></td></tr><tr><td>NucBlue</td><td>ThermoFisher Scientific</td><td>R37605</td><td>DAPI</td></tr><tr><td>Eclipse Ts2-FL (Inverted Routine Microscope)</td><td>Nikon</td><td></td><td>Inverted epi-fluorescence microscope or bright-field microscope</td></tr><tr><td>Nikon A1R-HD25</td><td>Nikon</td><td></td><td>Confocal microscope</td></tr><tr><td>NIS Elements- Basic Research</td><td>Nikon</td><td></td><td>manual imaging analysis software</td></tr><tr><td>Histogel</td><td>ThermoFisher Scientific</td><td>HG-4000-012</td><td></td></tr><tr><td>Disposable Base Molds</td><td>ThermoFisher Scientific</td><td>41-740</td><td></td></tr><tr><td>Lionheart FX</td><td>BioTek</td><td>BTLFX</td><td>Automated image system</td></tr><tr><td>Lionheart Cover</td><td>BioTek</td><td>BT1450009</td><td>Environmental Control Lid</td></tr><tr><td>Humidity Chamber</td><td>BioTek</td><td>BT1450006</td><td>Stage insert (environmental chamber)</td></tr><tr><td>Gas Controller for CO&lt;sub&gt;2&lt;/sub&gt; and O&lt;sub&gt;2&lt;/sub&gt;</td><td>BioTek</td><td>BT1210013</td><td>Gas controller</td></tr><tr><td>Microplate/Slide Stage Insert</td><td>BioTek</td><td>BT1450527</td><td>Slide holder</td></tr><tr><td>Gen5 Imaging Prime Software</td><td>BioTek</td><td>BTGEN5IPRIM</td><td>Automated imaging analysis software</td></tr><tr><td>4x Phase Contrast Objective</td><td>BioTek</td><td>BT1320515</td><td></td></tr><tr><td>10x Phase Contrast Objective</td><td>BioTek</td><td>BT1320516</td><td></td></tr><tr><td>LED Cube</td><td>BioTek</td><td>BT1225007</td><td></td></tr><tr><td>Filter Cube (DAPI)</td><td>BioTek</td><td>BT1225100</td><td>DAPI</td></tr><tr><td>CFTRinh-172</td><td>Selleck Chemicals</td><td>S7139</td><td></td></tr><tr><td>Forskolin</td><td>Sigma</td><td>F6886</td><td></td></tr><tr><td>IBMX</td><td>Sigma</td><td>I5879</td><td></td></tr><tr><td>&lt;strong&gt;Expansion Media&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>DMEM</td><td>ThermoFisher Scientific</td><td>11965</td><td></td></tr><tr><td>F12 Nutrient mix</td><td>ThermoFisher Scientific</td><td>11765</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>ThermoFisher Scientific</td><td>&amp;nbsp;16140-071</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>ThermoFisher Scientific</td><td>&amp;nbsp;15-140-122</td><td></td></tr><tr><td>Cholera Toxin</td><td>Sigma</td><td>&amp;nbsp;C8052</td><td></td></tr><tr><td>Epidermal Growth Factor (EGF)</td><td>ThermoFisher Scientific</td><td>&amp;nbsp;PHG0314</td><td></td></tr><tr><td>Hydrocortisone (HC)</td><td>Sigma</td><td>&amp;nbsp;H0888</td><td></td></tr><tr><td>Insulin</td><td>Sigma</td><td>&amp;nbsp;I9278</td><td></td></tr><tr><td>Adenine</td><td>Sigma</td><td>&amp;nbsp;A2786</td><td></td></tr><tr><td>Y-27632</td><td>Stemgent</td><td>&amp;nbsp;04-0012-02</td><td></td></tr><tr><td>&lt;strong&gt;Antibiotic Media&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Ceftazidime</td><td>Alfa Aesar</td><td>&amp;nbsp;J66460-03</td><td></td></tr><tr><td>Tobramycin</td><td>Alfa Aesar</td><td>&amp;nbsp;J67340</td><td></td></tr><tr><td>Vancomycin</td><td>Alfa Aesar</td><td>&amp;nbsp;J67251</td><td></td></tr><tr><td>Amphotericin B</td><td>Sigma</td><td>&amp;nbsp;A2942</td><td></td></tr><tr><td>&lt;strong&gt;Differentiation Media&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>DMEM/F-12 (1:1)</td><td>ThermoFisher Scientific</td><td>&amp;nbsp;11330-32</td><td></td></tr><tr><td>Ultroser-G</td><td>Pall</td><td>&amp;nbsp;15950-017</td><td></td></tr><tr><td>Fetal Clone II</td><td>Hyclone</td><td>&amp;nbsp;SH30066.03</td><td></td></tr><tr><td>Bovine Brain Extract</td><td>Lonza</td><td>&amp;nbsp;CC-4098</td><td></td></tr><tr><td>Insulin</td><td>Sigma</td><td>&amp;nbsp;I-9278</td><td></td></tr><tr><td>Hydrocortisone</td><td>Sigma</td><td>&amp;nbsp;H-0888</td><td></td></tr><tr><td>Triiodothyronine</td><td>Sigma</td><td>&amp;nbsp;T-6397</td><td></td></tr><tr><td>Transferrin</td><td>Sigma</td><td>&amp;nbsp;T-0665</td><td></td></tr><tr><td>Ethanolamine</td><td>Sigma</td><td>&amp;nbsp;E-0135</td><td></td></tr><tr><td>Epinephrine</td><td>Sigma</td><td>E-4250</td><td></td></tr><tr><td>&lt;em&gt;O&lt;/em&gt;-Phosphorylethanolamine</td><td>Sigma</td><td>P-0503</td><td></td></tr><tr><td>Retinoic Acid</td><td>Sigma</td><td>R-2625</td><td></td></tr><tr><td>&lt;strong&gt;Primary antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Human CFTR antibody</td><td>R&amp;amp;D Systems</td><td>MAB1660</td><td>Dilution: 100x</td></tr><tr><td>ZO-1 antibody</td><td>Thermo Fisher</td><td>MA3-39100-A647</td><td>Dilution: 1000x</td></tr><tr><td>Anti-MUC5B antibody</td><td>Sigma</td><td>HPA008246</td><td>Dilution: 100x</td></tr><tr><td>Anti-acetylated tubulin</td><td>Sigma</td><td>T7451</td><td>Dilution: 100x</td></tr><tr><td>Anti-beta IV Tubulin antibody</td><td>Abcam</td><td>Ab11315</td><td>Dilution: 100x</td></tr><tr><td>&lt;strong&gt;Secondary antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Donkey anti-Mouse IgG (H+L), Alexa Fluor 488</td><td>Invitrogen</td><td>A21202</td><td>Dilution: 2000x</td></tr><tr><td>Donkey anti-Rabbit IgG (H+L), Alexa Fluor 594</td><td>Invitrogen</td><td>A21207</td><td>Dilution: 2000x</td></tr></tbody></table>

