I aim to perform Halo assay on Adipocytes to detect DNA damage. The adipocytes are prepared in ibidi 8-chamber slide. They're attached to the bottom of the chamber and allowed to differentiate fully for about 12 days. So they contain a lot of lipid droplets.
I usually use common lysis buffer (1% lauryl-sarcosine, 2M NaCl, 1% Triton, 10 mM Tris, 100 mM EDTA) to lyze the cells. However I have troubles to prepare "clean" slides without lipid.
May I ask your advises how to prepare a better lysis solution in such a way that:
- Remove completely lipid droplets.
- Cellular DNA is kept intact, and should not be damaged by the lysis steps.
- Cells should not be detached away from the original position.
Using an organic solvent fixative would remove lipids without any need for permeabilization/lysis. See this page for some examples: http://www.ihcworld.com/_protocols/general_ICC/fixation.htm
Thank you for your advice.
I indeed tried Acetone couples of time. However the sample after acetone fixation still contains remnants of lipid.
In light of acetone fixation:
- Should I always use at -20 Celcius ?
- We cultured cells in idibi 8-chamber slide. Would acetone disturb the plastic materials in this kind of slide ? (I saw some kind of "cracks" and not sure if it's because of Acetone).
- Should the duration of Acetone fixation be longer or shorter than 10 minutes, or it doesn't matter at all? (given that I have about 10 000 fat cell in a chamber).
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