Stainning of peritoneal macrophages with Oil Red O - I'm not really sure of HOW to prepare the Oil Red O solution (Jul/22/2010 )

Hello everybody,

I am trying to stain peritoneal macrophages of mouse with Oil Red O. I did it once with an Oil Red O prepared 0,2% in Methanol 100% and NaOH 1M (0,07g Oil Red O, 25ml Mehanol, 10ml NaOH 1M), and everything was fine. The problem is now, every time I try to repeat that, with Oil Red prepared in the same way and with the same stainning protocol, the result of the stainning is awful, the Oil Red seems to form crystals inside the cells and the stainning seems to be unspecific. I filtered everytime I prepare it, and again before use it through 0,45 filter, so it is not the problem.

I would be very grateful if someone who usually stains macrophages with Oil Red to see foam cell formation, acLDL uptake or similar, could tell me his protocol to prepare the Oil Red. I use the Oil Red O O0625 of Sigma Aldrich.

I'm really desperate.

Thanks in advance for your help!

Gema.

GMA on Jul 22 2010, 04:47 PM said:

Hey Gema,

I have been staining THP-1 differentiated foam cells with ORO-staining, and it has worked quite well. Here it goes like this,

Materials:
• Cells
• 1xPBS
• 10% Formalin, neutral buffered
• Millipore water
• 60% Isopropyl alcohol (2-propanol)
• Mayer’s Hemalin
• Whatman filter paper
• Tap water

Protocol for preparing Oil Red O stain:
1. Weight 300mg of Oil Red O red and add it to 100ml of 99% isopropanol. This stock solution is stable for a year from this date onwards.
2. Mix 3 parts of ORO-Stock with 2 parts of Millipore water. Incubate in RT for 10min.
3. Filter ORO working solution completely through Whatman paper on funnel on proper vessel. This working solution is stable for 2 hours.

Protocol for staining cultured cells:

1. Remove cell cultures from incubator and transfer them to fume hood.
2. Transfer cover slips, which have adhered cells on them to wells containing 2ml 1xPBS. (If removing media, controls always first!!)
3. Remove 1xPBS and add 2ml 10% Formalin. Incubate 60min in fume hood.
4. After incubation remove formalin to waste container (preferably in fume hood.)
5. Gently rinse cells with 2ml Millipore water. Remove and discard.
6. Wash twice with 1ml 1xPBS.
7. Add 2ml 60% Isopropanol and let it sit for 5min.
8. Remove Isopropanol and add 2ml Oil Red O Working Solution per well. Be sure the coverslips are immersed in liquid. Incubate 5min RT.
9. Remove Oil Red O and rinse under RT tap water carefully with low pressure water flow until water rinses of clear.
10. Add 1ml of Mayers Hemalin. Let it sit 5 min. Rinse with tap water until rinses of clear
11. Mount cover slips on objective glasses cell downwards on drop of ProLong Antifade Gold (Aqua-based) to preserve fluorescence.

The amounts of solutions used to stain cells vary according the volume of plates used for staining (24-well,500µl;12-well,1000µl etc.), so that the cells are covered properly. Don't leave cells to dry more than 30 secs. Also avoid pipetting of the solution straight on the cells.
The tap water rinsing is quite harsh, though cells are usually quite rigid after formalin fixation, one can also use some rinsing bottle etc..

hope this helps. It worked quite nicely. The protocol is adapted from some Thermo-Scientific protocol.

Keep it Jolly,

tero

Great! thanks a lot! I'll try it! Although I have never used cover slips. I usually work with the macrophages adhered to the wells of the culture plate, but I suppose that you work with the cover slips because of the THP-1 cell line.

Thanks again for your help Tero!

Gema.

hey buddy plz i willbe very oblzed if u can tell me wat is ds mayers hemalin and wat is its function? bcz there is unavailability of mayers heamlin in our lab so if i can knw the function ill use some of its analogue available in our lab.

                                                                     

thanks