In vitro Coculture Assay to Assess Pathogen Induced Neutrophil Trans-epithelial Migration

Steps (1-3) should be performed in a sterile environment under a laminar flow hood.

1. Collagen Coating Transwells

1. Make a 30 µg/ml collagen solution. Dilute 3 mg/ml collagen stock 1:100 in 60% ethanol that has been passed through a 0.2 µm filter unit.
2. Vortex diluted 30 µg/ml solution. Note: It is not necessary to vortex the 3 mg/ml collagen stock solution, as this solution is quite viscous and air bubbles may be introduced, potentially decreasing accuracy when pipetting.
3. Remove 0.33 cm² growth area transwells with a pore size of 3 µm from outside packaging and place the 24-well plate containing 12 transwells upside down inside the hood.
4. Lift off bottom plate and place aside. Note: Transwells will now be upside down resting atop the lid of the 24-well plate
5. Turn on Bunsen burner. Flame hemostat for sterility and let cool.
6. With sterile hemostat, transfer transwells to a 150 mm x 25 mm Petri dish, keeping them in the inverted position. Note: Be sure to maintain sterility of the empty 24-well plate plus lid to be used to house transwells once they have been seeded with epithelial cells.
7. Pipette 70 µl of the 30 µg/ml collagen solution onto the filter membrane of each inverted transwell. Note: Surface tension should hold this volume of solution in place as there are no walls around the filter membrane when accessing the surface of the underside. Be careful that the collagen solution does not drip down the side of the transwell and avoid moving transwells during the coating procedure.
8. Leave the lid of the Petri dish off for a minimum of 4 hr to allow ethanol solution to evaporate. To maintain sterility during this process keep the hood fan and the ultraviolet light on.
9. After the transwells have dried, they can be used immediately or stored at room temperature for up to a week provided the Petri dish cover is in place and sterility is maintained.

2. Passage Flask of Epithelial Cells for Seeding Transwells

(This protocol specifically describes the handling of the lung epithelial cell line H292 for the generation of epithelial barriers grown on transwells. Other epithelial cell lines can be used with slight modifications.)

1. Prepare culture medium by adding 10% heat-inactivated Fetal Bovine Serum (HI-FBS) and 1x Penicillin/Streptomycin to RPMI 1640.
2. Warm media, trypsin-EDTA (T-EDTA), and D-PBS in 37 ºC water bath.
3. Remove T-75 flask containing cells from incubator and assess confluence visually with inverted microscope. Note: When using H292 cells, flasks should be close to or fully confluent.
4. Aspirate media from flask. Reduce length of time that cells are dry by having next solution in hood with cap loosened.
5. Add 5 ml D-PBS to T-75 flask and swirl. Avoid creating bubbles. Remove PBS by aspiration.
6. Add 3 ml T-EDTA to T-75 flask and swirl to cover bottom.
7. Remove most of T-EDTA, leaving approximately 0.3 ml in T-75 flask.
8. Distribute the small remaining volume of T-EDTA over the surface area of the cells and place in a humidified incubator under standard culture conditions (37 ºC, 5% CO₂).
9. After a period of time, remove flask from incubator. Note: Cells should readily detach from the surface of the flask by gently tapping the flask on its side. Cells should now be rounded and floating when visualized under the microscope. The average time this will take for H292 cells is 5-10 min.
10. Add 10 ml culture media to flask to neutralize T-EDTA and resuspend cells. Draw up the cell solution into pipette tip and eject onto surface of flask containing cells approximately five times to resuspend all cells.
11. Transfer resuspended cells to a 15 ml tube for seeding transwells. Note: The concentration of resuspended H292 cells prepared in this manner should range between approximately 1 x 10⁶-1.8 x 10⁶ cells/ml.

