This study has the following funding sources: National Institutes of Health (NIH) R01AI 169537 (S.R.W. and N.A.C.), NIH R01AI 140442 (S.R.W.), VA Merit Review CX001717 (N.A.C.), VA Merit Review BX005432 (S.R.W. and N.A.C.),&#160; Penn Center for Research on Coronaviruses and other Emerging Pathogens (S.R.W.), Laffey-McHugh Foundation (S.R.W. and N.A.C.), T32 AI055400 (CJO), T32 AI007324 (AF).

The use of nasal specimens was approved by the University of Pennsylvania Institutional Review Board (protocol # 800614) and the Philadelphia VA Institutional Review Board (protocol # 00781).
1. Infection of nasal ALI cultures
NOTE: Acquisition of clinical specimens, as well as growth and differentiation of nasal ALI cultures, is outside the scope of this paper. Specific methods for culturing primary nasal epithelial cells can be found in recently published works utilizing these cultures18,22,23. The below protocols can additionally be applied to commercially available nasal epithelial ALI cultures if desired. Protocols and volumes detailed below are applicable to 24-well plate transwell inserts (6.5&#160;mm diameter, 0.33 cm2 membrane surface area). If using ALI cultures grown on larger transwells (i.e., 12-well plates, 12 mm diameter, 1.12 cm2 surface area), adjust the volumes proportionally to reflect the transwell size.
<ol>
	<li>Day before infection:
	<ol>
		<li>Wash&#160;ALI cultures 3x with phosphate-buffered saline (PBS) apically (add ~200 &#956;L of warmed PBS, place in 37 &#176;C incubator for 5 min, aspirate PBS, and repeat).</li>
		<li>Replace basal medium (500 &#956;L).</li>
		<li>Allow cultures to equilibrate at the temperature at which infections will be conducted overnight (i.e., if infecting at 33 &#176;C, place cultures in a 33 &#176;C incubator after PBS washes). 
		NOTE: HCoVs associated with the common cold such as HCoV-229E and HCoV-NL63 are reported to replicate more efficiently at 33 &#176;C. Additionally, the temperature of the in vivo nasal epithelium is 33 &#176;C (this differs from the temperature of the lung, which is 37 &#176;C).</li>
	</ol>
	</li>
	<li>Dilute virus as needed in serum-free Dulbecco&#39;s Modified Eagle Medium (DMEM) to achieve the desired multiplicity of infection (MOI) in a total inoculum volume of 50 &#956;L. 
	NOTE: Infections have typically been conducted at MOI = 5 (high MOI); however, infections at MOI = 0.5 (low MOI) have also been used for SARS-CoV-2 and common cold-associated HCoVs, and these result in comparable peak viral titers, but with variable kinetics (either MOI is acceptable).</li>
	<li>Add inoculum apically, and place cultures back into the incubator for 1 h.</li>
	<li>Rock plates gently every 15 min during infection (holding plate firmly in both hands, rock forward and backward and side to side to ensure uniform adsorption of viral inoculum).</li>
	<li>After 1 h of incubation, aspirate viral inoculum, and wash each infected culture 3x with PBS to ensure removal of viral inoculum (for each wash, add 200 &#956;L of PBS, incubate for 5 min, and aspirate or remove with a pipette).</li>
	<li>If desired, collect the third PBS wash to confirm adequate removal of input virus.</li>
	<li>Replace the basal medium with fresh medium on infected ALIs every 72 h during infection.</li>
</ol>
2. Collection of apical surface liquid (ASL) and titration of shed virus
<ol>
	<li>At predetermined time points following infection, add 200 &#956;L of PBS to apical chamber of each infected transwell. 
	NOTE: Relevant time points vary depending on HCoV of interest and range from 24 h to 192 h post infection (see the representative results section for viral replication data for various HCoVs).</li>
	<li>Pipette PBS up and down 5x to ensure maximal collection of apically shed virus, and collect entire volume into a microcentrifuge tube (this is the ASL sample). 
	NOTE: ASL includes shed viral particles in addition to mucus and other products secreted apically from ALI cultures.</li>
	<li>Quantify infectious virus in ASL via&#160;standard viral plaque assay (serially dilute virus-containing samples in order to quantify the concentration of viral particles). 
	NOTE: Cell types and incubation period used for plaque assay will depend on the virus used: SARS-CoV-2 (VeroE6 cells); MERS-CoV (VeroCCL81 cells); HCoV-NL63 (LLC-MK2&#160;cells); HCoV-229E (Huh7 cells). Specifics on how to conduct viral plaque assays are beyond the scope of this manuscript but have been detailed previously in a Journal of Visualized Experiments (JoVE) publication24.</li>
	<li>If desired, collect the basal medium at various times post-infection to confirm the absence of basally released virus. HCoVs are typically released apically from nasal epithelial cells, but confirm this via plaque assay of undiluted basal medium.</li>
	<li>Store ASL samples at &#8722;80 &#176;C if quantification by&#160;plaque assay will not occur on the day of collection.</li>
</ol>
3. Quantification of intracellular virus
<ol>
	<li>After collecting ASL sample, move each transwell into a clean 24-well plate preloaded with 500 &#956;L of DMEM containing 2% fetal bovine serum (FBS) basally. 
	NOTE: DMEM with 2% FBS is used to stabilize the virus during subsequent freeze-thaw cycles.</li>
	<li>Wash each transwell 3x apically with PBS to ensure complete removal of apically shed virus.</li>
	<li>After aspirating to remove the final PBS wash, add 100 &#956;L of DMEM with 2% FBS to the apical compartment.</li>
	<li>Move the plate containing transwells with both apical and basal media into a &#8722;80 &#176;C freezer, and complete three consecutive freeze-thaw cycles to lyse the cells.</li>
	<li>After the final freeze-thaw cycle, pool apical (100 &#956;L) and basal (500 &#956;L) media into a clean tube.</li>
	<li>Centrifuge at 500 &#215; g for 10 min at 4 &#176;C to pellet any cellular debris.</li>
	<li>Collect the supernatant. This is the intracellular virus sample for titration via&#160;standard plaque assay. 
	NOTE: Three-fold dilution occurs during the collection process relative to ASL collection; ASL samples are collected in 200 &#956;L of PBS, while intracellular virus sample is collected into a total volume of 600 &#956;L.</li>
</ol>
4. Transepithelial electrical resistance (TEER) measurement
NOTE: For TEER measurement, PBS supplemented with calcium and magnesium (PBS + Ca2+/Mg2+) should be used. An epithelial volt/ohmmeter set to read in ohms is used (see the Table of Materials).
<ol>
