I've been piloting clonogenic survival assays to look at long-term effects of of drugs in colon cancer cell lines (SW620, SW480, HT-29). My initial pilot assays in 12-well plates were successful and was able to establish optimal seeding conditions (200 cells/well for SW620/HT-29, 50 cells/well SW480) and assay endpoint (10 days for all 3 cell lines). However, my student who is running these assays has run into some issues.

We run a lot of control (vehicle) treated wells (6 wells/plate) to account for plating efficiency but in some cases, we get very few colonies back. And when there are colonies, a large proportion of colonies are stuck to the edge of the well, which makes counting single colonies problematic

I'm pretty sure this is down to my student's plating technique as the assays work ok in my hands but I'm wondering if anybody has any advice on this/improvements to the method.

PROTOCOL:

Cells are counted via haemocytometer and then diluted 1:100 to allow for cell suspensions to be made. Cells are plated at 200 cells/ml (SW620/HT-29) or 50 cells/ml (SW480) per well of a 12-well plate (we use 3 technical replicates per run). Cells are left overnight to settle. Next day, media is aspirated and replaced with 500 ul media containing drug/vehicle and treated for 24h. Drug media is the replaced with 1 ml full culture media and colonies are left to develop for 10 days. Colonies are then washed in 50% methanol/PBS, followed by fixing in 100% methanol and staining with crystal violet (1:40 in PBS) for 45 minutes. Excess stain is washed off and colonies counted manually or in imageJ.