3. Seeding Collagen Coated Transwells with Epithelial Cells

1. Add 70 µl of resuspended cell solution from the 15 ml tube to each inverted collagen coated transwell. Note: H292 cell solutions with a concentration range between 1 x 10⁶-1.8 x 10⁶ cells/ml will yield approximately 70,000-120,000 cells/transwell.
2. Mix the cell suspension by gently inverting tube periodically to prevent cells from settling to the bottom of the 15 ml tube while seeding the transwells. Note: Do not vortex cells and be cautious that pipette tip does not come in direct contact with collagen coated membrane filter.
3. Once all inverted transwells have been seeded with epithelial cells, replace Petri dish cover and carefully place in the tissue culture incubator, humidified, 37 ºC, 5% CO₂. Take care to avoid disturbing the cell suspension that has been added to the collagen coated filter membrane.
4. Maintain inverted transwells in the tissue culture incubator, humidified, 37 ºC, 5% CO₂ overnight. Note: The bubble of liquid containing cells should remain associated with the permeable filter membrane of the transwell throughout the overnight incubation. If the filter membrane becomes dry due to evaporation of liquid or because the liquid drips down the side of the transwells, the epithelial layers may not grow properly.
5. The following day, add 1 ml media into the original sterile 24-well plate and flip each inverted transwell using a sterilized hemostat into each well containing 1 ml of media.
6. Add 0.2 ml media into the top chamber of each transwell and return to tissue culture incubator, humidified, 37 ºC, 5% CO₂.
7. H292 inverted transwells can be used from 8 days up to 1 month following seeding. Note: We suggest waiting a minimum of 8 days post seeding of transwells to ensure that the epithelial monolayers have fully formed and exhibit a functional barrier. One can test the H292 epithelial barrier by determining whether the monolayer is capable of restricting trans-epithelial flux of the protein horse radish peroxidase (HRP) following addition to the basolateral surface¹⁸.

4. Preparation of Bacteria for Infection of Epithelial Layers on Transwells

1. One day prior to experiment, extract one colony of P. aeruginosa PAO1 from a Pseudomonas Isolation Agar plate and one colony of K12 E. coli MC1000 from a Luria-Bertani (LB) Agar plate and place into separate tubes containing 3 ml of LB broth. CAUTION: P. aeruginosa is a human pathogen, standard BSL2 safety measures should be adopted when handling this organism.
2. Shake overnight at 225 rpm in a 37 ºC environment.
3. Next day, remove 1 ml from the dense bacterial cultures and place into 1.5 ml Eppendorf tubes.
4. Spin in microfuge at 15,800 x g for 5 min and then remove supernatant.
5. Prepare Hanks’ Balanced Salt Solution with calcium and magnesium (HBSS+) in advance. Add 23.8 g HEPES and 97.5 g HBSS+ powder to 9.5 L distilled water. Add small quantities of 10 M NaOH to solution, while monitoring pH until a stable pH of 7.4 is reached. Bring volume up to 10 L and store at 4 ºC.
6. Resuspend each bacterial pellet with 1 ml HBSS+. Vortex to be sure an even bacterial suspension is achieved.
7. Spin again at 15,800 x g for 5 min and remove supernatant.
8. Resuspend the PAO1 pellet with 0.6 ml HBSS+ and the MC1000 pellet with 0.5 ml HBSS+. Vortex the bacterial suspensions.
9. Further dilute the PAO1 and MC1000 resuspended pellets 1:100 in HBSS+. For example, add 50 µl of the 0.6 ml PAO1 resuspended pellet to 5 ml HBSS+. Note: Using optical density (OD) measurements to determine culture density as well as standard colony forming unit (CFU) dilution plating methodology, PAO1 and MC1000 solutions prepared in this manner consistently yield between 3 x 10⁷-6 x 10⁷ CFU/ml. The density of a bacterial culture is positively correlated with the OD measurement of the culture at 600 nm and this correlation can be used to estimate the bacterial cell concentration. To confirm the bacterial cell concentration, the culture can be diluted to a very low estimated concentration and plated on an agar plate such that one would expect between 30-200 bacterial cells to be present on the plate. The cells are spread across the plate and incubated overnight at 37 ºC such that each cell forms an individual colony of cells that is large enough to be visible and colonies can then be counted.

