Contact-Free Co-Culture Model for the Study of Innate Immune Cell Activation During Respiratory Virus Infection

NOTE: Refer to Table 1 for recipes of media used in this protocol.
NOTE: hNECS grown on 12-well transwell have been found to grow into more optimal thickness for soluble factors to reach the basal chamber readily when infected with Influenza virus. Hence use of 12-well sized transwell for co-culture is recommended. 

1. Establishment of the 3T3 feeder layer

1. Establishment from frozen stocks
   1. Thaw a cryovial of NIH/3T3 fibroblasts from frozen stocks. Add the contents of the cryovial to 2 mL of complete Dulbecco's Minimal Essential Media (DMEM) and resuspend the cells.
   2. Centrifuge for 5 min at 300 × g and room temperature/pressure (rtp), and remove the supernatant. Resuspend the cells in complete DMEM.
   3. Count the cells using trypan blue staining. Add 10 µL of trypan blue to 10 µL of the resuspended cell suspension. Mix thoroughly, and add 10 µL of the suspension to a hemocytometer to count the cells.
   4. Seed cells at a density of 1 × 10⁴ cells/cm² in an appropriate culture dish (e.g., T75 flask). Incubate the resulting culture flask for 3 days at 37 °C in a 5% CO₂ atmosphere.
2. Mitomycin C treatment
   1. At 3 days, ensure confluency is 60%-80%, remove the medium, and wash the cells with 1x phosphate-buffered saline (PBS).
      ​NOTE: It is important that the 3T3 feeder layer does not reach full confluency; otherwise, the mitomycin C treatment will not be effective.
   2. Add mitomycin C-supplemented complete DMEM to the flask, and incubate for 3.5 h at 37 °C in a 5% CO₂ atmosphere. Remove the mitomycin C-containing medium, and wash the cells 2x with 1x PBS.
3. Seeding 3T3 cells in 6-well plates
   1. Add 3 mL of 1x trypsin-EDTA to the flask for 3-5 min to disassociate the cells. Collect the disassociated cells in a fresh 15 mL tube. Centrifuge the 15 mL tube for 5 min at 300 × g, rtp; discard the supernatant. Resuspend the cells in complete DMEM.
   2. Count the cells using trypan blue staining. Seed the cells in a 6-well plate at 7.5 × 10⁵-2.5 × 10⁶ cells/well. Incubate the plate overnight at 37 °C in a 5% CO₂ atmosphere.
      ​NOTE: The 3T3 feeder layer is considered ready if the cells are healthy. At this point, seed the human nasal epithelial stem/progenitor cells (hNESPCs) onto the feeder layer for expansion.
   3. Add Complete DMEM to the flask and incubate at 37°C, 5% CO² overnight for 3T3 cells to recover from mitomycin C treatment.

2. Establishment of human nasal epithelial cell (hNEC) culture

NOTE: Clinical samples should be obtained from patients who are free of symptoms of upper respiratory tract infection.