culture and imaging of human nasal epithelial organoids

Individualized therapy for cystic fibrosis (CF) patients can be achieved with an in vitro disease model to understand baseline Cystic Fibrosis Transmembrane conductance Regulator (CFTR) activity and restoration from small molecule compounds. Our group recently focused on establishing a well-differentiated organoid model directly derived from primary human nasal epithelial cells (HNE). Histology of sectioned organoids, whole-mount immunofluorescent staining, and imaging (using confocal microscopy, immunofluorescent microscopy, and bright field) are essential to characterize organoids and confirm epithelial differentiation in preparation for functional assays. Furthermore, HNE organoids produce lumens of varying sizes that correlate with CFTR activity, distinguishing between CF and non-CF organoids. In this manuscript, the methodology for culturing HNE organoids are described in detail, focusing on the assessment of differentiation using the imaging modalities, including the measurement of baseline lumen area (a method of CFTR activity measurement in organoids that any laboratory with a microscope can employ) as well as the developed automated approach to a functional assay (which requires more specialized equipment).

HNEs expansion is essential for a thriving organoid culture. HNEs from a successful sample collection should expand to over 70% confluence around 10 days. An example of successful and unsuccessful samples is shown in Figure 1A and Figure 1B, respectively. The cells must be discarded if they cannot reach 70% confluence by 14 days after co-culture with irradiated 3T3 cells. Any contaminated cells are to be immediately discarded if unable to rescue...

Watch this Scientific Journal Video about Culture and Imaging of Human Nasal Epithelial Organoids at JoVE.com

Culture and Imaging of Human Nasal Epithelial Organoids

A detailed protocol is presented here to describe an in vitro organoid model from human nasal epithelial cells. The protocol has options for measurements requiring standard laboratory equipment, with additional possibilities for specialized equipment and software.

Individualized therapy for cystic fibrosis (CF) patients can be achieved with an in vitro disease model to understand baseline Cystic Fibrosis ...

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3.6K Views.  University of Alabama at Birmingham (UAB). A detailed protocol is presented here to describe an in vitro organoid model from human nasal epithelial cells. The protocol has options for measurements requiring standard laboratory equipment, with additional possibilities for specialized equipment and software.

Video: Culture and Imaging of Human Nasal Epithelial Organoids

Cultivate Primary Nasal Epithelial Cells from Children and Reprogram into Induced Pluripotent Stem Cells

Nasal epithelial cells (NECs) are the part of the airways that respond to air pollutants and are the first cells infected with respiratory viruses. They are also involved in many airway diseases through their innate immune response and interaction with immune and airway stromal cells. NECs are of particular interest for studies in children due to their accessibility during clinical visits. Human induced pluripotent stem cells (iPSCs) have been generated from multiple cell types and are a powerful tool for modeling human development and disease, as well as for their potential applications in regenerative medicine. This is the first protocol to lay out methods for successful generation of iPSCs from NECs derived from pediatric participants for research purposes. It describes how to obtain nasal epithelial cells from children, how to generate primary NEC cultures from these samples, and how to reprogram primary NECs into well-characterized iPSCs. Nasal mucosa samples are useful in epidemiological studies related to the effects of air pollution in children, and provide an important tool for studying airway disease. Primary nasal cells and iPSCs derived from them can be a tool for providing unlimited material for patient-specific research in diverse areas of airway epithelial biology, including asthma and COPD research.

This publication demonstrates methods for successful sampling and culture of nasal epithelial mucosa from children, and reprogramming these cells to induced Pluripotent Stem Cells (iPSCs).

Nasal epithelial cells (NECs) are the part of the airways that respond to air pollutants and are the first cells infected with respiratory viruses. They ...

Developmental Biology

Three-Dimensional Imaging of Organoids to Study Primary Ciliogenesis During ex vivo Organogenesis

Organoids are stem cell-derived three-dimensional structures that reproduce ex vivo the complex architecture and physiology of organs. Thus, organoids represent useful models to study the mechanisms that control stem cell self-renewal and differentiation in mammals, including primary ciliogenesis and ciliary signaling. Primary ciliogenesis is the dynamic process of assembling the primary cilium, a key cell signaling center that controls stem cell self-renewal and/or differentiation in various tissues. Here we present a comprehensive protocol for the immunofluorescence staining of cell lineage and primary cilia markers, in whole-mount mouse mammary organoids, for light sheet microscopy. We describe the microscopy imaging method and an image processing technique for the quantitative analysis of primary cilium assembly and length in organoids. This protocol enables a precise analysis of primary cilia in complex three-dimensional structures at the single cell level. This method is applicable for immunofluorescence staining and imaging of primary cilia and ciliary signaling in mammary organoids derived from normal and genetically modified stem cells, from healthy and pathological tissues, to study the biology of the primary cilium in health and disease.

