I am trying to evaluate the cytotoxicity of my effector cells against an adherent target cell line. I plated 10,000 target cells and incubated them overnight to allow them to attach. I then stained them with 15uM calcein AM in complete medium and incubated the cells for 45 min at 37C. I washed three times with complete medium, then added my effector cells at a 1:1 ratio. I had a negative control (target cells alone) and a positive control (target cells with lysis buffer). I had two separate plates - after 4 and 24 hours, I transferred the supernatant to a black opaque 96 well plate, and added lysis buffer to lyse any effector cells that might've uptaken calcein. I have calcein AM blue, so read at 360/449 nm. When I read the signal, I saw very high background signal in my negative control wells, and did not see much difference between my experimental wells and positive control. Could this be due to insufficient washes? Or is there something else contributing to such high noise?
Why not consider using fluorescence microscopy to observe what's happening with your cells? You can also include specific death staining, such as EtBr for the red channel, or a variety of death reagents available on the market. This approach would allow you to quantify changes relatively easily in the images. For the plate reader approach, I’d recommend switching the wavelength to match Calcein Green AM 488/535 nm. Additionally, have you tested for potential autofluorescence in your media or plate? This can sometimes interfere with imaging and quantification, so confirming it could help clarify your results.
I don't have a clue about your case.
Did you collect only the supernatant for reading the signal? Another important factor is to be very careful during the plate handling and centrifugation in order to avoid spurious cell death.
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