1. Processing nasal tissue into single-cell suspension
   1. Wash nasal tissue in 1x Dulbecco's PBS (dPBS) (PBS without Mg²⁺ and Ca²⁺) containing 100 µL/mL of an antibiotic-antimycotic mixture. Cut the tissue into small fragments, and treat the sample in 10 mg/mL of a neutral protease overnight at 4 °C with shaking.
   2. Centrifuge for 5 min at 200 × g, and remove the supernatant after centrifugation. Incubate the pellet in 1-2 mL of 1x trypsin-EDTA (37 °C, 15 min), and quench by adding a volume of fetal bovine serum equal to 10% of the volume in the tube.
   3. Mechanically dissociate the digested tissue into single-cell aggregates by pipetting, and then pass the resulting suspension through a 70 µm cell strainer. Resuspend the cells in complete DMEM.
      ​NOTE: After this step, the cell suspension is a mixture of terminally differentiated cells and stem cells that are referred to as human nasal epithelial stem/progenitor cells (hNESPCs). The objective of the subsequent steps is to select for and enrich the hNESPC population from the mixture of different cell populations.
2. Seeding a single-cell suspension onto the 3T3 feeder layer for selection of hNESPCs
   1. Centrifuge the cell suspension from step 2.1.3 (300 × g, 5 min, rtp), and remove the supernatant. Resuspend the cells in 3-5 mL of medium 3.
      ​NOTE: Medium 3 is formulated to promote the selective growth of hNESPCs.
   2. Count the cells via trypan blue staining. Seed 1 x 10⁶ cells in 2mL of medium 3 per well onto 6-well plate containing prepared 3T3 feeder layer after removing the media in the 6 well plate. Incubate the resulting co-culture at 37 °C in a 5% CO₂ atmosphere.
      ​NOTE: After seeding, take care not to agitate the plate as it will affect the attachment of the hNESPCs.
   3. Change medium 3 (2 mL) every 2-4 days by removing the old medium entirely and replacing it with fresh medium 3.
      ​NOTE: Medium is changed to replenish nutrients and to prevent overly acidic conditions from negatively influencing the growth of the hNESPCs. The interval between each medium change depends on how acidic (yellow) the medium becomes. Intervals should be shortened if the medium turns acidic quickly.
   4. After 7-10 days, observe that hNESPCs are at a confluency suitable for transferring them onto membrane inserts. See Figure 1 for representative morphology of hNESPCs at 3 different timepoints (2 days, 5 days, and 10 days).
3. Transferring hNESPCs onto membrane inserts
   ​NOTE: Due to the mitomycin C treatment, the 3T3 feeder layer will slowly degrade over the hNESPC expansion period. This will result in a weakened adhesion to the surface of the well, allowing their dislodgement by flushing with a pipette, leaving behind only the healthy hNESPCs.
   1. Remove the medium, and add 500 µL of 1x dPBS to each well. Flush the cells 3x with a micropipette to dislodge the 3T3 cells, and discard the 1x dPBS. Observe under the microscope that the round hNESPCs remain while the spindle-shaped 3T3 feeder layer is dislodged.
   2. Add 500 µL of a cell disassociation reagent per well, and incubate at 37 °C in a 5% CO₂ atmosphere for 5-10 min, or until the hNESPCs have detached from the well surface.
   3. Collect the hNESPC suspension, centrifuge (300 × g, 5 min, rtp), and remove the supernatant.
   4. Resuspend the pellet in medium 3. Count the cells via trypan blue staining. Seed 3 × 10⁴ cells in 150 µL of medium 3 per membrane insert (24-well plate) or 1 × 10⁵ cells in 300 µL of medium 3 per membrane insert (12-well plate) (Figure 2). Incubate the cells at 37 °C in a 5% CO₂ atmosphere.
   5. Change the medium (medium 3) of the apical and basal chambers every 2 days. Navigate a pipette tip between the supporting arms of the membrane insert to access the basal chamber, remove the old medium, and then reintroduce fresh medium by the same method. Although the apical chamber is readily accessible, take care not to disturb the growing hNESPC layer.
      ​NOTE: Do not disturb the medium in the apical chamber for at least 2 days after seeding as the cells need time to attach to the membrane.
4. Differentiation of hNESPCs into hNECs
   1. When hNESPCs reach 100% confluency (approximately 3-7 days from seeding on the membrane insert), start the ALI culture. Change the basal medium to differentiation medium, and remove the medium from the apical chamber. Only add medium to the basal chamber without adding it to the apical chamber when changing the medium every 2-3 days, as indicated in Figure 3 and step 2.3.5.
      NOTE: Medium is changed to differentiation medium to promote the differentiation of the hNESPCs into hNECs in ALI. The hNESPCs take approximately 3-4 weeks to reach full maturity to become hNECs in ALI, at which point the cells would have grown into a multilayer with different populations of cells in each layer. Cilia should also be observed at a magnification of 400x (cilia can be identified by their beating movement, which cause the microscope field to appear like it is vibrating). At this point, the cells are ready to be used for the co-culture experiments. Refer to Figure 4 for the cross-section of a fully differentiated hNEC layer.
   2. After obtaining mature hNECs, wash the cells in the apical chamber 3x with 1x dPBS at 2-3-day intervals for 1 week prior to the co-culture experiment. Synergize the wash step with the basal medium changes, and perform the washes as follows.
      1. Add 50 µL (24-well)/150 µL (12-well) of 1x dPBS into the apical chamber of the membrane insert, and incubate the cells at 37 °C for 10 min. Remove the dPBS after 10 min of incubation to remove accumulated mucus and dead cells over the course of differentiation.
         ​NOTE: Depending on the nature of the experiment, sodium bicarbonate solution can be used to wash off the mucus layer entirely. However, for viral infection in co-culture experiments, the mucus layer is retained, and only excess mucus is removed with dPBS wash. This retention is to mimic the physiological mucus layer present on the nasal epithelia that will interact with the incoming viral infection.