5. Isolation of Neutrophils from Whole Blood

1. Acid citrate dextrose solution (ACD) is used as an anticoagulant and prepared by adding 6.9 g citric acid, 12.5 g sodium citrate, and 10 g dextrose to 500 ml distilled water. Once all components are dissolved, the ACD solution is sterilized by passing through a 0.2 µm filter unit. Store ACD solution at 4 ºC.
2. Prior to drawing blood, add 5 ml ACD to 50 ml syringe and attach 21 x ¾ G butterfly needle with 12 in tubing.
3. Apply tourniquet, wipe vein with alcohol, and insert needle. Note: Any research protocol involving human subjects must be approved by an institutional review board and written informed consent must be provided from each blood donor. For example, experiments conducted using this protocol were approved by Massachusetts General Hospital Institutional Review Board and assigned protocol #1999-P-007782.
4. Slowly draw 45 ml of blood ensuring that blood mixes with ACD once in the syringe.
5. When 45 ml has been drawn (50 ml total volume), untie the tourniquet before removing the needle. Immediately apply pressure to wound with sterile gauze when needle is removed.
6. Transfer blood slowly down the side of a 50 ml tube over approximately 5-10 sec.
7. Spin in centrifuge at 1,000 x g with the brake deactivated for 20 min at room temperature. Note: An additional step involving dilution of whole blood in PBS- and careful overlay of diluted blood onto Ficoll-Paque prior to centrifugation can be employed to achieve a slightly higher level of neutrophil purity by more efficiently separating the buffy coat from the granulocytes⁸. Omitting this step serves to save time without drastically impacting yield or purity for the purposes of this assay, and thus we favor not including the blood overlay onto Ficoll-Paque step.
8. While blood is spinning, very slowly add 1 g gelatin into 50 ml heated Hanks’ Balanced Salt Solution without calcium and magnesium (HBSS(-)) to prepare 2% gelatin solution. Keep solution at 37 ºC.
9. Aspirate and discard plasma from the 50 ml tube of blood using a glass pipette tip that has been treated with Sigmacote.
10. Continue aspirating very carefully to remove the buffy coat, which contains white blood cells, including lymphocytes.
11. As the buffy coat is slowly being removed, the whitish yellow cloudy material above the red blood cell (RBC) layer will disappear. Cease aspiration once buffy coat is removed and RBC layer can be clearly visualized when viewing from above the tube.
12. Try not to disturb the RBC layer as granulocytes, mostly neutrophils, are near the surface of the RBC layer, but are not visible.
13. To a volume of 15 ml RBC layer, add 30 ml warm 2% gelatin solution (2x volume of the RBC layer).
14. Invert tube gently to mix, taking care to minimize formation of bubbles.
15. Incubate at 37 ºC for 25 min. Note: RBCs should aggregate and settle to the bottom. As an alternative to gelatin, large molecular weight dextrans can be employed to sediment RBCs⁸.
16. After incubation, transfer top layer light reddish solution that contains granulocytes into a new 50 ml tube with pipette, being careful not to disturb the settled dark RBC layer during transfer. Once separated, discard dark RBC layer.
17. Spin transferred light reddish solution containing granulocytes and some RBCs in centrifuge at 1,000 x g with brake activated for 10 min at room temperature in order to pellet cells.
18. Pour off or aspirate the supernatant carefully, as the reddish cell pellet that will form following centrifugation is generally loose.
19. Prepare RBC lysis buffer in advance and store at 4 ºC by adding 8.29 g ammonium chloride (NH₄Cl), 1 g sodium bicarbonate (NaHCO₃), and 0.038 g EDTA to 1 L distilled water in a beaker. Store RBC lysis buffer at 4 ºC.
20. Resuspend and break up reddish cell pellet with a small volume of cold RBC lysis buffer. Once the cell pellet is in solution, fill tube to 50 ml with RBC lysis buffer. Note: At this point and going forward, keep the cells on ice or at 4 ºC.
21. Spin in centrifuge at 300 x g for 10 min at 4 ºC and then pour off or aspirate the supernatant. Note: The supernatant will be somewhat reddish containing lysed RBCs. The pellet should be yellowish white at this point. If significant RBCs remain in the pellet as indicated by a reddish pellet, step 5.20-5.21 can be repeated. Significant RBCs remain in the cell pellet if the reddish pellet from step 5.20 is not broken up sufficiently.
22. Resuspend and break up whitish pellet with a small volume of HBSS(-) and then fill tube to 50 ml with HBSS(-). Note: HBSS(-) and not HBSS+ should be used to resuspend the whitish cell pellet. HBSS(-) lacks calcium and magnesium.
23. Spin again in centrifuge at 300 x g for 10 min at 4 ºC.
24. Pour off supernatant and resuspend the pellet to a final volume of 1 ml HBSS(-).
25. Add 5 µl of the 1 ml cell suspension to 250 µl HBSS(-) in a 1.5 ml Eppendorf tube and mix gently up and down by pipette.
26. Add 25 µl diluted cell suspension to 25 µl 0.4% Trypan blue.
27. Add 12 µl of mixture to each side of a standard hemocytometer.
28. Count the number of living cells that have excluded the Trypan blue dye and are present in the middle square of both sides of the hemocytometer.
29. To calculate the cell concentration, multiply the average of the 2 squares by 100 (dilutional factor) and then multiply by 10⁴ to give the number of cells/ml.
30. Adjust the final concentration to 5 x 10⁷ cells/ml. If greater than 5 x 10⁷ cells/ml, adjust by adding HBSS(-). If less, pellet the cells and resuspend to appropriate volume.
31. Keep cells on ice until ready for migration assay. Note: These cells represent the granulocyte population of white blood cells. The majority of the granulocytes isolated from whole human blood are neutrophils. Cells are kept on ice suspended in HBSS(-) until addition to the migration assay in order to maintain cells in a quiescent state.