	<li>Clean, equilibrate, and blank the EVOM instrument according to the manufacturer&#39;s instructions; use an &#34;empty&#34; transwell with no nasal cells added for blanking. Record blank TEER measurement. 
	NOTE: If using multiple viruses, the EVOM instrument must be cleaned stringently between conditions to avoid cross-contamination (washes with 70% ethanol followed by deionized water are sufficient).</li>
	<li>Move each infected transwell into a prelabeled, clean 24-well plate with 500 &#956;L of PBS + Ca2+/Mg2+ basally to wash residual basal medium from the transwells.</li>
	<li>Add 200 &#956;L of PBS + Ca2+/Mg2+ to the apical compartment of each transwell.</li>
	<li>Add 1 mL of PBS + Ca2+/Mg2+ to the Endohm-6 measurement chamber.</li>
	<li>Move each transwell into the Endohm-6 measurement chamber, and replace the lid of the chamber so that the apical electrode rests in the 200 &#956;L of PBS in the apical compartment; the basal electrode is built into the bottom of the Endohm-6 chamber.</li>
	<li>Allow the EVOM reading to stabilize, and record raw TEER measurement.</li>
	<li>Collect ASL sample as described above after taking TEER measurement if titering is desired (collect in the 200 &#956;L of PBS + Ca2+/Mg2+ that was added for TEER measurement). 
	NOTE: ASL sample collection can induce micro-tears in the epithelial barrier that may confound TEER readings, so ASL must be collected after TEER measurement. Further, basal medium must not be changed immediately before TEER readings as this may also impact TEER values.</li>
	<li>To convert raw TEER readings to final measurements in Ohms/cm2, subtract blank TEER value, and multiply this value by the surface area of the transwell membrane using equation (1): 
	TEER = [TEER reading - blank TEER value] &#215; (surface area of transwell)&#160; &#160; (1)</li>
	<li>For HCoVs, evaluate TEER every 24 h or 48 h following infection; the kinetics of TEER changes often vary among viruses and necessitate troubleshooting at various time points.</li>
	<li>When measuring TEER, always include&#160;mock-infected cultures&#160;and evaluate at each time point (negative control). 
	NOTE: For mock cultures, TEER measurements should remain stable or may slightly increase from baseline after differentiation of the cultures is completed. Surface area of membrane supports will vary depending on size and manufacturer of transwells; 24-well transwells typically have a surface area of 0.33 cm2.</li>
</ol>
5. Measurement of cytotoxicity during infection via&#160;lactate dehydrogenase (LDH) assay
NOTE: In this work, LDH content in ASL samples was quantified using a commercially available cytotoxicity detection kit. LDH signal in basal medium was often below the limit of detection and often less reproducible than LDH quantified in ASL samples from HCoV-infected cultures.
<ol>
	<li>Prepare additional controls necessary for LDH assay.
	<ol>
		<li>Background control:&#160;use PBS (use PBS containing calcium and magnesium if ASL samples were collected in this way).</li>
		<li>Positive control (ceiling value):&#160;treat ALIs apically with Triton X-100. Collect Triton control wells (typically use three ALI cultures for this at each time point).
		<ol>
			<li>Add 200 &#956;L of 2% Triton X-100 in PBS directly to the apical compartment of the transwell.</li>
			<li>Incubate for 10-15 min to allow cells to completely lyse.</li>
			<li>Collect the entire volume as a Triton ceiling sample.</li>
		</ol>
		</li>
		<li>Negative control (low control/baseline LDH release):&#160;collect ASL from mock-infected ALI cultures derived from the same donor as infected cultures.
		<ol>
			<li>Collect negative control mock ASL following ASL collection procedure above. 
			NOTE: The same transwells can be used for this negative control for all time points, whereas fresh transwells will be necessary for each time point for the Triton control.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>To allow for quantification of shed virus in addition to&#160;LDH readings from each ASL sample, load an optically clear flat-bottom black 96-well plate as follows:
	<ol>
		<li>Dilute all samples in PBS, using 45 &#956;L of sample and 55 &#956;L of PBS (100 &#956;L total volume).</li>
		<li>Treat Triton positive control and mock background control samples in the same manner. 
		NOTE: This dilution allows for triplicate loading of each experimental sample (45 &#956;L x 3) and enough residual ASL volume for titering via&#160;plaque assay.</li>
		<li>Load&#160;LDH plate in triplicate: Triton ceiling, mock background control, PBS control, and experimental samples.</li>
		<li>Prepare the reaction mixture as indicated by the manufacturer (dye solution + catalyst).
		<ol>
			<li>Add 100 &#956;L of reaction mixture to each well, and incubate the plate at room temperature for 20 min protected from light.</li>
		</ol>
		</li>
		<li>After 20 min, measure absorbance at 492 nm.</li>
		<li>Calculate percentage cytotoxicity relative to the Triton ceiling value using equation (2). 
		Cytotoxicity (%) &#160;<img alt="Equation 1" src="/files/ftp_upload/64868/64868eq01.jpg" style="margin: auto;" />&#160; &#160; (2)</li>
	</ol>
	</li>
</ol>
6. Preparation of nasal ALI cultures for immunofluorescence (IF) imaging
<ol>
	<li>Move the transwell into a fresh 24-well plate, and wash 3x apically with PBS (collect the first PBS wash as ASL if titering; then, perform two additional PBS washes to remove excess shed virus that might hinder IF quality)</li>
	<li>After the final PBS wash, cover the transwell apically and basally in 4% PFA.</li>
	<li>Incubate for 30 min to fix in 4% PFA, and then remove and wash 3x with PBS.</li>
	<li>Excise the transwell support containing the cells using a sharpened razor blade or scissors.</li>
	<li>To increase the number of IF targets for each transwell, cut each membrane in half, and stain each half with different antibody combinations.</li>
	<li>Permeabilize with 0.2% Triton X-100 in PBS for 10 min.</li>
	<li>Block with 10% normal donkey serum and 1% BSA in PBST (PBS + 0.2% Triton X-100) for 60 min at room temperature. 
	NOTE: Protect samples from light exposure from this step forward.</li>
	<li>Incubate in primary antibody solution overnight at 4 &#176;C. Dilute all antibodies 1:1,000 in blocking buffer. 
	NOTE: Representative antibodies for HCoV nucleocapsid staining, as well as epithelial cell type markers and cytoskeletal stains, are listed below (see the Table of Materials for manufacturer information and catalog numbers).
	<ol>
		<li>For HCoV antigen staining, use antibodies targeting SARS-CoV-2 nucleocapsid, MERS-CoV nucleocapsid, and HCoV-NL63 nucleocapsid.</li>
