Hey everybody. I'm hoping someone out there can help me out with a question I have. I am working with a pore forming toxin, and am looking for ways to fluorescently stain cells that have been perforated. The pores are too small to allow the passage of antibodies, so I am looking to use different small molecules.
My protocol requires a fixation step. I am looking to expose cells to the pore forming molecule, fix the cells at different time points, then stain them with a fluorescent molecule. Fluorescence would indicate pore formation, no fluorescence would indicate that pores have not yet been formed, or that they were formed but the cells repaired their membranes prior to treatment with the fluorescent probe.
I first tried the assay with EthD-1 but found that all of the control cells that were not treated with the pore forming agent, were still stained with EthD-1. I know that EthD-1 stains dead cells, but I cannot find anymore information regarding the mechanism by which it does so. My intial assumption was that a dead cell would have a compromised membrane which EthD-1 would be able to pass through, but that my untreated cells fixed with PFA, while they are dead, would not have compromised membranes. They were not treated with anything in order to kill them before the PFA treatment, so I figured their intact membranes would be fixed in place. I've performed plenty of other staining protocols using PFA as a fixative agent and have never had issues with it disrupting cell membranes and allowing probes to pass through. Can someone out there tell me exactly how EthD-1 selectively stains dead cells, and why it's use is not compatible with a fixation step? Also, would I have this same problem if I use propidium iodide instead of EthD-1 to stain nucleic acids?
EthD-1 live/cead cell staining
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