Live or Dead™ Cell Viability Assay Kit *Red/Blue Dual Fluorescence*

1. Prepare cells with test compounds
2. Add dye-working solution
3. Incubate at room temperature or 37°C for 30 minutes to 1 hour
4. Monitor fluorescence intensity (bottom read mode) at Ex/Em = 610/650 nm (Cutoff = 630 nm, Red) and Ex/Em = 360/450 nm (Cutoff =420 nm, Blue) or fluorescence microscope with Cy5 channel (live) and DAPI channel (dead)

Thaw all the kit components at room temperature before starting the experiment.

For guidelines on cell sample preparation, please visit:

https://www.aatbio.com/resources/guides/cell-sample-preparation.html

Add 5 µL of 200X Cellbrite™ Red (Component A) and 5 µL of 200X Nuclear Blue™ DCS1 (Component C) into 1 mL of Assay Buffer (Component B) and mix well to make dye-working solution. This dye-working solution is stable for at least 1 hour at room temperature.

Note: As the optimal staining conditions may vary depending on different cell types, it’s recommended to determine the appropriate concentration of Component A and C individually.

1. Prepare cells according to the standard protocol. Note: We treated HeLa cells with staurosporine (SS) for 4 hours at 37ºC to induce cell apoptosis. See Figure 1 for details.
2. Replace growth medium with 100 µL/well (96-well plate) or 25 µL/well (384-well plate) of dye-working solution.
3. Incubate the dye-working solution plate at room temperature or 37°C for 30 minutes to 1 hour, protected from light.
4. Wash cells with HHBS, PBS or buffer of your choice twice.
5. Add 100 µL/well (96-well plate) or 25 µL/well (384-well plate) of Assay Buffer (Component B) into the cells.
6. Monitor the fluorescence signal under a fluorescence microscope with Texas Red or Cy5 filter for live cells, and DAPI filter for dead cells. The fluorescence intensity can also be analyzed with a fluorescence microplate reader (bottom read mode) at Ex/Em = 610/650 nm (Cutoff = 630 nm, Red) and Ex/Em = 360/450 nm (Cutoff =420 nm, Blue).