3. Transepithelial electrical resistance (TEER) measurement

NOTE: Confirmation of epithelial integrity is important to ensure that an intact and healthy epithelial layer is obtained. An intact epithelial layer is determined through TEER measurement performed on 4 random wells using a voltohmmeter.

1. Rinse the electrode with 70% ethanol, and sterilize it with ultraviolet light for 15 min before use to ensure sterility. Rinse with 1x dPBS after sterilization.
2. Add 100 µL (for 24-well) or 300 µL (for 12-well) of 1x dPBS to the apical chamber of the membrane insert. Incubate the plate at 37 °C for 10 min; then, remove the dPBS.
3. For each well to be measured, prepare an unused well with 1 mL (for 24-well) or 3 mL (for 12-well) of prewarmed (to 37 °C) differentiation medium. Place the membrane insert in the prepared well, and add 200 µL (for 24-well) or 600 µL (for 12-well) of prewarmed differentiation medium to the apical chamber. Equilibrate at 37 °C (subject to the incubation temperature of the type of virus added, e.g., 35 °C for Influenza) for 15 min.
   NOTE: Prepare a blank (empty membrane insert) in the same way for the calculation of the TEER.
4. Equilibrate a 15 mL tube of prewarmed differentiation medium at the same temperature for 15 min as well. Place the electrodes in the 15 mL tube, and switch on the voltohmmeter.
   NOTE: At this point, the reading should be 0 resistance, as the electrodes are in the same medium. If any other reading is obtained, equilibrate the electrode in the 15 mL tube until the reading is 0.
5. Starting from the blank, position the electrodes in each well such that one electrode is submerged in the apical chamber medium and the other in the basal chamber medium. Record the reading only when a constant reading is obtained for a period of at least 5 min.
6. After each measurement, wash the electrode with PBS before the next measurement. For each well tested, take measurements for three samples at different positions of the membrane. To calculate the net TEER of each sample, subtract the background resistance, given by the blank membrane insert, from each measurement.
7. Calculate the total TEER reading for a well using the following equation:
   Epithelial Integrity (Ωcm²) = Net TEER(ohms) × Area of membrane (cm²)
   NOTE: The TEER values of hNECs for viral infection experiments should be >1000 Ωcm^2 13,15,16.

4. Isolation of peripheral blood monocytes and NK cells

NOTE: Blood samples should be obtained from healthy volunteers and used on the same day of isolation.

1. Collect 30-40 mL of whole blood from each donor in 10 mL blood collection tubes.
2. Isolate PBMCs using density gradient centrifugation, and obtain PBMCs from the buffy coat after the following steps (see also the Table of Materials).
   1. Thoroughly mix ~10 mL of blood from the blood collection tube in a vacutainer before diluting with an equal volume of balanced salt solution (PBS).
   2. Add 15 mL of density gradient medium to the base of a 50 mL tube.
      ​NOTE: The ratio of density gradient medium to diluted blood should be 2:3-3.5.
   3. Layer 35 mL of diluted blood on the density gradient medium with a pipette gun (with the dispense setting set to the lowest possible) by tilting the tube by 45° and allowing the diluted blood to fall dropwise onto the inner wall of the tube so that the layer of density gradient medium is not disturbed.
   4. Centrifuge lysate at 500 × g, 18-20 °C for 30 min (brake: off). Carefully remove and discard the upper plasma layer without disturbing the lower layer.
   5. Transfer the buffy coat to a new tube without mixing the red blood cell layer, gradient density medium layer, or the buffy coat. Once the buffy coat has been collected in a new 50 mL tube, top up the volume to 50 mL with 1x PBS.
   6. Centrifuge lysate at 800 × g, 18-20 °C, 8 min (brake: off), and remove the supernatant with a pipette gun. Resuspend the cell pellet with 50 mL of 1x PBS, and centrifuge at 120 × g, 18-20 °C, 10 min to remove platelets.
   7. Discard the supernatant by pipetting, without disturbing the cell pellet.
      ​NOTE: As the cell pellet might be loose, it is inadvisable to discard the supernatant by pouring.
   8. Resuspend the cell pellet with complete RPMI (Roswell Park Memorial Institute) medium. Perform a cell count by adding 10 µL of 3% acetic acid with methylene blue to 10 µL of the resuspended cell suspension. Mix thoroughly, and add 10 µL of the mixture to a hemocytometer to count the cells.
3. Dilute PBMCs with complete RPMI medium to a density of 2 × 10⁶/mL (for 24-well) or 4 × 10⁶/mL (for 12-well).