		<li>To identify epithelial cell types, use antibodies targeting goblet cell marker MUC5AC&#160;and ciliated cell marker type IV &#946;-tubulin.</li>
		<li>For cytoskeletal markers, use antibodies directed against phalloidin (binds F-actin) and epithelial cell adhesion marker: EpCAM (CD326).</li>
	</ol>
	</li>
	<li>Incubate in secondary antibody solution for 60 min at room temperature; use secondary antibody dyes diluted 1:1,000 in blocking buffer.</li>
	<li>After&#160;staining is complete, transfer the membrane onto a glass slide with a spatula, orient the transwell with the apical side toward the slide, and add mounting solution. Allow to settle for 15-30 min prior to applying clear nail polish around edges.</li>
	<li>Acquire images using a confocal microscope (Z-axis step: 0.5 &#956;m; sequential scanning)1,16,17. 
	&#8203;NOTE: After fixation of cultures in 4% PFA and washing with PBS, fixed cultures can be stored at 4 &#176;C for weeks to months prior to staining and preparation for imaging. After staining and mounting of membranes, samples can be stored long-term (&#62;2 years) at 4 &#176;C in the dark.</li>
</ol>
7. Collection of intracellular protein for western immunoblotting or RNA for RT-qPCR analysis
<ol>
	<li>Collect ASL as described above if quantifying viral titers (protocol section 2).</li>
	<li>Move the transwell to a clean 24-well plate, as scraping the membrane can lead to breaking the insert.</li>
	<li>For western blot analysis, collect total protein lysate in 125 &#956;L of RIPA buffer (50 mM Tris, pH 8, 150 mM NaCl, 0.5% deoxycholate, 0.1% SDS, 1% NP40) supplemented with protease inhibitors and phosphatase inhibitors.</li>
	<li>Add 125 &#956;L of RIPA buffer to the apical compartment, and incubate for 5-10 min.</li>
	<li>Scrape the membrane using a P200 pipette tip to remove any remaining attached cells, and collect the entire volume (scrape vigorously across the entire surface of the membrane, and then pipette up and down multiple times to collect the entire sample).</li>
	<li>Incubate protein lysate samples on ice for 10 min, and then centrifuge at maximum speed (20,000 &#215; g) for 10 min at 4 &#176;C.</li>
	<li>Mix the supernatant with 4x Laemmli sample buffer with &#946;-mercaptoethanol (reducing agent) according to manufacturer protocols.</li>
	<li>Boil protein samples at 95 &#176;C for 5 min, and then run using traditional western blotting protocols1,16,17.</li>
	<li>For collection of total RNA, use a commercially available RNA extraction kit of choice. 
	NOTE: The apical compartment of 24-well transwell inserts has a maximal volume of ~200 &#956;L; thus, we perform two sequential washes on each culture to reach total recommended volume. Details below correspond to recommended collection of RNA samples in a 350 &#956;L total volume.</li>
	<li>Add 200 &#956;L of lysis buffer to the apical compartment of the infected transwell.</li>
	<li>Let sit for 5-10 min, scrape any remaining cells from the membrane using a pipette tip, and collect the entire volume into a labeled microcentrifuge tube.</li>
	<li>Add 150 &#956;L of additional lysis buffer to the apical compartment of the transwell, and pipette up and down before collecting into same microcentrifuge tube.</li>
	<li>Extract RNA according to the manufacturer&#39;s protocol.</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=Airway+epithelial+differentiation+and+mucociliary+clearance.">Airway epithelial differentiation and mucociliary clearance.</a> Annals of the American Thoracic Society. 15, S143-S148 (2018).</li><li>Gao, N., Raduka, A., Rezaee, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Respiratory+syncytial+virus+disrupts+the+airway+epithelial+barrier+by+decreasing+cortactin+and+destabilizing+F-actin.">Respiratory syncytial virus disrupts the airway epithelial barrier by decreasing cortactin and destabilizing F-actin.</a> Journal of Cell Science. 135 (16), 259871 (2022).</li><li>Schmidt, H., 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=IL-13+impairs+tight+junctions+in+airway+epithelia.">IL-13 impairs tight junctions in airway epithelia.</a> International Journal of Molecular Sciences. 20 (13), 3222 (2019).</li><li>Huang, Z. Q., 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=Interleukin-13+alters+tight+junction+proteins+expression+thereby+compromising+barrier+function+and+dampens+rhinovirus+induced+immune+responses+in+nasal+epithelium.">Interleukin-13 alters tight junction proteins expression thereby compromising barrier function and dampens rhinovirus induced immune responses in nasal epithelium.</a> Frontiers in Cell and Developmental Biology. 8, 572749 (2020).</li><li>Saatian, B., 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=Interleukin-4+and+interleukin-13+cause+barrier+dysfunction+in+human+airway+epithelial+cells.">Interleukin-4 and interleukin-13 cause barrier dysfunction in human airway epithelial cells.</a> Tissue Barriers. 1 (2), e24333 (2013).</li><li>Coles, J. 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=A+revised+protocol+for+culture+of+airway+epithelial+cells+as+a+diagnostic+tool+for+primary+ciliary+dyskinesia.">A revised protocol for culture of airway epithelial cells as a diagnostic tool for primary ciliary dyskinesia.</a> Journal of Clinical Medicine. 9 (11), 3753 (2020).</li><li>Baldassi, D., Gabold, B., Merkel, O. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Air&#8722;liquid+interface+cultures+of+the+healthy+and+diseased+human+respiratory+tract:+Promises,+challenges,+and+future+directions.">Air&#8722;liquid interface cultures of the healthy and diseased human respiratory tract: Promises, challenges, and future directions.</a> Advanced NanoBiomed Research. 1 (6), 2000111 (2021).</li><li>Seibold, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interleukin-13+stimulation+reveals+the+cellular+and+functional+plasticity+of+the+airway+epithelium.">Interleukin-13 stimulation reveals the cellular and functional plasticity of the airway epithelium.</a> Annals of the American Thoracic Society. 15, S98-S106 (2018).</li><li>Morrison, C. B., 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=SARS-CoV-2+infection+of+airway+cells+causes+intense+viral+and+cell+shedding,+two+spreading+mechanisms+affected+by+IL-13.">SARS-CoV-2 infection of airway cells causes intense viral and cell shedding, two spreading mechanisms affected by IL-13.</a> Proceedings of the National Academy of Sciences of the United States of America. 119 (16), 2119680119 (2022).</li></ol>

