Cell Navigator® F-Actin Labeling Kit *Red Fluorescence*

Protocol summary

1. Prepare samples (microplate wells)
2. Remove the liquid from the plate
3. Add 100 µL/well of iFluor™ 594-Phalloidin working solution
4. Stain the cells at RT for 15 to 60 minutes
5. Wash the cells
6. Examine the specimen under fluorescence microscope at Ex/Em = 594/610 nm (Texas Red channel)

Important notes
Thaw all the components at room temperature before starting the experiment.

Add 10 μL of iFluor™ 594-Phalloidin (Component A) to 10 mL of Labeling Buffer (Component B) to make 1X iFluor™ 594-Phalloidin working solution. Protect from light. Note: Different cell types might be stained differently. The concentration of iFluor™ 594-Phalloidin working solution should be prepared accordingly.

For guidelines on cell sample preparation, please visit
https://www.aatbio.com/resources/guides/cell-sample-preparation.html

1. Perform formaldehyde fixation. Incubate the cells with 3.0 – 4.0% formaldehyde in PBS at room temperature for 10 – 30 minutes. Note: Avoid any methanol containing fixatives since methanol can disrupt actin during the fixation process. The preferred fixative is methanol-free formaldehyde.
   
   
2. Rinse the fixed cells 2 – 3 times in PBS.
   
   
3. Optional: Add 0.1% Triton X-100 in PBS into fixed cells for 3 to 5 minutes to increase permeability. Rinse the cells 2 – 3 times in PBS.
   
   
4. Add 100 µL/well (96-well plate) of iFluor™ 594-Phalloidin working solution into the fixed cells.
   
   
5. Stain the cells at room temperature for 15 to 60 minutes.
   
   
6. Rinse cells gently with PBS 2 to 3 times to remove excess dye before plate sealing.
   
   
7. Image cells using a fluorescence microscope with Texas Red channel (Ex/Em = 594/610 nm).