I am also trying to stain cultured cells (osteoblasts and adipocytes) and am not having any luck. I have no experience with staining and no physical guidance and am just working from protocols I find online.
I have tried Alizarin Red S staining and failed miserably. Has anyone had any luck staining using this protocol or a similar protocol:
Cells were grown in a Nunc 24 well plate.
Add 500ul freshly prepared 4% paraformaldehyde/DPBS to each well. Incubate 5 minutes at room temperature on a Nutator.
Aspirate paraformaldehyde and wash the cells 3 times with 1mL 1xDPBS. Aspirate DPBS.
Stain cells with 750ul per well 2% Alizarin Red S solution (pH 4.2) for 2-3 minutes on a Nutator (i also tried staining for 2-3 hours with no luck). Rinse the wells 3 times with distilled water.
Visualize under light microscope and capture images for qualitative or quantitative purposes. Calcium salts appear intense reddish orange.
2% Alizarin Red S Solution: 2g Alizarin Red S, C.I. 58005, 100mL Distilled water
Stir dye into the distilled water so that only a few small grains of dye remain undissolved. Adjust pH to 4.2 with ammonium hydroxide. Solution is stable for one year.
4% paraformaldehyde: 0.4g paraformaldehyde powder, 10mL 1xDPBS, 25ul 5M NaOH. Heat to 65oC until dissolved, approximately 5 minutes, then cool to room temperature before use.
Is there something wrong with this protocol?
Also, I want to try to stain cultured osteoblasts using von Kossa staining, but I'm having difficulty finding a protocol that is specific for cultured cells. Can I use a tissue protocol using cells grown in a 24 well plate? fixing the cells at described above, and then following this protocol:
Solutions and Reagents:
1% Aqueous Silver Nitrate Solution:
Silver nitrate ------------------------- 1 g
Distilled water ---------------------- 100 ml
5% Sodium Thiosulfate:
Sodium thiosulfate ---------------- 5 g
Distilled water -------------------- 100 ml
0.1% Nuclear Fast Red Solution:
Nuclear fast red ------------------- 0.1 g
Aluminum sulfate------------------ 5 g
Distilled water ----------------------100 ml
Dissolve aluminum sulfate in water. Add nuclear fast red and slowly heat to boil and cool. Filter and add a grain of thymol as a preservative.
3. Incubate cells with 1% silver nitrate solution. place under ultraviolet light for 20 minutes (or in front of a 60-100 watt light bulb for 1 hour or longer). Note: If stain was weak or rinsed off in washing steps, it indicated the UV light was not strong enough. Longer staining is required for up to several hours.
4. Rinse in several changes of distilled water.
5. Remove un-reacted silver with 5% sodium thiosulfate for 5 minutes.
6. Rinse in distilled water.
7. Counterstain with nuclear fast red for 5 minutes.
8. Rinse in distilled water.
9. Dehydrate through graded alcohol and clear in xylene.
Has anyone actually used von Kossa staining on cultured cells in a plate (not on glass)? Any tips?
Lastly, I want to stain cultured adipocytes using Oil Red O. After fixing as described above, I want to use the following protocol. Has anyone used a protocol like this, did it work, and do you have any suggestions?
Oil Red O stock
FW 408.5, Sigma O-0625
0.7 g Oil Red O
200 ml Isopropanol
Stir O/N, then filter with 0.2 μm and store at +4°C
Oil Red O Working Solution
6 parts Oil Red O stock
4 parts dH2O
Mix and let sit at room temp for 20 min
Filter 0.2 μm
10% Formalin in PBS
Wash wells with 60% isopropanol.
Let the wells dry completely
Add Oil Red O working solution for 10 min (do not touch walls of the wells)
Remove all Oil Red O and IMMEDIATELY add dH2O, wash with H2O 4 times (you can wash under running tap water)
Take pictures if desired
I appreciate your time and any help you can offer.