Susan Weiss is on the Scientific Advisory Boards for Ocugen. Noam A. Cohen consults for GSK, AstraZeneca, Novartis, Sanofi/Regeron, and Oyster Point Pharmaceuticals and has a US Patent, &#34;Therapy and Diagnostics for Respiratory Infection&#34; (10,881,698 B2, WO20913112865), and a licensing agreement with GeneOne Life Sciences.

The methods detailed here describe a primary epithelial culture system in which patient-derived nasal epithelial cells are grown at an air-liquid interface and applied to the study of HCoV-host interactions. Once differentiated, these nasal ALI cultures recapitulate many features of the in vivo nasal epithelium, including a heterogeneous cellular population with ciliated, goblet, and basal cells represented, as well as intact mucociliary function with robustly beating cilia and mucus secretion. This heterogeneou...

Seven human coronaviruses (HCoVs) have been identified to date and cause a range of respiratory diseases2. The common or seasonal HCoVs (HCoV-NL63, -229E, -OC43, and -HKU1) are typically associated with upper respiratory tract pathology and cause an estimated 10%-30% of common cold cases annually. Though this is the typical clinical phenotype associated with the common HCoVs, these viruses can cause more significant lower respiratory tract disease in at-risk populations, including children, older adults, and immunocompromised individuals3,4. Three pathogenic HCoVs have emerged and caused significant public health emergencies in the last 20 years, including severe acute respiratory syndrome (SARS)-CoV, Middle East respiratory syndrome (MERS)-CoV, and SARS-CoV-2. Lethal HCoVs are associated with more severe respiratory tract pathology, which is clearly illustrated by the &#62;34% case-fatality rate associated with MERS-CoV cases (894 deaths from over 2,500 cases since its emergence in 2012)5,6. It is important to note that the lethal HCoVs also cause a range of respiratory tract diseases, from asymptomatic infections to lethal pneumonia, as seen with the ongoing COVID-19 pandemic7.
HCoVs, like other respiratory pathogens, enter the respiratory tract and establish a productive infection in the nasal epithelium8. Spread to the lower airway is thought to be associated with aspiration from the oral/nasal cavity to the lung, where HCoVs cause more significant lower respiratory tract pathology9,10,11. Thus, the nose serves as the initial portal for viral entry and is the primary barrier to infection with its robust mucociliary clearance machinery and unique innate immune mechanisms aimed at preventing further viral spread to the lower airway12,13. For example, nasal epithelial cells have been reported to express higher than average basal levels of antiviral interferons and interferon-stimulated genes, indicating that nasal cells may be primed for early responses to respiratory viruses14,15,16.
We have previously utilized patient-derived primary nasal epithelial cells grown at an air-liquid interface (ALI) to model HCoV-host interactions in the nose, where HCoV infections begin. Nasal ALI cultures are permissive to both pathogenic (SARS-CoV-2 and MERS-CoV) and common HCoVs (HCoV-NL63 and HCoV-229E) and offer various advantages over traditional airway epithelial cell lines such as A549 (a lung adenocarcinoma cell line)16,17. After differentiation, nasal ALI cultures contain a heterogeneous cellular population and exhibit many of the functions expected of the in vivo nasal epithelium, such as mucociliary clearance machinery18. Nasal cells also offer advantages over lower airway culture systems (such as human bronchial epithelial cells, HBECs), as the acquisition of nasal epithelial cells via cytologic brushing is significantly less invasive compared with using techniques such as bronchoscopy for attaining HBECs19,20,21.
This paper describes methods for utilizing this nasal ALI culture system to characterize HCoV-host interactions in the nasal epithelium. We have applied these methods in recently published works to compare SARS-CoV-2, MERS-CoV, HCoV-NL63, and HCoV-229E1,16,17. Though these methods and representative results emphasize the study of HCoVs in this nasal cell model, the system is highly adaptable to other HCoVs, as well as other respiratory pathogens. Further, these methods can be applied more broadly to other ALI culture systems in order to investigate viral replication and cellular tropism, as well as cytotoxicity and innate immune induction following infection.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Alexa Fluor secondary antibodies (488, 594, 647)</td>
			<td>Invitrogen</td>
			<td>Various</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA (bovine serum albumin)</td>
			<td>Sigma-Aldrich</td>
			<td>A7906</td>
			<td></td>
		</tr>
		<tr>
			<td>cOmplete mini EDTA-free protease inhibitor</td>
			<td>Roche</td>
			<td>11836170001</td>
			<td></td>
		</tr>
		<tr>
			<td>Cytotoxicity detection kit</td>
			<td>Roche</td>
			<td>11644793001</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM (Dulbecco&#39;s Modified Eagle Media)</td>
			<td>Gibco</td>
			<td>11965-084</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS (Dulbecco&#39;s Phosphate Buffered Saline)</td>
			<td>Gibco</td>
			<td>14190136</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS + calcium + magnesium</td>
			<td>Gibco</td>
			<td>14040-117</td>
			<td></td>
		</tr>
		<tr>
			<td>Endohm-6G measurement chamber</td>
			<td>World Precision Instruments</td>
			<td>ENDOHM-6G</td>
			<td></td>
		</tr>
		<tr>
			<td>Epithelial cell adhesion marker (EpCAM; CD326)</td>
			<td>eBiosciences</td>
			<td>14-9326-82</td>
			<td></td>
		</tr>
		<tr>
			<td>Epithelial Volt/Ohm (TEER) Meter (EVOM)</td>
			<td>World Precision Instruments</td>
			<td>300523</td>
			<td></td>
		</tr>
		<tr>
			<td>FBS (Fetal Bovine Serum)</td>
			<td>HyClone</td>
			<td>SH30071.03</td>
			<td></td>
		</tr>
		<tr>
			<td>FV10-ASW software for imaging</td>
			<td>Olympus</td>
			<td>Version 4.02</td>
			<td></td>
		</tr>
		<tr>
			<td>HCoV-NL63 (Human coronavirus, NL63)</td>
			<td>BEI Resources</td>
			<td>NR-470</td>
			<td></td>
		</tr>
		<tr>
			<td>HCoV-NL63 nucleocapsid antibody</td>
			<td>Sino Biological</td>
			<td>40641-V07E</td>
			<td></td>
		</tr>
		<tr>
			<td>Hoescht stain</td>
			<td>Thermo Fisher</td>
			<td>H3570</td>
			<td></td>
		</tr>
		<tr>
			<td>Laemmli sample buffer (4x)</td>
			<td>BIO-RAD</td>
			<td>1610747</td>
			<td></td>
		</tr>
		<tr>
			<td>LLC-MK2 cells</td>
			<td>ATCC</td>
			<td>CCL-7</td>
			<td>To titrate HCoV-NL63</td>
		</tr>
		<tr>
			<td>MERS-CoV (Human coronavirus, Middle East Respiratory Syndrome Coronavirus (MERS-CoV), EMC/2012)</td>
			<td>BEI Resources&#160;</td>
			<td>NR-44260</td>
			<td></td>
		</tr>
		<tr>
			<td>MERS-CoV nucleocapsid antibody</td>
			<td>Sino Biological</td>
			<td>40068-MM10</td>
			<td></td>
		</tr>
		<tr>
			<td>MUC5AC antibody</td>
			<td>Sigma-Aldrich</td>
			<td>AMAB91539</td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus Fluoview confocal microscope</td>
			<td>Olympus</td>
			<td>FV1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Phalloidin-iFluor 647 stain</td>
			<td>Abcam</td>
			<td>ab176759</td>
			<td></td>
		</tr>
		<tr>
			<td>PhosStop easy pack (phosphatase inhibitors)&#160;</td>
			<td>Roche</td>
			<td>PHOSS-RO</td>
			<td></td>
		</tr>
		<tr>
			<td>Plate reader&#160;</td>
			<td>Perkin Elmer</td>
			<td>HH34000000</td>
			<td>Any plate reader or ELISA reader is sufficient; must be able to read absorbance at 492 nm</td>
		</tr>
		<tr>
			<td>RIPA buffer (50 mM Tris pH 8; 150 mM NaCl; 0.5% deoxycholate; 0.1% SDS; 1% NP40)</td>
			<td>Thermo Fisher</td>
			<td>89990</td>
			<td>Can prep in-house or purchase</td>
		</tr>
		<tr>
			<td>RNeasy Plus Kit</td>
			<td>Qiagen</td>
			<td>74134</td>
			<td></td>
		</tr>
		<tr>
			<td>SARS-CoV-2 (SARS-Related Coronavirus 2, Isolate USA-WA1/2020)</td>
			<td>BEI Resources</td>
			<td>NR-52281</td>
			<td></td>
		</tr>
		<tr>
			<td>SARS-CoV-2 nucleocapsid antibody</td>
			<td>Genetex</td>
			<td>GTX135357</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton-X 100</td>
			<td>Fisher Scientific</td>
			<td>BP151100</td>
			<td></td>
		</tr>
		<tr>
			<td>Type IV &#946;- tubulin antibody</td>
			<td>Abcam</td>
			<td>ab11315</td>
			<td></td>
		</tr>
		<tr>
			<td>VeroCCL81 cells</td>
			<td>ATCC</td>
			<td>CCL-81</td>
			<td>To titrate MERS-CoV</td>
		</tr>
		<tr>
			<td>VeroE6 cells</td>
			<td>ATCC</td>
			<td>CRL-1586</td>
			<td>To titrate SARS-CoV-2</td>
		</tr>
	</tbody>
</table>