6. Preparation of Epithelial Cell Layers for Migration Assay

(These steps do not need to be performed within a sterile hood.)

1. Remove from incubator 24-well plate supporting epithelial layers that have been cultured on the underside of transwell filter membranes for at least eight days.
2. Grasp the edge of each transwell with a hemostat, lift out of the 24-well plate and invert the transwell over a waste bucket to discard culture media from the inner chamber of the transwell. Dip each transwell into a wash beaker containing HBSS+ to fill the inner chamber of the transwell with HBSS+. Discard fluid from inner chamber into a waste bucket.
3. Repeat step 6.2 for a second wash in HBSS+. After two washes, place transwells into a new 24-well plate containing 1 ml HBSS+ in each well. Note: All washes should be performed with HBSS+ that is warmed to 37 ºC.
4. Observe the top chamber of each transwell to assess epithelial cell layer integrity. Note: HBSS+ should not leak into and fill the top chamber of the transwell when placed in the well of a 24-well plate containing 1 ml of HBSS+.
5. After carefully observing each transwell, add 0.2 ml of HBSS+ into top chamber.
6. Place in incubator at 37 ºC, 5% CO₂ in a humidified chamber for 30-60 min to equilibrate epithelial cell layers.

7. Neutrophil Trans-epithelial Migration Assay

1. Remove from incubator 24-well plate of washed transwells equilibrating in HBSS+.
2. Grasp each transwell with a hemostat and discard fluid in top well into a waste bucket.
3. Place each transwell upside down in a 150 mm x 25 mm Petri dish.
4. To six of the inverted Transwells, add 25 µl of HBSS+. To three of the inverted transwells, infect with 25 µl P. aeruginosa (PAO1) diluted in HBSS+ prepared as described in step 4. To the final three inverted transwells, infect with 25 µl E. coli K12 (MC1000) diluted in HBSS+ prepared as described in step 4. This is approximately 0.8 x 10⁶-1.5 x 10⁶ CFUs/epithelial layer for each bacterial species since the concentration of the bacterial solutions prepared in step 4 is approximately 3 x 10⁷-6 x 10⁷ CFU/ml.
5. Incubate at 37 ºC, 5% CO₂ in a humidified chamber for 60 min.
6. Prepare fMLP as a positive control for neutrophil migration by adding 5 µl of 10 mM fMLP stock to 5 ml HBSS+ giving a 10 µM solution.
7. Wash transwells by grasping with a hemostat and flipping back over into a 24-well wash plate that contains 1 ml HBSS+ in the bottom chambers. Add 0.2 ml of HBSS+ to the top chambers.
8. Grasp transwell with a hemostat and remove from the first wash plate. Prior to placing transwells into a second 24-well wash plate containing 1 ml HBSS+, discard fluid from top chamber into a waste bucket. Add 0.2 ml of HBSS+ to the top chamber.
9. Repeat steps 7.8 one more time for a total of 3 washes. Note: All transwells should be subjected to the same handling and washing regimen, whether or not they have been infected with bacteria, to ensure that all transwells being assayed have been manipulated in the same manner. A 60 min infection with either PAO1 or MC1000 does not reduce viability or alter the barrier properties of the H292 monolayer as assessed by a lactate dehydrogenase based toxicology assay and the HRP flux assay, respectively^18,22.
10. Prepare 24-well migration plate by adding 1 ml HBSS+ into 12 wells of the 24-well plate. Note: This plate can be prepared in advance.
11. After the third wash, discard fluid from top chambers of transwells into a waste bucket using a hemostat and place three of the uninfected transwells into three wells of the 24-well migration plate containing 1 ml HBSS+, which represents the negative control.
12. Pipette 10 µl of a 10 µM solution of fMLP into each of three wells of the 24-well migration plate containing 1 ml HBSS+ (100 nM).
