An Air-liquid Interface Bronchial Epithelial Model for Realistic, Repeated Inhalation Exposure to Airborne Particles for Toxicity Testing

1. Preparing cell culture medium (CCM)

1. Prepare a bottle of 500 mL of minimum essential medium (MEM) supplemented with glutamine.
2. Add 5 mL of penicillin-streptomycin (i.e., 100 U/mL penicillin and 100 µg/mL streptomycin).
3. Add 5 mL of non-essential amino acids (NEAA) solution.
4. Add 10 mL of amphotericin B (optional).
5. Add 50 mL of FBS (heat inactivated, please follow the ATCC protocol for heat inactivation; (https://www.atcc.org/~/media/PDFs/Culture%20Guides/AnimCellCulture_Guide.ashx, page 19)).

2. Subculturing of Calu-3 cells

NOTE: Calu-3 cells are cultured in T75 or T175 cell culture flasks at 37 °C and 5% CO₂. Cells are passaged at 60–80% confluency every 7 days with CCM renewal every 2–3 days. CCM is poured off and fresh CCM (T25 = 5 mL, T75 = 15 mL, and T175 = 25 mL) is pipetted into the flask and the flask is placed back into the incubator. Cells should be passaged at least 2x after thawing, before using in experiments, or before freezing, and they should be passaged no more than 25x in total.

1. Confirm if flask is 60–80% confluent by checking under a light microscope.
2. Pour off the CCM from the flask.
3. Wash the cells 2x with 5 mL of 1x Hanks’ Balanced Salt Solution (HBSS) without calcium and without magnesium. Discard the HBSS after each wash. HBSS removes serum, which inhibits trypsin.
4. Add 3 mL of trypsin-EDTA for a T75 (4 mL for a T175) and place the flask back into the incubator at 37 °C and 5% CO₂ for 10–15 min. Check after 10 min, ensuring the cells have become detached from the flask surface. In case the cells are grown to >80% confluency, they will not detach using trypsin 0.05% and trypsin 0.25% could be used.
5. Add 6 mL (i.e., double the trypsin-EDTA volume originally added) of CCM to the flask and gently rock the flasks to ensure proper mixing. This is to ensure the trypsin has been neutralized by the FBS in the CCM and its activity on the cells halted. If trypsin is allowed to remain in contact with the cells for too long they will not reattach when put into a new cell culture flask.
6. Pour the complete contents of the flask into a 50 mL centrifuge tube.
7. Centrifuge the cells for 5 min at 130 x g, ensuring that the centrifuge is correctly balanced.
8. Return the vial containing the cells back to aseptic conditions and remove the supernatant gently, without disturbing the pellet. The supernatant can be poured off and the remainder pipetted off, ensuring the pellet is not disturbed.
9. Resuspend the cell pellet in 1 mL of CCM by pipetting up and down until all cells are suspended (i.e., no pellet or cell agglomerates are observed). Additional CCM can be added to dilute the cell suspension.
10. Count the cells, both dead and alive, in 1 mL of CCM using a hemocytometer. If needed for proper counting, dilute the cells in 3 or 4 mL.
11. Suspend the cells into the CCM volume required and add the cell suspension into each flask. To achieve about 80% confluency in a week, seed 2 x 10⁶ cells in a T75 or 6 x 10⁶ cells in a T175.
12. Gently rock the flask and then place it back into the incubator at 37 °C and 5% CO₂.
13. Replace with fresh CCM every 2–3 days and subculture when the cells reach 60–80% confluency.

3. Seeding Calu-3 cells onto culture inserts

NOTE: Inserts are available with different pore sizes. Small pore sizes (e.g., 0.4 µm) have the advantage that the cells grow more easily and can achieve a good barrier already after 5 days culturing under submerged conditions, as measured by Trans Epithelial Electrical Resistance (TEER). However, when interested in particle translocation, these pores are too small and will trap the particles. Therefore, larger pore sizes (e.g., 3 µm) are usually chosen to test particles. When using a larger pore size, the cells need longer time periods (e.g., 7–10 days culturing under submerged conditions) to achieve a good TEER.