5. hNEC Viral infection and transition to hNEC:PBMC co-culture

NOTE: H3N2 (A/Aichi/2/1968) is used as the representative strain of infection in this protocol. Multiplicity of infection (MOI) of 0.1 is used as the representative MOI in this protocol.

1. Day 0 (Infection of hNECs)
   NOTE: One well from the hNECs grown from the same donor is used to obtain a representative cell count for every well used in the experiment.
   1. In the representative well, add 150 µL of 1x trypsin-EDTA to the apical chamber of the membrane insert and 350 µL of 1x trypsin-EDTA to the basal chamber, and incubate at 37 °C for 10 min, or until the cells detach from the membrane.
   2. Flush the cells on the membrane by pipetting up and down, and collect the suspension in a 1.5 mL centrifuge tube. Add 200 µL of complete DMEM to quench trypsin activity. Count cells via trypan blue staining to obtain the cell count per well.
   3. Calculate the required MOI of the virus based on the cell count per well, and dilute the virus stock accordingly with complete RPMI on ice.
      NOTE: MOI 0.1 of 1.26 × 10⁶ hNECs per well = 1.26 × 10⁵ H3N2 viral particles per well in 30 µL (for 24-well)/100 µL (for 12-well)
   4. For the remaining wells for the infection experiment, add 50 µL (for 24-well)/150 µL (for 12-well) of 1x dPBS into the apical chambers of the membrane inserts, incubate at 37 °C for 10 min, and remove the 1x dPBS.
   5. Change the basal medium in the membrane inserts to complete RPMI medium by transferring the inserts to a new plate with complete RPMI added to the wells (350 µL (for 24-well)/700 µL (for 12-well)).
      NOTE: When hNESPCs have fully differentiated to hNECs, they are tolerant/permissive to different media for up to 72 h, with no changes to morphology or organization when the medium is switched to RPMI¹². RPMI is used to support the growth and maintenance of the PBMC population.
   6. Add the prepared 30 µL (for 24-well)/100 µL (for 12-well) of the virus inoculum into the apical chamber of the membrane insert, incubate at 35 °C in a 5% CO₂ atmosphere for 1 h, and remove the viral inoculum from the apical chamber.
   7. Change the basal medium of membrane insert to fresh complete RPMI medium and incubate for at 35 °C in a 5% CO₂ atmosphere for 24 h.
2. Day 1 (establishment of hNECs + PBMCs co-culture)
   1. Seed the required number of PBMCs in 150 µL (for 24-well)/300 µL (for 12-well) of complete RPMI medium by directly adding the PBMC suspension into the basal chamber of each well of uninfected or infected hNECs from Day 0 (1.5 × 10⁶ for 24-well plates and 3 × 10⁶ for 12-well plates). Incubate at 37 °C for 24/48 h.
      NOTE: The final volume in the basal chamber after the establishment of the co-culture is 500 µL (for 24-well)/1000 µL (for 12-well).
3. Days 2-3 (harvesting of PBMCs 48/72 h post-viral infection)
   1. Collection of apical supernatants (48/72 h post-viral infection)
      1. Add 50 µL (for 24-well)/150 µL (for 12-well) of 1x dPBS to each apical chamber, and incubate at 37 °C for 10 min. Collect the 1x dPBS in 1.5 mL tubes. Aliquot 25 µL (for 24-well)/50 µL (for 12-well) of the supernatant for plaque assay in a new 1.5 mL tube, and immediately freeze both the stock and the aliquots at -80 °C.
   2. Collection of cellular RNA from hNECs (48/72 h post-viral infection)
      1. Transfer the membrane insert to a clean well, add 300 µL (for 24-well)/600 µL (for 12-well) of RNA lysis buffer to the apical chamber, and incubate at rtp for 5 min. Collect the supernatant in a 1.5 mL centrifuge tube, and store it at -80 °C until RNA extraction for molecular analysis.
   3. Harvesting PBMCs (48/72 h post-viral infection)
      1. With the broad base of a sterile pipette tip, gently scrape the surface of the well to dislodge the activated PBMCs that may be adherent to the well surface. Collect the basal medium containing PBMCs in a 2 mL centrifuge tube.
      2. Flush the wells 2x with 300 µL of 1x dPBS, and collect the wash in the same 2 mL tube. Centrifuge the 2 mL tube (500 × g, 5 min, rtp), and collect the supernatant in a fresh 2 mL tube without disturbing cell pellet. Store the supernatant at -80 °C for cytokine and chemokine analysis; resuspend the cell pellet in 200 µL of 1x dPBS.

