When I seeded 0.1 million cancer cells in a 60mm plate and treated it with 3ul of fixed concentrations of a drug, it gave 40-50% cell death. But when we seeded with 0.6 million cells in 100mm plates and treated with 10ul of the same fixed concentration of the drug, it gives only 2-5% cell death. (As we use 3ml media for 60mm plates and 10ml media for 100mm plates). What should I do to solve this problem?
0,6 million of cells in 100mm plates have a cell density higher than 0.1 million in 60 mm plates, usually the effect of drugs on cell cultures strongly depend on cell density, so to maintain the same condition with respect to the first experiment you should seed about 0.3 million of cell in 100 mm plates, I hope that this will be useful for you
The cancer cells must be actively dividing to observe an anti-proliferative effect of a (cancer) drug, if they become too confluent they become senescent/stop dividing and don't care what drug you are throwing at it. Try a 'colony formation assay' - very low density seeding over 7-14 days, quantification via colony counting or CellTiter-Glo (proliferation assay, like MTT or staining with crystal violet or sulforhodamine B) - ie start from 500 cells/well in a 6-well dish - depends on the cell line though....
Hi
Its depends on the doubling time of your cells, and as said by Richard, on the mechanism of action of you drug, and of the time of treatment
Try to verify the cell cycle distribution of your cells plated at these 2 densities, one or tow days after.
Best
I agree with Maria, your seeding density was too high on the 100 mm plate (it should be 0.28 million cells to be equal to the 60 mm plate). Did the cells on the 100 mm plate have the room to double in cell numbers? If they did not, the drug may not work, especially if it interferes with DNA replication.
Is your drug in 100% DMSO? Some drugs breakdown after a freeze/thaw in DMSO. I have had that problem several times. When I went back to the drug powder and made a fresh DMSO stock the activity was restored.
The duplication of the cells is the key to see effects. So you need surface area (28.26 cm2 for your 60mm plates and 78.5cm2 for your 100mm dishes). the doubling time also is another factor as Stephane said. Where they the same cells you were testing? If not, you need to allow for them to reach their doubling time. You can find some of the doubling times by Googling it. Some kind researchers have gone through the trouble. If not you have to figure out the doubling time as you might need a longer incubation in some cells compared to the others.
In addition to the previously made points, I'd like to add that going from a 50% to 5% reduction in cellular viability is quiet the change in activity - which connotes a possible solubility and/or stability issue. Is aqueous solubility a challenge with this drug? Is the drug stored as a solution in DMSO? Are multiple freeze/thaw cycles used? (With each freeze/thaw cycle, condensation enters DMSO solutions, and can adversely affect stability and solubility.) Such low volumes implies the stock solutions are highly concentrated relative to the treatment solution. The drug may be precipitating out of solution when added to the cells. What method are you using to assay for cellular viability? Is the increased number of cells used in the 100 mm dish still within the linear range of your detection method?
Thanks Maria, and all for your precious suggestions.
As Daniel asked, my drug is generally in-soluble in aqueous solution and even DMSO it takes 15-20 min to dissolve with a brief exposure of Sonication and each time I prepare a fresh stock and use. Generally, I use Trypan blue Exclusion assay to detect % cell death and the linearity is maintained when we use higher concentration of Drug.
Thanks once again
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