I'm wondering if anyone has stumbled on a similar phenomenon, as I? Or if you may have an explanation of it.
I was using ethidium and calcein-AM (LiveDead assay) on retinablastoma cells (Y79). Before the LiveDead assays, the cells had been made to adhere to coverslips, with use of ornithin and fibronectin. They had also been treated with a cytostatic agent or with DMSO.
I found that live cells were mainly present at the periphery of the coverslips, while dead cells were found more centrally (see pictures). At first, I thought the phenomenon was probably caused by my method of staining. I was adding the ethidium and calcein-AM into the 24-well-plate, where my coveslips were. Due to surface tension, the LiveDead solution was being pulled toward the walls of the well, making the solution unequally distributed. I tried switching my method of staining. I moved my coverslips from the 24-well-plates, onto water-repellent parafilm were I made my new assays. I placed 60 uL of the staining solution on top of each coverslip, and let the solution remain of top by its surface tension. However, the phenomenon remained.
What is this phenomenon? Is it avoidable?
Hi Karl, I'm guessing that you are using 10% FBS serum in all of your experiments.
The live cells are quickly adhering to a surface coated with serum (mostly through vitronectin or fibronectin) and staying there. The dead cells can't adhere so they are just moving with the flow towards the inside.
I would predict that if you used serum-free media in your experiments and plating, the demarcation would not be so extreme.
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