Three-Dimensional Imaging of Organoids to Study Primary Ciliogenesis During ex vivo Organogenesis

Stem cell-derived organoids facilitate the analysis of molecular and cellular processes that regulate stem cell self-renewal and differentiation during organogenesis in mammalian tissues. Here we present a protocol for the analysis of the biology of the primary cilium in mouse mammary organoids.

Organoids are stem cell-derived three-dimensional structures that reproduce ex vivo the complex architecture and physiology of organs. Thus, organoids ...

Genetics

Generating and Imaging Mouse and Human Epithelial Organoids from Normal and Tumor Mammary Tissue Without Passaging

Organoids are a reliable method for modeling organ tissue due to their self-organizing properties and retention of function and architecture after propagation from primary tissue or stem cells. This method of organoid generation forgoes single-cell differentiation through multiple passages and instead uses differential centrifugation to isolate mammary epithelial organoids from mechanically and enzymatically dissociated tissues. This protocol provides a streamlined technique for rapidly producing small and large epithelial organoids from both mouse and human mammary tissue in addition to techniques for organoid embedding in collagen and basement extracellular matrix. Furthermore, instructions for in-gel fixation and immunofluorescent staining are provided for the purpose of visualizing organoid morphology and density. These methodologies are suitable for myriad downstream analyses, such as co-culturing with immune cells and ex vivo metastasis modeling via collagen invasion assay. These analyses serve to better elucidate cell-cell behavior and create a more complete understanding of interactions within the tumor microenvironment.

This protocol discusses an approach for generating epithelial organoids from primary normal and tumor mammary tissue through differential centrifugation. Furthermore, instructions are included for three-dimensional culturing as well as immunofluorescent imaging of embedded organoids.

Organoids are a reliable method for modeling organ tissue due to their self-organizing properties and retention of function and architecture after ...

Cancer Research

Culturing of Human Nasal Epithelial Cells at the Air Liquid Interface

In vitro models using human primary epithelial cells are essential in understanding key functions of the respiratory epithelium in the context of microbial infections or inhaled agents. Direct comparisons of cells obtained from diseased populations allow us to characterize different phenotypes and dissect the underlying mechanisms mediating changes in epithelial cell function. Culturing epithelial cells from the human tracheobronchial region has been well documented, but is limited by the availability of human lung tissue or invasiveness associated with obtaining the bronchial brushes biopsies. Nasal epithelial cells are obtained through much less invasive superficial nasal scrape biopsies and subjects can be biopsied multiple times with no significant side effects. Additionally, the nose is the entry point to the respiratory system and therefore one of the first sites to be exposed to any kind of air-borne stressor, such as microbial agents, pollutants, or allergens. 
Briefly, nasal epithelial cells obtained from human volunteers are expanded on coated tissue culture plates, and then transferred onto cell culture inserts. Upon reaching confluency, cells continue to be cultured at the air-liquid interface (ALI), for several weeks, which creates more physiologically relevant conditions. The ALI culture condition uses defined media leading to a differentiated epithelium that exhibits morphological and functional characteristics similar to the human nasal epithelium, with both ciliated and mucus producing cells. Tissue culture inserts with differentiated nasal epithelial cells can be manipulated in a variety of ways depending on the research questions (treatment with pharmacological agents, transduction with lentiviral vectors, exposure to gases, or infection with microbial agents) and analyzed for numerous different endpoints ranging from cellular and molecular pathways, functional changes, morphology, etc. 
In vitro models of differentiated human nasal epithelial cells will enable investigators to address novel and important research questions by using organotypic experimental models that largely mimic the nasal epithelium in vivo.

Nasal epithelial cells, obtained through superficial scrape biopsy of human volunteers, are expanded and transferred onto tissue culture inserts. Upon reaching confluency, cells are grown at air liquid interface, yielding cultures of ciliated and non-ciliated cells. Differentiated nasal epithelial cell cultures provide viable experimental models for studying the respiratory mucosa.

In vitro models using human primary  epithelial cells are essential in understanding key functions of the respiratory  epithelium in the context of ...

