Overview

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1. Prepare cells:

1.1    For adherent cells: Plate cells overnight in growth medium at 10,000 to 40,000 cells/well/100 µL for 96-well plates or 2,500 to 10,000 cells/well/25 µL for 384-well plates.

1.2    For non-adherent cells: Centrifuge the cells from the culture medium and then suspend the cell pellets in culture medium at 50,000-100,000 cells/well/100 µL for 96-well poly-D lysine plates or 10,000-25,000 cells/well/25 µL for 384-well poly-D lysine plates. Centrifuge the plates at 800 rpm for 2 minutes with brake off prior to the experiment.

Note: Each cell line should be evaluated on an individual basis to determine the optimal cell density.

2. Prepare Calcein UltraBlue™ stain solution:

2.1    Prepare Calcein UltraBlue™ stock solution: Add 20 µL of DMSO into Calcein UltraBlue™ vial (Component A) and mix them well.

Note: 10 µL of Calcein UltraBlue™ stock solution is enough for one plate. Unused Calcein UltraBlue™ stock solution can be aliquoted and stored at < -20 ^oC for more than 2 weeks if the tubes are sealed tightly. Avoid repeated freeze-thaw cycles and protect from light.

2.2    Prepare Calcein UltraBlue™ working solution for one cell plate: Add 10 µL of DMSO reconstituted Calcein UltraBlue™ stock solution (from Step 2.1) into 10 mL of HHBS (Component B), and mix them well.

3. Stain the cells:

3.1    Remove the growth medium from the cell plates.

3.2    Add 100 µL/well (96-well plate) or 25 µL/well (384-well plate) Calcein UltraBlue™ working solution (from Step 2.2) into the cell plate.

3.3    Incubate the cells in a 37 ^oC, 5% CO₂ incubator for 30 minutes to 2 hours.

3.4    Wash cells with HHBS buffer (Component B), and add growth medium or HHBS back to the cells.

3.5    Image the cells using a fluorescence microscope with DAPI filters (Ex/Em = 360/445 nm).

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