6. Flow cytometry

NOTE: This section of the protocol is continued directly from the previous section using the PBMC cell suspension from step 5.3.3.2. Ensure minimal light exposure during the following steps in this section. A sample panel of surface staining markers can be found in Table 2.

1. Surface staining
   1. Transfer the resuspended PBMC cell pellet into a 96-V bottom well plate. Centrifuge lysate at 800 × g for 3 min, and remove the supernatant.
   2. Incubate all the PBMCs (except for the "unstained") with 100 µL of a viability stain for 15 min at rtp. Top up to 200 µL with 150 µL of Magnetic-Activated Cell Sorting (MACS) buffer. Centrifuge lysate at 800 × g for 3 min, and discard the supernatant.
   3. Prepare a panel of surface staining antibodies of interest with appropriate dilution ratios and a final volume of 50 µL per reaction (top-up with MACS buffer to obtain the final volume). Perform surface staining by adding 50 µL of the prepared antibody mix to the cells using a multi-channel pipette. Incubate at 4 °C for 15 min in the dark.
   4. Wash by adding 150 µL of MACS buffer. Centrifuge lysate at 800 × g for 3 min, and discard the supernatant. If intracellular staining is not required, proceed directly to the 15 min incubation in step 6.3.
2. Intracellular staining (If required)
   1. Add 100 µL of a fixation and permeabilization solution to each well. Incubate at 4 °C for 20 min in the dark. Top up the wells with 100 µL of 1x MACS buffer.
   2. Centrifuge (800 × g, 3 min, 25 °C), and remove the supernatant. Repeat the wash by adding 200 µL of 1x Permeabilization Wash buffer. Centrifuge (500 × g, 3 min, 25 °C), and remove the supernatant.
   3. Prepare the dilutions for the antibody panel of interest to achieve a final volume of 50 µL using 1x Permeabilization Wash buffer. Incubate on ice for 30 min in the dark. Add 200 µL of 1x Permeabilization Wash buffer.
   4. Centrifuge the cells (800 × g, 3 min, 4 °C), and remove the supernatant. Resuspend the cells in 100 µL of fluorescence-activated cell sorting buffer.
3. Pipette 200 µL of a lysing solution into each well. Incubate for 15 min at rtp. Centrifuge lysate at 800 × g for 3 min and discard the supernatant.
4. Resuspend the cell pellets in 200 µL of MACS buffer. Perform flow cytometry in accordance to the settings outlined previously¹³, or store at 4 °C in the dark.

7. Determination of cytokine and chemokine levels

1. Process 25 µL of the basal chamber supernatant using an immunology multiplex assay according to the manufacturer's protocol¹⁸.
2. Calculate the concentration of each analyte using multiplex manager software utilizing a curve-fitting algorithm (5 parameters) for the standard curve.