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 representative figures are partially adapted from data that can be found in the manuscript Otter et al.1. Nasal ALI cultures derived from four or six donors were infected with one of four HCoVs (SARS-CoV-2, MERS-CoV, HCoV-NL63, and HCoV-229E) according to the protocols described above, and the average apically shed viral titers for each virus are depicted in Figure 1A. While all four of these HCoVs replicate productively in nasal ALI cultures, SARS-CoV-2 and HCoV-...

Watch this Scientific Journal Video about Infection of Primary Nasal Epithelial Cells Grown at an Air-Liquid Interface to Characterize Human Coronavirus-Host Interactions at JoVE.com

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

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 ...

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1.1K Views.  University of Pennsylvania. 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.

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

Application of a Mouse Ligated Peyer&#x2019;s Patch Intestinal Loop Assay to Evaluate Bacterial Uptake by M cells

The inside of our gut is inhabited with enormous number of commensal bacteria. The mucosal surface of the gastrointestinal tract is continuously exposed to them and occasionally to pathogens. The gut-associated lymphoid tissue (GALT) play a key role for induction of the mucosal immune response to these microbes1, 2. To initiate the mucosal immune response, the mucosal antigens must be transported from the gut lumen across the epithelial barrier into organized lymphoid follicles such as Peyer's patches. This antigen transcytosis is mediated by specialized epithelial M cells3, 4. M cells are atypical epithelial cells that actively phagocytose macromolecules and microbes. Unlike dendritic cells (DCs) and macrophages, which target antigens to lysosomes for degradation, M cells mainly transcytose the internalized antigens. This vigorous macromolecular transcytosis through M cells delivers antigen to the underlying organized lymphoid follicles and is believed to be essential for initiating antigen-specific mucosal immune responses. However, the molecular mechanisms promoting this antigen uptake by M cells are largely unknown. We have previously reported that glycoprotein 2 (Gp2), specifically expressed on the apical plasma membrane of M cells among enterocytes, serves as a transcytotic receptor for a subset of commensal and pathogenic enterobacteria, including Escherichia coli and Salmonella enterica serovar Typhimurium (S. Typhimurium), by recognizing FimH, a component of type I pili on the bacterial outer membrane 5. Here, we present a method for the application of a mouse Peyer's patch intestinal loop assay to evaluate bacterial uptake by M cells. This method is an improved version of the mouse intestinal loop assay previously described 6, 7. The improved points are as follows: 1. Isoflurane was used as an anesthetic agent. 2. Approximately 1 cm ligated intestinal loop including Peyer's patch was set up. 3. Bacteria taken up by M cells were fluorescently labeled by fluorescence labeling reagent or by overexpressing fluorescent protein such as green fluorescent protein (GFP). 4. M cells in the follicle-associated epithelium covering Peyer's patch were detected by whole-mount immunostainig with anti Gp2 antibody. 5. Fluorescent bacterial transcytosis by M cells were observed by confocal microscopic analysis. The mouse Peyer's patch intestinal loop assay could supply the answer what kind of commensal or pathogenic bacteria transcytosed by M cells, and may lead us to understand the molecular mechanism of how to stimulate mucosal immune system through M cells.