1. Prepare cell suspension with known concentration following steps 2.1–2.10.
2. Dilute cells to a concentration of 5 x 10⁵ cells/mL in prewarmed CCM for 6 well inserts or 2.5 x 10⁵ cells/mL for 12 well inserts.
3. Take a cell culture plate with inserts and place under aseptic conditions.
4. Fill the basolateral side with 2 mL of prewarmed CCM for 6 well inserts, or 1 mL for 12 well inserts. While adding the culture medium, take the insert out using tweezers.
5. Carefully mix the cell suspension by pipetting up and down. Pipette 1.0 mL for 6 well inserts and 500 µL for 12 wells inserts (equivalent to 100,000 cells/cm²) on the top of the membrane in the cell culture insert with 0.4 µm pores. For 3.0 µm pores, increase the cell density to 128,000 cell/cm².
6. Cover the cell culture plate and incubate at 37 °C and 5% CO₂.
7. Change the CCM every 2–3 days.
8. Let the cells become confluent for 7 days under submerged conditions before continuing to culture at the ALI.
9. Measure TEER.
   1. Take an Epithelial Voltohmmeter supplemented with Chopstick Electrode Set and charge the battery system overnight.
   2. Disconnect the Voltohmmeter from the charger and connect the chopstick electrode.
   3. Clean the electrode with 70% ethanol.
   4. Place the electrode in the CCM by putting the longer electrode in the external culture media until it touches the bottom of the dish and putting the shorter electrode in the media without touching the membrane.
   5. Start with an empty insert without cells. Wait until the measurement stabilizes and write down the resistance in Ohms. This measurement is the resistance of the insert membrane without any cells (i.e., blank resistance).
   6. Repeat the measurement for each insert and subtract the blank resistance to obtain the true resistance.
   7. For data analysis, multiply the resistance values by the surface area of the insert into Ω x cm². For a 6 well insert, the surface area is 4.67 cm². Thus, if a resistance of 600 Ohm is measured and the background is 120 Ohm, the resistance is 480 Ohm, which is then multiplied by the surface area of 4.67 cm² for a total of 2,241.6 Ohm x cm². The TEER should be >1,000 Ω x cm² to continue.
10. Remove the CCM from the apical side of the inserts.
11. Add 2 mL of prewarmed CCM for 6 well and 1 mL for 12 well inserts to the basolateral side of the well (i.e., under the cell culture insert). The CCM should touch the membrane from the bottom, but not leak onto the top of the insert.
12. At this point cells are apically exposed to air, which is referred to as culturing at the ALI.
13. Culture cells at the ALI in the incubator at 37 °C and 5% CO₂ for 7 days prior to exposure.
14. Change the basolateral CCM every 2–3 days. The cells can be used at the ALI for 6 weeks.

4. Preparing the exposure setup

NOTE: Sections 5–7 describe preparations for particle exposure using an automated exposure station (AES, see Table of Materials). The setup for particle nebulization and characterization is also compatible with other ALI exposure systems from other manufacturers. As an example, the exposure to particles is described below. Such systems can also be used for other exposures, such as sensitizers, cigarette smoke, and diesel exhaust. Figure 1 shows the AES and an exposure module. Figure 2 shows a schematic representation of the exposure setup including all other instruments.

1. Before starting an exposure using the AES, connect the system to several instruments to measure aerosol characteristics; these are measured in a side stream just before the aerosols enter the cabinet.
   NOTE: A flow splitter is used to connect the side stream to the exposure characterization equipment. Generally, the following equipment is used: scanning mobility particle sizer (SMPS), optical particle sizer (OPS), condensation particle counter (CPC), tapered element oscillating microbalance (TEOM). The SMPS and OPS measurements are performed 1x per hour and use the same tubing from the flow splitter. The CPC and TEOM perform continuous measuring and data from both are logged on a Squirrel model 2020 data logger. In addition, gravimetric mass concentration is determined using a microbalance in controlled relative humidity (40–70%) and temperature (21–23 °C) conditions. Teflon filters are weighed before and after each exposure to confirm the exposure concentration. To capture the exhaust, a HEPA filter is used. The setup including engineered nanomaterial (ENM) suspensions and nebulizer are all placed in a flow cabinet to prevent any exposure to people. The AES can be used for testing many different types of ENMs, including metals and metal oxides.
2. Prepare a nanomaterial suspension shortly before exposure. Usually a 1% suspension is prepared as a stock solution. For example, suspend 100 mg of nanomaterial in 10 mL of pure water.
   NOTE: For DQ12 exposure, 300 mg is used to achieve about 2 µg/cm². This amount can be used in a single exposure or divided over repeated exposures (e.g., 300 mg is suspended in 30 mL for a single exposure or 21.5 mg is suspended in 2.15 mL freshly every day for 3 weeks of repeated exposure). The suspension is freshly prepared on each exposure day. The particle suspension is sonicated for 16 min using probe sonication. The volume of the 1% stock solution is adjusted to a total volume of 100 mL by adding pure water.
3. Put the ENM suspension in a small bottle with a cap and a magnetic stirrer to prevent settling of the particles. Connect the bottle to a peristaltic pump via a small tube and adjust the flow to 25 mL/h.
4. Connect the peristaltic pump to a spray nozzle and adjust the settings to allow continuous aerosolization.
5. The spray nozzle is mounted to a 60 cm long aluminium tube (mixing chamber, diameter 15 cm, heated to 60 °C). The setup is connected to the AES via a 1.5 meter long copper tube (diameter 15 cm). On top of the AES an impactor removes all aerosols larger than 2.5 µm.
6. Connect the spray nozzle to 3 bar compressed air through two mass flow controllers (MFC) to allow nebulization of suspensions. One flow of 14 L/min is used for the spray nozzle, the other MFC for mixing of the air in the tube.
7. The day before the start of the exposure, turn on the AES to allow the cabinet to reach a temperature of 37 °C.
8. Turn on the air flow and the humidity in the cabinet 2 h before start of the exposure, to reach 85% humidity. Turn on the heating of the exposure chambers in which the inserts with the cells will be placed.
9. Turn on the quartz microbalance (QCM) and set the initial value at 0 using the software. Start logging. Every 10 s the mass is measured and expressed as ng/cm².
10. Warm the cell culture media to 37 °C in a water bath (~20–30 min).

