Hi all,
I'm completely new to live cell imaging and have been trying to optimise a protocol to visualise cell contractile responses using calcein, however I've been having some issues with these orb-like artifacts appearing (image attached). I was wondering if anyone with more experience using calcein, or live cell imaging in general, knows what these could be and how I can prevent them from appearing?
I incubate my cells (retinal endothelial) in 1µM calcein dissolved in HEPES buffered HBSS for an hour, followed by washing. The cells are then imaged in HEPES buffered HBSS, at 37°C with 5% carbon dioxide.
The orbs definitely aren't caused by any treatment I'm using, as it also occurs in my controls.
Thank you.
I saw these dots in some other images in literature depending upon cell line. May be these might be due to some exocytosis secreted vesicles which might also be carrying esterases from cytoplasm that convert Calcein AM to green fluorescent. If you want to confirm, take another kind of cell line, these dots may be less in number or more in number depending on cell line. This is not contamination in my opinion
Hi, I agree with Dr. Juneja - many endothelial cells / cell lines does that. It is the extend that varies according to the well-being of your cells.
I think the problem is when you use HEPES and CO2. I have not measured the pH of the supernatant, but I know that my cells die in less than 24 h if I use HEPES for an experiment outside the incubator and put them back into 5% CO2.
So use HEPES or media with Hanks salts when outside 5% CO2 only. Otherwise use Earls salts or other buffer system intended for 5% CO2.
In regard to the calcein, I think 15 min incubation should be enough, and I use only 0,25 µM calcein in PBS / media and it works fine.
For live cell I would recommend complete media, preferably without phenol-red.
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