Application of a Mouse Ligated Peyer&amp;#8217;s Patch Intestinal Loop Assay to Evaluate Bacterial Uptake by M cells

M cells in a specialized follicle-associated epithelium covering Peyer&#x2019;s patches play an important role for the mucosal immunosurveillance in gut-associated lymphoid tissue. Here we described the evaluation method for bacterial transcytosis by M cells in vivo. This method provides a method to understand M-cell function in the immune system.

The inside of our gut is inhabited with enormous number of commensal bacteria. The mucosal surface of the gastrointestinal tract is continuously exposed ...

An In vitro Co-infection Model to Study Plasmodium falciparum-HIV-1 Interactions in Human Primary Monocyte-derived Immune Cells

Plasmodium falciparum, the causative agent of the deadliest form of malaria, and human immunodeficiency virus type-1 (HIV-1) are among the most important health problems worldwide, being responsible for a total of 4 million deaths annually1. Due to their extensive overlap in developing regions, especially Sub-Saharan Africa, co-infections with malaria and HIV-1 are common, but the interplay between the two diseases is poorly understood. Epidemiological reports have suggested that malarial infection transiently enhances HIV-1 replication and increases HIV-1 viral load in co-infected individuals2,3. Because this viremia stays high for several weeks after treatment with antimalarials, this phenomenon could have an impact on disease progression and transmission.
The cellular immunological mechanisms behind these observations have been studied only scarcely. The few in vitro studies investigating the impact of malaria on HIV-1 have demonstrated that exposure to soluble malarial antigens can increase HIV-1 infection and reactivation in immune cells. However, these studies used whole cell extracts of P. falciparum schizont stage parasites and peripheral blood mononuclear cells (PBMC), making it hard to decipher which malarial component(s) was responsible for the observed effects and what the target host cells were4,5. Recent work has demonstrated that exposure of immature monocyte-derived dendritic cells to the malarial pigment hemozoin increased their ability to transfer HIV-1 to CD4+ T cells6,7, but that it decreased HIV-1 infection of macrophages8. To shed light on this complex process, a systematic analysis of the interactions between the malaria parasite and HIV-1 in different relevant human primary cell populations is critically needed.
Several techniques for investigating the impact of HIV-1 on the phagocytosis of micro-organisms and the effect of such pathogens on HIV-1 replication have been described. We here present a method to investigate the effects of P. falciparum-infected erythrocytes on the replication of HIV-1 in human primary monocyte-derived macrophages. The impact of parasite exposure on HIV-1 transcriptional/translational events is monitored by using single cycle pseudotyped viruses in which a luciferase reporter gene has replaced the Env gene while the effect on the quantity of virus released by the infected macrophages is determined by measuring the HIV-1 capsid protein p24 by ELISA in cell supernatants.

An In vitro Co-infection Model to Study Plasmodium falciparum-HIV-1 Interactions in Human Primary Monocyte-derived Immune Cells

We have developed an in vitro malaria-HIV-1 co-infection model to study the impact of Plasmodium falciparum on the HIV-1 replicative cycle in human primary monocyte-derived macrophages. This versatile system can easily be adapted to other primary cell types susceptible to HIV-1 infection.

Plasmodium falciparum, the causative agent  of the deadliest form of malaria, and human immunodeficiency virus type-1 (HIV-1) are among the most important ...

Flow Cytometric Isolation of Primary Murine Type II Alveolar Epithelial Cells for Functional and Molecular Studies

Throughout the last years, the contribution of alveolar type II epithelial cells (AECII) to various aspects of immune regulation in the lung has been increasingly recognized. AECII have been shown to participate in cytokine production in inflamed airways and to even act as antigen-presenting cells in both infection and T-cell mediated autoimmunity 1-8. Therefore, they are especially interesting also in clinical contexts such as airway hyper-reactivity to foreign and self-antigens as well as infections that directly or indirectly target AECII. However, our understanding of the detailed immunologic functions served by alveolar type II epithelial cells in the healthy lung as well as in inflammation remains fragmentary. Many studies regarding AECII function are performed using mouse or human alveolar epithelial cell lines 9-12. Working with cell lines certainly offers a range of benefits, such as the availability of large numbers of cells for extensive analyses. However, we believe the use of primary murine AECII allows a better understanding of the role of this cell type in complex processes like infection or autoimmune inflammation. Primary murine AECII can be isolated directly from animals suffering from such respiratory conditions, meaning they have been subject to all additional extrinsic factors playing a role in the analyzed setting. As an example, viable AECII can be isolated from mice intranasally infected with influenza A virus, which primarily targets these cells for replication 13. Importantly, through ex vivo infection of AECII isolated from healthy mice, studies of the cellular responses mounted upon infection can be further extended.
Our protocol for the isolation of primary murine AECII is based on enzymatic digestion of the mouse lung followed by labeling of the resulting cell suspension with antibodies specific for CD11c, CD11b, F4/80, CD19, CD45 and CD16/CD32. Granular AECII are then identified as the unlabeled and sideward scatter high (SSChigh) cell population and are separated by fluorescence activated cell sorting 3.
In comparison to alternative methods of isolating primary epithelial cells from mouse lungs, our protocol for flow cytometric isolation of AECII by negative selection yields untouched, highly viable and pure AECII in relatively short time. Additionally, and in contrast to conventional methods of isolation by panning and depletion of lymphocytes via binding of antibody-coupled magnetic beads 14, 15, flow cytometric cell-sorting allows discrimination by means of cell size and granularity. Given that instrumentation for flow cytometric cell sorting is available, the described procedure can be applied at relatively low costs. Next to standard antibodies and enzymes for lung disintegration, no additional reagents such as magnetic beads are required. The isolated cells are suitable for a wide range of functional and molecular studies, which include in vitro culture and T-cell stimulation assays as well as transcriptome, proteome or secretome analyses 3, 4.