8. Assessment of viral contamination

1. Extract viral RNA using an extraction kit on 4 sets of solutions: the stock H3N2 solution, the MOI 0.1 dilution for infection, the 10x serial dilutions of the MOI 0.1, and the basal medium from the samples (both 48 and 72 h post-viral infection)¹².
2. Select non-structural gene (NS1) and matrix (M1) as targets for polymerase chain reaction (PCR) (see the Table of Materials for primers used in the representative experiment).
3. Perform reverse-transcription-PCR (RT-PCR) (42 °C, 60 min) to convert the viral RNA to cDNA before performing quantitative PCR (qPCR) to measure NS1 and M1 levels as a proxy for viral presence, and correlate the levels to the standard curve obtained from the RT-qPCR performed on the 10x serial dilution of the MOI 0.1 aliquot. Refer to Table 3 for the composition of the reaction mixes, and use the following conditions: preincubation at 95 °C for 10 min, followed by a 3-step amplification for 40 cycles: 95 °C for 10 s, 60 °C for 10 s, 72 °C for 30 s; melting curve: 95 °C for 10 s, 65°C for 60 s, and 97°C for 1 s.

9. Plaque assay

1. Day 0: seeding MDCK (Madin Darby Canine Kidney) in 24-well plates
   1. Remove the medium from a T75 Flask of confluent MDCK cells. Wash 2x with 1x PBS to remove all traces of serum. Trypsinize the MDCK cells by adding 10 mL of trypsin and incubating at 37 °C in a 5% CO₂ atmosphere for 20-30 min until the cells detach.
      ​NOTE: Do not tap the flask as this might result in clumping.
   2. Pipette the cell suspension into a 15 mL tube containing 2 mL of complete Eagle's Minimal Essential Medium (EMEM). Centrifuge (300 × g, 5 min, rtp), and remove the supernatant.
   3. Resuspend the cells in 6 mL of complete EMEM. Count the cells by trypan blue staining.
   4. Dilute the MDCK cell suspension to a concentration of 1 × 10⁵ cells/mL, and seed 1 mL of the diluted suspension in each well of a sterile 24-well plate. Incubate at 37 °C in a 5% CO₂ atmosphere for 24 h to obtain a confluent monolayer of MDCK cells in each well.
2. Day 1: infection of MDCK cells
   1. Prepare the infection medium. Remove the medium from the MDCK monolayer in the 24-well plate, and wash 2x with 1x dPBS. For the second wash, leave the PBS in the wells while preparing serial dilutions of the virus.
   2. Thaw the virus samples on ice, and serially dilute them in a 24-well plate to achieve serial dilutions from 10^-1 to 10^-6.
      ​NOTE: As an example, fill each well in a 24-well plate with 270 µL of infection medium.
   3. Add 30 µL of the virus sample to the first well in the row. With a new pipette tip, mix well and transfer to the next well in the row to dilute the sample by 10x. Proceed until 10^-6 dilution has been achieved; perform steps 9.2.1-9.2.3 for all virus samples.
   4. Remove PBS from the MDCK plate, and infect in duplicate with 100 µL of the prepared viral dilutions. For control wells, add 100 µL of the infection medium (without the viral sample). Incubate at 35 °C in a 5% CO₂ atmosphere for 1 h, shaking the plate to eliminate dry spots every 15 min.
   5. Remove the viral inoculum, and add 1 mL of liquid overlay for each well. Incubate at 35 °C in a 5% CO₂ atmosphere for 72 h.
3. Day 4 (72 h post-infection): plaque visualization
   1. Remove the liquid overlay, and fix the cells with 4% formaldehyde in 1x PBS for 1 h. Remove the formaldehyde solution, and wash once with 1x PBS or distilled water.
   2. Stain the fixed cells by adding 1% crystal violet solution for 15 min. Remove the crystal violet dye, and wash the cells with running water. Dry the plate at rtp.
   3. Once dry, count plaques, and calculate the viral titer according to the following formula:
      Number of Plaques × Dilution Factor = Number of Plaque – Forming Units in 100 μL