Biology

Infection of Primary Nasal Epithelial Cells Grown at an Air-Liquid Interface to Characterize Human Coronavirus-Host Interactions

Three highly pathogenic human coronaviruses (HCoVs) - SARS-CoV (2002), MERS-CoV (2012), and SARS-CoV-2 (2019) - have emerged and caused significant public health crises in the past 20 years. Four additional HCoVs cause a significant portion of common cold cases each year (HCoV-NL63, -229E, -OC43, and -HKU1), highlighting the importance of studying these viruses in physiologically relevant systems. HCoVs enter the respiratory tract and establish infection in the nasal epithelium, the primary site encountered by all respiratory pathogens. We use a primary nasal epithelial culture system in which patient-derived nasal samples are grown at an air-liquid interface (ALI) to study host-pathogen interactions at this important sentinel site. These cultures recapitulate many features of the in vivo airway, including&#160;the cell types present, ciliary function, and mucus production. We describe methods to characterize viral replication, host cell tropism, virus-induced cytotoxicity, and innate immune induction in nasal ALI cultures following HCoV infection, using recent work comparing lethal and seasonal HCoVs as an example1. An increased understanding of host-pathogen interactions in the nose has the potential to provide novel targets for antiviral therapeutics against HCoVs and other respiratory viruses that will likely emerge in the future.

The nasal epithelium is the primary barrier site encountered by all respiratory pathogens. Here, we outline methods to use primary nasal epithelial cells grown as air-liquid interface (ALI) cultures to characterize human coronavirus-host interactions in a physiologically relevant system.

Three highly pathogenic human coronaviruses (HCoVs) - SARS-CoV (2002), MERS-CoV (2012), and SARS-CoV-2 (2019) - have emerged and caused significant public ...

Immunology and Infection

Advanced Two-Layered 3D Co-Culture System for Epithelial and Fibroblast Cells

Establishing a Three-Dimensional Coculture Module of Epithelial Cells Using Nanofibrous Membranes

Technical hurdles in a culture of epithelial cells include dedifferentiation and loss of function. Biomimetic three-dimensional (3D) cell culture methods can enhance cell culture efficiency. This study introduces an advanced two-layered culture system intended to cultivate epithelial cells as tissue-like layers with the culture of fibroblasts within a 3D environment. Polyvinyl alcohol (PVA) and poly(&#949;-caprolactone) (PCL) nanofibrous membranes (NMs) were fabricated via electrospinning and utilized as a physiologically relevant extracellular matrix for the culture of epithelial cells and fibroblasts, respectively. In the upper insert wells, lung epithelial cells were cultivated on the PVA NM, and in the lower chambers, fibroblasts were cultured on the PCL NM. This configuration eliminates direct cell-cell contact and facilitates the examination of paracrine signaling mediated by soluble factors. Confocal microscopy was employed to analyze the distribution, growth pattern, and expression of intracellular proteins, including zona occludens in epithelial cells. Z-stacking techniques enabled detailed 3D reconstructions, providing precise insights into the integrity of tight junctions and spatial organization within the epithelial layer. Scanning electron microscopy (SEM) assessed the morphological characteristics of cell types on the nanofibrous membranes. SEM imaging revealed intricate cell surface structures and interactions with the nanofibers, offering a comprehensive perspective on cellular architecture and cell interaction with nanofibrous structure. The Cell Counting Kit-8 (CCK-8) assay is a simple method for measuring epithelial cell and fibroblast growth rates over time. It provides the proliferative behaviors and potential synergistic effects of coculturing these cells. These findings highlight the effectiveness of a simple insert co-culture system for simultaneous culture of fibroblasts and epithelial cells, which is crucial in various physiological and pharmacological contexts, including epithelial tissue regeneration, tumor microenvironment with endothelial, immune, and other stroma cells, toxicity assay, and drug activity test.

Author Spotlight: Advancing 3D Coculture Systems with PVA-PCL Nanofibrous Membranes 

Here, we demonstrate how epithelial cells cultured with fibroblasts in the nanofibrous membrane-based two-layer system can stably adhere to and grow on the membrane.