We describe the rapid isolation of primary murine type II alveolar epithelial cells (AECII) by flow cytometric negative selection. These AECII show high viability and purity and are suitable for a wide range of functional and molecular studies regarding their role in respiratory conditions such as autoimmune or infectious diseases.

Throughout  the last years, the contribution of alveolar type II epithelial cells (AECII)  to various aspects of immune regulation in the lung has been ...

Isolation of Human Monocytes by Double Gradient Centrifugation and Their Differentiation to Macrophages in Teflon-coated Cell Culture Bags

Human macrophages are involved in a plethora of pathologic processes ranging from infectious diseases to cancer. Thus they pose a valuable tool to understand the underlying mechanisms of these diseases. We therefore present a straightforward protocol for the isolation of human monocytes from buffy coats, followed by a differentiation procedure which results in high macrophage yields. The technique relies mostly on commonly available lab equipment and thus provides a cost and time effective way to obtain large quantities of human macrophages. Briefly, buffy coats from healthy blood donors are subjected to a double density gradient centrifugation to harvest monocytes from the peripheral blood. These monocytes are then cultured in fluorinated ethylene propylene (FEP) Teflon-coated cell culture bags in the presence of macrophage colony-stimulating factor (M-CSF). The differentiated macrophages can be easily harvested and used for subsequent studies and functional assays. Important methods for quality control and validation of the isolation and differentiation steps will be highlighted within the protocol. In summary, the protocol described here enables scientists to routinely and reproducibly isolate human macrophages without the need for cost intensive tools. Furthermore, disease models can be studied in a syngeneic human system circumventing the use of murine macrophages.

We present a simple and efficient protocol for the generation of human macrophages. Buffy coats are processed by double density gradient centrifugation and isolated monocytes are then differentiated to macrophages in Teflon-coated cell culture bags. This maximizes macrophage yields and facilitates cell harvesting for subsequent experiments.

Human macrophages are involved in a plethora of pathologic processes ranging from infectious diseases to cancer. Thus they pose a valuable tool to ...

Biomimetic Materials to Characterize Bacteria-host Interactions

Bacterial attachment to host cells is one of the earliest events during bacterial colonization of host tissues and thus a key step during infection. The biochemical and functional characterization of adhesins mediating these initial bacteria-host interactions is often compromised by the presence of other bacterial factors, such as cell wall components or secreted molecules, which interfere with the analysis. This protocol describes the production and use of biomimetic materials, consisting of pure recombinant adhesins chemically coupled to commercially available, functionalized polystyrene beads, which have been used successfully to dissect the biochemical and functional interactions between individual bacterial adhesins and host cell receptors. Protocols for different coupling chemistries, allowing directional immobilization of recombinant adhesins on polymer scaffolds, and for assessment of the coupling efficiency of the resulting &#8220;bacteriomimetic&#8221; materials are also discussed. We further describe how these materials can be used as a tool to inhibit pathogen mediated cytotoxicity and discuss scope, limitations and further applications of this approach in studying bacterial - host interactions.

Bacterial attachment to host cells is a key step during host colonization and infection. This protocol describes the generation of polymer-coupled recombinant adhesins as biomimetic materials which allow analysis of the contribution of individual adhesins to these processes, independent of other bacterial factors.

Bacterial attachment to host cells is one of the earliest events during bacterial colonization of host tissues and thus a key step during infection. The ...

Human Placental and Decidual Organ Cultures to Study Infections at the Maternal-fetal Interface

The placenta shows a large degree of interspecies anatomic variability. To best understand biology and pathophysiology of the human placenta, it is imperative to design experiments using human cells and tissues. An advantage of organ culture is maintenance of three-dimensional (3D) structural organization and extracellular matrix. The goal of the method described here is successful establishment of ex vivo human gestational tissue organ cultures and their healthy culture maintenance for 72-96 hr. The protocol details the immediate processing of research-consented, placental and decidual specimens fresh from the operating suite. These are abundant specimens that would otherwise be discarded. Detailed instructions on the sterile collection of these samples, including morphologic details on how to select appropriate tissues to establish 3D organ cultures, is provided. Placental villous and decidual tissues are microdissected into 2-3 mm3 pieces and placed separately on matrix-lined transwell filters and cultured for several days. Villous and decidual organ cultures are well suited for the study of human host-pathogen interaction. As compared to other model organisms, these human cultures are particularly advantageous to examine mechanism of infection for pathogens that demonstrate variable patterns of host specificity. As an example, we demonstrate infection of placental and decidual organ cultures with the clinically relevant, facultative intracellular bacterial pathogen Listeria monocytogenes.

A simple method to establish primary human placental (villous) and decidual organ cultures is described. Villous and decidual organ cultures are invaluable tools for studying pathogenesis at the human maternal-fetal interface. Infection with the facultative intracellular bacterium Listeria monocytogenes is demonstrated.

The placenta shows a large degree of interspecies anatomic variability. To best understand biology and pathophysiology of the human placenta, it is ...

Assessment of the Synaptic Interface of Primary Human T Cells from Peripheral Blood and Lymphoid Tissue

The current understanding of the dynamics and structural features of T-cell synaptic interfaces has been largely determined through the use of glass-supported planar bilayers and in vitro-derived T-cell clones or lines1,2,3,4. How these findings apply to the primary human T cells isolated from blood or lymphoid tissues is not known, partly due to significant difficulties in obtaining a sufficient number of cells for analysis5. Here we address this through the development of a technique exploiting multichannel flow slides to build planar lipid bilayers containing activating and adhesion molecules. The low height of the flow slides promotes rapid cell sedimentation in order to synchronize cell:bilayer attachment, thereby allowing researchers to study the dynamic of the synaptic interface formation and the kinetics of the granules release. We apply this approach to analyze the synaptic interface of as few as 104 to 105 primary cryopreserved T cells isolated from lymph nodes (LN) and peripheral blood (PB). The results reveal that the novel planar lipid bilayer technique enables the study of the biophysical properties of primary human T cells derived from blood and tissues in the context of health and disease.