5. Preparing Calu-3 cells for exposure

NOTE: For a typical ALI exposure using the AES, 15–20 inserts with a confluent cell layer are needed. These consist of 3 clean air controls, 3 incubator controls that will be handled similarly to the other inserts without exposure in the AES, 6–8 inserts for aerosol exposure (depending on the use of 0, 1, or 2 microbalances), 1–3 inserts for control measurements (such as maximum LDH release), and 3 spare inserts in case the TEER of some of the inserts is not sufficient. The cells should have a TEER of >1,000 Ω x cm² to continue.

1. On the first day of exposure, wash cells 1x with CCM, check cell morphology, and measure the TEER of the cell model using a Voltohmmeter. The cells should form a tight monolayer without gaps.
2. Put 2 mL/1 mL of HEPES buffered CCM without FCS to the basolateral side of 6 well/12 well plates and transfer the culture inserts with cells to the new plates.
   NOTE: During exposure, no CO₂ is present in the AES. Therefore, HEPES buffered culture medium (25 mM HEPES) is used during transport and exposure. This medium is used for both the exposed cells in the AES as well as the incubator control cells.
3. In case the time to transport the cell cultures to the AES is more than 5 min, put the cells in a portable incubator of 37 °C during transport.

6. Handling the AES during an exposure

1. At the AES, fill the exposure modules with HEPES-buffered CCM without fetal calf serum (FCS). The amount of CCM depends on the unit used, and the type of cell culture insert. Keep in mind that to keep cells at the ALI, the medium should reach the bottom of the membrane, but should not leak on top of the membrane. When using 6 well inserts, add 6 mL of HEPES-buffered CCM to the exposure modules.
2. Transfer the inserts with cells from the plates to the exposure modules using sterile tweezers. Check that there are no air bubbles at the basolateral side of the cells and remove any CCM on the apical side of the inserts. In case there are some air bubbles at the basolateral side, gently turn the inserts using the sterile tweezers until they are removed. Keep the plates containing CCM in an incubator for transfer after an exposure.
3. Use the touchscreen display to choose exposure duration, air flow rate, and electrostatic deposition enhancement. The display can also be used to check humidity and temperature. Usually, an exposure duration of 4 h is chosen, with a flow rate of 50 mL/min on the inserts at 37 °C and 85% humidity.
   NOTE: The modules in the first level (Figure 1) are used for clean air exposure; inserts in this level are used as clean air exposure controls. The other modules in the second and third level can be used for aerosol exposure, including two modules for quartz crystal microbalances (QCM) to measure deposition online.
4. The leak test should be conducted before starting exposure. The leakage needs to be less than 5 mL/min. Follow the instructions on the AES display. When the leak test has finished, the exposure can be started. In case of a leak, the tubing should be checked.
5. At the end of exposure, open the door of an AES module, open the exposure modules, place the cell culture inserts back to the cell culture plates, and transfer the plates to the portable incubator.
6. Collect the media from the modules (i.e., exposed samples) and from the plates (i.e., incubator controls) for later analysis, such as lactate dehydrogenase (LDH) measurement.
7. Back at the cell culture lab, transfer the cell culture inserts to plates filled with fresh standard CCM. Put the cell culture plates in the incubator until the next exposure or until analysis.
8. After the final exposure day, put the inserts in the incubator until the next day.
9. At the day after the final exposure, add CCM to the apical side to measure TEER using a Voltohmmeter. Collect both the apical and the basolateral CCM separately for analysis of cytokines.
10. Remove all CCM and perform a cell viability assay by adding, for example, a proliferation reagent to the apical side.

7. Cleaning the AES

1. Fill the exposure modules with water, wait for 1 min, and remove the water. Next, fill the modules with 70% ethanol, leave it for 10 min, and remove the ethanol. Clean the exposure trumpets also with 70% ethanol.
2. Stop the 85% humidity control, but leave the temperature of the cabinet at 37 °C for the next experiment.