Materials Required

1. Phosphate Buffered Saline (PBS):

   To 800 mL of distilled water, add:

   8 g of NaCl

   0.2 g of KCl

   1.44 g of Na₂HPO₄

   0.24 g of KH₂PO₄

Adjust the pH to 7.4 using HCl. Add distilled water to bring the final volume to 1 liter. Dispense the solution into aliquots and sterilize by autoclaving at 121°C for 20 minutes (liquid cycle). Store at room temperature.

2. 2% Paraformaldehyde

3. 0.2% Bovine Serum Albumin (BSA) and 0.75% Saponin

4. Fluorescent Phallotoxin

Procedure

1. Pellet 50,000 cells onto each coverslip placed inside a 12-well plate.
2. Incubate the cells for 24 hours at 37 °C in a carbon dioxide incubator.
3. Remove the plate and check for 90% confluency under a phase contrast microscope.
4. Perform the scratch assay.
5. Remove all media from the wells using a pipette tip attached to a vacuum pump.
6. Wash the wells twice with PBS.
7. Fix the cells with 500 µL of 2% paraformaldehyde supplemented with 1 mM magnesium chloride and 0.1 mM calcium chloride.
8. Incubate the plate for 30 minutes in the dark.
9. Wash the wells thoroughly three times with PBS.
10. Block with 0.2% BSA and 0.75% saponin and incubate in the dark for 15 minutes.
11. Incubate with PBA and phalloidin in a 1:200 ratio for 20 minutes. To prevent evaporation, place the well plate inside a covered container during incubation.
12. Wash three times with PBS.
13. Carefully remove the coverslips from the wells using forceps. Invert and mount them onto glass slides. Take care to avoid breakage.
14. Fix the coverslip onto the slide using nail polish. Gently press the center of the coverslip with forceps and apply the nail polish around the edges, ensuring it touches both the coverslip and the slide.
15. View the samples under a fluorescence microscope at 20X magnification.