Live or Dead™ Cell Viability Assay

Protocol summary

1. Prepare cells with test compounds 
2. Add the same volume of CytoCalcein™ Green/Propidium Iodide dye-working solution (100 µL/well/ 96-well plate or 25 µL/well/384-well plate)
3. Incubate at room temperature or 37°C for 1 hour
4. Monitor fluorescence at intensity (bottom read mode) Ex/Em = 490/525 nm (Cutoff = 515 nm, live) and 540/620 nm (Cutoff = 590 nm, dead), fluorescence microscope with FITC filter (live) and TRITC filter (dead), or flow cytometer with FL1 and FL2 channels

Important notes
Thaw one of each kit component at room temperature before starting the experiment.

1. CytoCalcein™ Green stock solution:
Add 20 µL of DMSO (Component C) into the vial of CytoCalcein™ Green (Component A) and mix well to make CytoCalcein™ Green stock solution. Protect from light. Note: 20 µL of CytoCalcein™ Green stock solution is enough for one plate. For storage, seal tubes tightly.

Add the whole content (20 µL) of CytoCalcein™ Green stock solution and 20 µL Propidium Iodide (Component B) into 10 mL of Assay Buffer (Component C) and mix well to make CytoCalcein™ Green/Propidium Iodide dye-working solution. The CytoCalcein™ Green/Propidium Iodide dye-working solution is stable for at least 2 hours at room temperature. Note: If the cells such as CHO cells contain organic-anion transporters which cause the leakage of the fluorescent dye over time, a probenecid stock solution should be prepared and added to the loading buffer at a final in-well working concentration ranging from 1 to 2.5 mM. Unused probenecid stock solution can be stored at ≤ -20 ^oC. As the optimal staining conditions may vary depending on different cell types, it’s recommended to determine the appropriate concentration of Component A and B individually.

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

Run the cell viability assay with plate reader or fluorescence microscope:

1. Treat cells with test compounds as desired. Note: It is not necessary to wash cells before adding compound. However, if tested compounds are serum sensitive, growth medium and serum factors can be aspirated away before adding compounds. Add 100 µL/well/96-well plate and 25 µL/well/384-well plate of 1X Hank’s salt solution and 20 mM Hepes buffer (HHBS) or the buffer of your choice after aspiration. Alternatively, cells can be grown in serum-free media.
   
   
2. Add 100 µL/well (96-well plate) or 25 µL/well (384-well plate) of CytoCalcein™ Green/Propidium Iodide dye-working solution.
   
   
3. Incubate the plate at room temperature or 37°C for 30 minutes to 1 hour, protected from light. (The incubation time could be from 15 minutes to overnight. We got the optimal results with the incubation time less than 4 hours). Note: The appropriate incubation time depends on the individual cell type and cell concentration used. Optimize the incubation time for each experiment. DO NOT wash the cells after loading. For non-adherent cells, it is recommended to centrifuge cell plates at 800 rpm for 2 minutes with brake off after incubation.
   
   
4. Monitor the fluorescence intensity with a fluorescence plate reader (bottom read mode) at Ex/Em = 490/525 nm (Cutoff = 515 nm, live) and Ex/Em = 540/620 nm (Cutoff = 590 nm, dead) or fluorescence microscope with FITC filter for live cells or TRITC filter for dead cells.

Run the cell viability assay with a flow cytometer:

1. Treat cells with test compounds for a desired period of time.
   
   
2. Centrifuge the cells to get 1 - 5 × 10⁵ cells/tube.
   
   
3. Resuspend cells in 500 µL of CytoCalcein™ Green/Propidium Iodide dye-working solution.
   
   
4. Incubate at room temperature or 37°C for 10 to 30 minutes, protected from light.
   
   
5. Optional: Wash the cells with HHBS or buffer of your choice. Resuspend cells in 500 µL of HHBS to get 1 - 5 × 10⁵ cells per tube.
   
   
6. Monitor the fluorescence intensity with flow cytometer at Ex/Em = 490/525 nm and  Ex/Em = 490/620 nm (FL1 and FL2 channels).