I have been trying to detect mineralisation in human osteoblasts and the hFOB cell line for some time now, but have been unsuccessful. My old method was to seed out 20,000 cells/cm2 and leave them for 24 hours to attach, before adding osteogenic media (50uM ascorbate-2-phosphate, 10uM B(beta)-glycerophosphate and 0.1uM dexamethasone). Cells were then fed twice weekly and left for 21 or 28 days before being washed with PBS and fixed in 10% PFA for 30 minutes. Cells were then stained with 40mM alizarin red (pH 4.2), for 40 mins in the dark at room temperature. Finally, monolayers were washed with distilled water three times and left to dry.
I think I made one obvious mistake - from what I have read the cells should be at confluency before adding osteogenic media and will likely have to be left longer in future.
I have a few questions:
Is it OK to wash cells with PBS? Or should I use Tris and try and eliminate Ca and Mg ions?
Is 10% PFA too much and should I instead be using neutral buffered formalin at a lower percentage?
I have been culturing cells in 96 well plates (to limit use of extracts), but many protocols mention 24 wells instead. Is a 24 well plate a better option?
Anything else I might be doing wrong would be a great help, as this lack of mineralisation is really holding up work.
Thanks so much in advance for any help,
Matt
This may have been a typo on your part but you said 10% PFA, what you should be using is either 4% PFA or 10% neutral buffered formalin. Attached is a protocol I was just looking at that may help you.
Alizarin Red staining for osteogenesis:
1) Preparation of Alizarin Red solution (2%):
a. 1 gr dissolve in 50 ml water
b. Then filter and adjust pH (4.1 – 4.3)
c. Adjustment must be performed by ammonium hydroxide 0.5% / HCL
d. Keep the solution at 4.c
2) Staining protocol
a. Cells wash with PBS
b. Fixation with 10% formalin (5 min)
c. Wash with dH2O for 2 times
d. Stain with alizarin red 2% (5 min)
e. Wash with dH2O for 2 times
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