13. Place three of the uninfected transwells into three wells of the 24-well migration plate containing 100 nM fMLP, which represents a positive control for the ability of isolated neutrophils to migrate.
14. Place three transwells infected with PAO1 and three transwells infected with MC1000 into three wells each respectively of the 24-well migration plate containing 1 ml HBSS+. Add 0.1 ml of HBSS+ to the top chamber of every transwell.
15. On top of the 0.1 ml HBSS+ in each transwell, carefully add 20 µl of the neutrophil suspension (5 x 10⁷ cells/ml) prepared as described in step 5. Note: This represents approximately 1 x 10⁶ neutrophils/transwell.
16. Incubate at 37 ºC, 5% CO₂ in a humidified chamber for 2 hr to allow trans-epithelial migration of neutrophils.
17. Prepare a standard curve for determining the number of neutrophils that have migrated after 2 hr. Add 40 µl of the neutrophil suspension (5 x 10⁷ cells/ml) into 2 ml of HBSS(+) and transfer 1 ml into 1 ml HBSS(+) and perform serial 2-fold dilutions eight additional times for a total of 10 standards ranging from approximately 2,000 cell/ml to 1 x 10⁶ cells/ml. Include 1 ml HBSS(+) for blank.
18. After 2 hr of migration, lift the transwells by grasping with a hemostat and tap gently against the inside walls of the 24-well migration plate. Discard transwells.
19. Assess the migration of neutrophils grossly in the wells by viewing wells of the 24-well migration plate under an inverted microscope using the 10X objective (See Figure 1 for images of migrated neutrophils in each condition).
20. Add 50 µl of 10% triton X-100 to each well used in the 24-well migration plate, each standard, and the blank.
21. Rotate the 24-well migration plate, standards, and blank at low speed for 20 min at 4 ºC.
22. Prepare 1 M citrate buffer in advance. Add 52.5 g citric acid and 73.5 g sodium citrate to 400 ml distilled water in a beaker. Bring pH to 4.2. Add distilled water for final solution of 500 ml. Store solution at 4 ºC.
23. Prepare ABTS substrate solution fresh that day. Add 3 ABTS tablets (10 mg each) and 5 ml of 1 M citrate buffer to 45 ml distilled water. Allow tablets to fully dissolve in solution. Withhold addition of 50 µl of 30% hydrogen peroxide until the ABTS substrate solution is needed (see step 7.27).
24. Add 50 µl of citrate buffer to each well used in the 24-well migration plate, each standard, and the blank.
25. Transfer 100 µl of each sample well in duplicate to a 96-well plate. Note: It is very important to thoroughly mix contents in each sample well prior to moving to the 96-well plate. Pipette the solution up and down at least five times prior to transfer.
26. Transfer 100 µl of each standard and the blank to the 96-well plate after mixing (blank in duplicate).
27. Add 50 µl of 30% hydrogen peroxide to the ABTS solution and vortex.
28. Add 100 µl of ABTS substrate solution to each sample on the 96-well plate.
29. Allow plate to develop for 5-10 min in the dark. Note: A green color should develop in certain wells, correlating with the number of neutrophils present in each well.
30. Read at wavelength of 405 nm using a microplate reader.
31. Calculate the number of neutrophils that have migrated across the epithelial layer using the standards whereby a linear positive correlation exists between the known numbers of neutrophils prepared as standards and the amount of peroxidase activity as measured by optical density at 405 nm (See Figures 2 and 3 for representative data). Note: This method of neutrophil quantification takes advantage of the large amount of peroxidase activity exhibited by neutrophils due to myeloperoxidase expression. Alternative approaches to quantify neutrophils can be considered, such as methods involving prelabeling neutrophils with radioactivity or fluorescence compounds.