Technical hurdles in a culture of epithelial cells include dedifferentiation and loss of function. Biomimetic three-dimensional (3D) cell culture methods ...

Bioengineering

Collection, Expansion, and Differentiation of Primary Human Nasal Epithelial Cell Models for Quantification of Cilia Beat Frequency

Measurements of cilia function (beat frequency, pattern) have been established as diagnostic tools for respiratory diseases such as primary ciliary dyskinesia. However, the wider application of these techniques is limited by the extreme susceptibility of ciliary function to changes in environmental factors e.g., temperature, humidity, and pH. In the airway of patients with Cystic Fibrosis (CF), mucus accumulation impedes cilia beating. Cilia function has been investigated in primary airway cell models as an indicator of CF Transmembrane conductance Regulator (CFTR) channel activity. However, considerable patient-to-patient variability in cilia beating frequency has been found in response to CFTR-modulating drugs, even for patients with the same CFTR mutations. Furthermore, the impact of dysfunctional CFTR-regulated chloride secretion on ciliary function is poorly understood. There is currently no comprehensive protocol demonstrating sample preparation of in vitro airway models, image acquisition, and analysis of Cilia Beat Frequency (CBF). Standardized culture conditions and image acquisition performed in an environmentally controlled condition would enable consistent, reproducible quantification of CBF between individuals and in response to CFTR-modulating drugs. This protocol describes the quantification of CBF in three different airway epithelial cell model systems: 1) native epithelial sheets, 2) air-liquid interface models imaged on permeable support inserts, and 3) extracellular matrix-embedded three-dimensional organoids. The latter two replicate in vivo lung physiology, with beating cilia and production of mucus. The ciliary function is captured using a high-speed video camera in an environment-controlled chamber. Custom-built scripts are used for the analysis of CBF. Translation of CBF measurements to the clinic is envisioned to be an important clinical tool for predicting response to CFTR-modulating drugs on a per-patient basis.

This protocol describes nasal epithelial cell collection, expansion, and differentiation to organotypic airway epithelial cell models and quantification of cilia beat frequency via live-cell imaging and custom-built scripts.

Measurements of cilia function (beat frequency, pattern) have been established as diagnostic tools for respiratory diseases such as primary ciliary ...

Mucociliary Epithelial Organoids from Xenopus Embryonic Cells: Generation, Culture and High-Resolution Live Imaging

Mucociliary epithelium provides the first line of defense by removing foreign particles through the action of mucus production and cilia-mediated clearance. Many clinically relevant defects in the mucociliary epithelium are inferred as they occur deep within the body. Here, we introduce a tractable 3D model for mucociliary epithelium generated from multipotent progenitors that were microsurgically isolated from Xenopus laevis embryos. The mucociliary epithelial organoids are covered with newly generated epithelium from deep ectoderm cells and later decorated with distinct patterned multiciliated cells, secretory cells, and mucus-producing goblet cells that are indistinguishable from the native epidermis within 24 h. The full sequences of dynamic cell transitions from mesenchymal to epithelial that emerge on the apical surface of organoids can be tracked by high-resolution live imaging. These in vitro cultured, self-organizing mucociliary epithelial organoids offer distinct advantages in studying the biology of mucociliary epithelium with high-efficiency in generation, defined culture conditions, control over number and size, and direct access for live imaging during the regeneration of the differentiated epithelium.

We describe a simple protocol to develop mucociliary epithelial organoids from deep ectoderm cells isolated from Xenopus laevis embryos. The multipotent progenitors regenerate epithelial goblet cell precursors and allow live tracking of the initiation and progression of the cell transitions on the surface of organoids.

Mucociliary epithelium provides the first line of defense by removing foreign particles through the action of mucus production and cilia-mediated ...

