I have to coat my 12, 24 and 96 well plates with collagen and I`m not so sure if this is working very well and equally on the whole plate. Is there some kind of staining or other method to visualize that coating?
You can measure the amount of protein that was adsorbed to the tissue culture plastic using a BCA assay. Calculate the difference in total protein in solution pre- vs. post-adsorption.
Use enough (not too much) volume just to cover the surfaces of your wells, to allow even distribution.
I am not familiar with visualizing coatings, it might inhibit cell attachment and growth. For a simple test, you might stain the odd well with e.g. eosin, Safranine O, Sirius red or ponceau S?
Maybe this paper is of use:
Article Simple and High Yielding Method for Preparing Tissue Specifi...
Try staining your plates after collagen coating with species-specific antibody to type-I collagen. Use a brightfield label (AEC or DAB) to visualize binding. It will show location as well as density of coating.
When we coated our plates and flasks with type-I collagen we noted that a swirling action would deposit the collagen along the periphery and leave the center of the plates and flasks collage-free. We determined that moving the plates/flasks back-and-forth (5x), then side-to-side (5x), then back-and-forth (5x), and then side-to-side (5x), followed by 30 min incubation at rest before removing excess solution gave us the best and most consistent coating as assessed by type-I antibody staining.
You could mix some phenol red in with your collagen solution (the dye indicator in most media). I attached a collagen recipe that I got from the Turkish Cell Culture Legend Dr. Altintas, it works like a charm.
Vincent -
Can you explain how the phenol red helps to monitor/prove collagen 1 deposition? I could imagine that if the soluble collagen is acting to buffer the coating medium, then the pH might change as collagen deposits on the TC surface. Is this what you mean?
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