The protocol describes a technique to study the ability of primary polyclonal human T cells to form synaptic interfaces using planar lipid bilayers. We use this technique to show the differential synapse formation capability of human primary T cells derived from lymph nodes and peripheral blood.

The current understanding of the dynamics and structural features of T-cell synaptic interfaces has been largely determined through the use of ...

Pneumococcus Infection of Primary Human Endothelial Cells in Constant Flow

Interaction of Streptococcus pneumoniae with the surface of endothelial cells is mediated in blood flow via mechanosensitive proteins such as the Von Willebrand Factor (VWF). This glycoprotein changes its molecular conformation in response to shear stress, thereby exposing binding sites for a broad spectrum of host-ligand interactions. In general, culturing of primary endothelial cells under a defined shear flow is known to promote the specific cellular differentiation and the formation of a stable and tightly linked endothelial layer resembling the physiology of the inner lining of a blood vessel. Thus, the functional analysis of interactions between bacterial pathogens and the host vasculature involving mechanosensitive proteins requires the establishment of pump systems that can simulate the physiological flow forces known to affect the surface of vascular cells.
The microfluidic device used in this study enables a continuous and pulseless recirculation of fluids with a defined flow rate. The computer-controlled air-pressure pump system applies a defined shear stress on endothelial cell surfaces by generating a continuous, unidirectional, and controlled medium flow. Morphological changes of the cells and bacterial attachment can be microscopically monitored and quantified in the flow by using special channel slides that are designed for microscopic visualization. In contrast to static cell culture infection, which in general requires a sample fixation prior to immune labeling and microscopic analyses, the microfluidic slides enable both the fluorescence-based detection of proteins, bacteria, and cellular components after sample fixation; serial immunofluorescence staining; and direct fluorescence-based detection in real time. In combination with fluorescent bacteria and specific fluorescence-labeled antibodies, this infection procedure provides an efficient multiple component visualization system for a huge spectrum of scientific applications related to vascular processes.

This study describes the microscopic monitoring of pneumococcus adherence to von Willebrand factor strings produced on the surface of differentiated human primary endothelial cells under shear stress in defined flow conditions. This protocol can be extended to detailed visualization of specific cell structures and quantification of bacteria by applying differential immunostaining procedures.

Interaction of Streptococcus pneumoniae with the surface of endothelial cells is mediated in blood flow via mechanosensitive proteins such as the Von ...

P. aeruginosa Infected 3D Co-Culture of Bronchial Epithelial Cells and Macrophages at Air-Liquid Interface for Preclinical Evaluation of Anti-Infectives

fDrug research for the treatment of lung infections is progressing towards predictive in vitro models of high complexity. The multifaceted presence of bacteria in lung models can re-adapt epithelial arrangement, while immune cells coordinate an inflammatory response against the bacteria in the microenvironment. While in vivo models have been the choice for testing new anti-infectives in the context of cystic fibrosis, they still do not accurately mimic the in vivo conditions of such diseases in humans and the treatment outcomes. Complex in vitro models of the infected airways based on human cells (bronchial epithelial and macrophages) and relevant pathogens could bridge this gap and facilitate the translation of new anti-infectives into the clinic. For such purposes, a co-culture model of the human cystic fibrosis bronchial epithelial cell line CFBE41o- and THP-1 monocyte-derived macrophages has been established, mimicking an infection of the human bronchial mucosa by P. aeruginosa at air-liquid interface (ALI) conditions. This model is set up in seven days, and the following parameters are simultaneously assessed: epithelial barrier integrity, macrophage transmigration, bacterial survival, and inflammation. The present protocol describes a robust and reproducible system for evaluating drug efficacy and host responses that could be relevant for discovering new anti-infectives and optimizing their aerosol delivery to the lungs.

P. aeruginosa Infected 3D Co-Culture of Bronchial Epithelial Cells and Macrophages at Air-Liquid Interface for Preclinical Evaluation of Anti-Infectives

We describe a protocol for a three-dimensional co-culture model of infected airways, using CFBE41o- cells, THP-1 macrophages, and Pseudomonas aeruginosa, established at the air-liquid interface. This model provides a new platform to simultaneously test antibiotic efficacy, epithelial barrier function, and inflammatory markers.

fDrug research for the treatment of lung infections is progressing towards predictive in vitro models of high complexity. The multifaceted presence of ...

Co-Culture of Murine Small Intestine Epithelial Organoids with Innate Lymphoid Cells

Complex co-cultures of organoids with immune cells provide a versatile tool for interrogating the bi-directional interactions that underpin the delicate balance of mucosal homeostasis. These 3D, multi-cellular systems offer a reductionist model for addressing multi-factorial diseases and resolving technical difficulties that arise when studying rare cell types such as tissue-resident innate lymphoid cells (ILCs). This article describes a murine system that combines small intestine organoids and small intestine lamina propria derived helper-like type-1 ILCs (ILC1s), which can be readily extended to other ILC or immune populations. ILCs are a tissue-resident population that is particularly enriched in the mucosa, where they promote homeostasis and rapidly respond to damage or infection. Organoid co-cultures with ILCs have already begun shedding light on new epithelial-immune signaling modules in the gut, revealing how different ILC subsets impact intestinal epithelial barrier integrity and regeneration. This protocol will enable further investigations into reciprocal interactions between epithelial and immune cells, which hold the potential to provide new insights into the mechanisms of mucosal homeostasis and inflammation.

This protocol offers detailed instructions for establishing murine small intestine organoids, isolating type-1 innate lymphoid cells from the murine small intestine lamina propria, and establishing 3-dimensional (3D) co-cultures between both cell types to study bi-directional interactions between intestinal epithelial cells and type-1 innate lymphoid cells.

Complex co-cultures of organoids with immune cells provide a versatile tool for interrogating the bi-directional interactions that underpin the delicate ...