Generation and Characterization of Murine Oral Mucosal Organoid Cultures

The mucous lining covering the inside of our mouth, the oral mucosa, is a highly compartmentalized tissue and can be subdivided into the buccal mucosa, gingiva, lips, palate, and tongue. Its uppermost layer, the oral epithelium, is maintained by adult stem cells throughout life. Proliferation and differentiation of adult epithelial stem cells have been intensively studied using in vivo mouse models as well as two-dimensional (2D) feeder-cell based in vitro models. Complementary to these methods is organoid technology, where adult stem cells are embedded into an extracellular matrix (ECM)-rich hydrogel and provided with a culture medium containing a defined cocktail of growth factors. Under these conditions, adult stem cells proliferate and spontaneously form three-dimensional (3D) cell clusters, the so-called organoids. Organoid cultures were initially established from murine small intestinal epithelial stem cells. However, the method has since been adapted for other epithelial stem cell types. Here, we describe a protocol for the generation and characterization of murine oral mucosal organoid cultures. Primary epithelial cells are isolated from murine tongue tissue, embedded into an ECM hydrogel, and cultured in a medium containing: epidermal growth factor (EGF), R-spondin, and fibroblast growth factor (FGF) 10. Within 7 to 14 days of initial seeding, the resulting organoids can be passaged for further expansion and cryopreservation. We additionally present strategies for the characterization of established organoid cultures via 3D whole-mount imaging and gene-expression analysis. This protocol may serve as a tool to investigate oral epithelial stem cell behavior ex vivo in a reductionist manner.

We present a method for the generation and characterization of oral mucosal organoid cultures derived from the tongue epithelium of adult mice.

The mucous lining covering the inside of our mouth, the oral mucosa, is a highly compartmentalized tissue and can be subdivided into the buccal mucosa, ...

Establishing Human Lung Organoids and Proximal Differentiation to Generate Mature Airway Organoids

The lack of a robust in vitro model of the human respiratory epithelium hinders the understanding of the biology and pathology of the respiratory system. We describe a defined protocol to derive human lung organoids from adult stem cells in the lung tissue and induce proximal differentiation to generate mature airway organoids. The lung organoids are then consecutively expanded for over 1 year with high stability, while the differentiated airway organoids are used to morphologically and functionally simulate human airway epithelium to a near-physiological level. Thus, we establish a robust organoid model of the human airway epithelium. The long-term expansion of lung organoids and differentiated airway organoids generates a stable and renewable source, enabling scientists to reconstruct and expand the human airway epithelial cells in culture dishes. The human lung organoid system provides a unique and physiologically active in vitro model for various applications, including studying virus-host interaction, drug testing, and disease modeling.

The protocol presents a method to derive human lung organoids from primary lung tissues, expand the lung organoids and induce proximal differentiation to generate 3D and 2D airway organoids that faithfully phenocopy the human airway epithelium.

The lack of a robust in vitro model of the human respiratory epithelium hinders the understanding of the biology and pathology of the respiratory system. ...

Culture and Imaging of Human Nasal Epithelial Organoids

Summary

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Chapters in this video

Cultivate Primary Nasal Epithelial Cells from Children and Reprogram into Induced Pluripotent Stem Cells

Three-Dimensional Imaging of Organoids to Study Primary Ciliogenesis During ex vivo Organogenesis

Generating and Imaging Mouse and Human Epithelial Organoids from Normal and Tumor Mammary Tissue Without Passaging

Culturing of Human Nasal Epithelial Cells at the Air Liquid Interface

Infection of Primary Nasal Epithelial Cells Grown at an Air-Liquid Interface to Characterize Human Coronavirus-Host Interactions

Establishing a Three-Dimensional Coculture Module of Epithelial Cells Using Nanofibrous Membranes

Collection, Expansion, and Differentiation of Primary Human Nasal Epithelial Cell Models for Quantification of Cilia Beat Frequency

Mucociliary Epithelial Organoids from Xenopus Embryonic Cells: Generation, Culture and High-Resolution Live Imaging

Generation and Characterization of Murine Oral Mucosal Organoid Cultures

Establishing Human Lung Organoids and Proximal Differentiation to Generate Mature Airway Organoids