Oil red stain may be difficult to interpret when looking at adipogenic differentiated cells .

Prepare oil red O- 0.5g in 100% isopropanol.

Fix cells using 4% paraformaldehyde (4 degrees Celsius) or 10% phosphate buffered formalin (room temp).  After fixing for 15-30 min remove fixative and rinse with phosphate buffered saline.  Remove and allow to air dry.  Meantime, mix oil red o stock at 6:4 ratio with dH2O and let stand for 10 min.  Filter with coffee filter or other fast filter then use a 0.2 micron syringe filter to add oil red o to cells.  If the oil red o is not filtered properly you will have a lot of background staining.  Leave the oil red o for 15 min then remove and rinse well with dH20.  Leave the final dH2O rinse on the cells for microscopy and analysis.

Hello Sallam,

Oil red O staining is very easy and I am doing the same since years. A detailed protocol is provided by Jodi and that seems promising. My protocol is for readymade Oil Red O which I use from Sigma and that's quiet simple. After fixation and washing of cells as suggested above make a dilution of 1:2 (1 part Oil red O, 2 parts ddH2O) of oil red O in ddH20. Make it in a tube e.g. you can use 1ml Oil Red O and put 2ml water in it. Now incubate cells for 15 minutes in this solution. After 15 min, decant out stain and give 4 washings with water or PBS. Now cells are ready to observe under microscope and you can see intense red droplets with no background.

Hope it helps !

Thank you all for your help.

Hi Ajay Kumar, 

Do you use any specific mounting media for this. I have used similar Oil Red O solution from Sigma. But it doesn't  work.

Thanks

Hi people and thanks for your info!

I would like to ask how many colorations, wells, cells, or what else, you can do with the quantity defined by Jodi and Ajay?

Many thanks.

Hi

I would like to know if anybody has done the spectrophotometric determination of lipids in cultured cells using Oil Red O.

Hi Shaima,

The protocol is very simple and similar to one described above. The difference is that you use 96-well plate with the respective volumes of water, oil red stain and i-propanol. After the cells are stained and washed, finally add 100 µL of 2-propanol for 10 minutes in order to extract the stain from the cells and measure the absorbance at 540 nm. Use undifferentiated cells as a negative control and differentiated adipocytes or standart treatment as a positive one.

Best of luck,

Stoyan

@ A. Nasibah,

Sorry to respond late as I almost forgot this thread. Generally we fix the cells in Neutral Buffered Formalin for 30 minutes and then wash with PBS. Take care not to add any alcohol based solvent like ethanol or isopropanol at this point because that will dissolve the oil and only left will be empty pockets created after oil droplet extraction. It might give you a false impression that droplets are there and they are not staining, while they are just empty pockets. I think in your protocol, you lost the droplets dear. Try the method again using same fixative. It should work.

@ Antonio Cervadoro ,

Generally you can use 100ul stain for 96 well, 300ul for 24 well and 500ul for 12-well. You can make up quantity as per your requirement then.

Hope it helps!

Hello,

I have a doubt, pls see the attached protocol link below on oil red ad o. The major differences is, 1 of them saying not o air dry the cells ever, the other says to air dry.I m confused.

Also, true, that we should not use alcohol based solvent, as that might dissolve oil, but arent the cells already fixed and also the 4%PFA is made in 100% isopropanol. Pls help me figure these things out, as I am staining the 3T3L1 cells with oil red and O. file:///Users/srijitamukhopadhyay/Downloads/Oil%20Red%20O%20Stain%20for%20In%20Vitro%20Adipogenesis_Original_28564.pdf

Hello Srijita,

I also came across two different protocols about the oil red o staining (respective let the cells dry or not), but I cannot follow your link unfortunately. I tried both of them and there is not a big difference, so you just can peak one you like. I do not see a problem in using i-prop for fixation step as normally the stain is dissolved in i-prop based solution. Maybe you should avoid washing of the stained cells with i-prop, as it will extract the stain from them... Anyway if it disturbs you for some reason you can use formalin-PBS solution for the fixation step. Hope it helps!

Best wishes,

Stoyan

Hello Ajay, the stain from Sigma is ready made in 0.5% isopropanol , why do you need to dissolve and dilute in water again, I dont understand

@ Srijita,

Yes it is ready made and you can use it directly by incubating cells for 15 minutes. But sometimes people have lots of assay wells and reagent is a limitation. That's why 1:1 dilution and incubation for 30 to 40 minutes will also give similar results. Also there is over-staining sometime if people leave the ready made stain in cells for too long. So in my PhD I tried both and when you can achieve a good staining with less volume, why to waste.Though its fine to use it anyway.

Hope it helps!

hi Ajay,

With specific type of sigma buffer you used because it is not really mentioned ( Neutral 10%, Carbonate Buffered , Formalin, 10%, Neutral Buffered with 0.03% Eosin , Formalin solution, neutral buffered, 10%

histological tissue fixative etc.....)

let me know thanks

Hi,

I used 10% NBF from Sigma (formalin solution neutral buffered 10% in above case).

Hope it helps!

Hi everyone, this is a really helpful thread, so thank you. I tried staining adipo-differentiated bone marrow MSCs today and the staining didn't work. I used differentiation media for 21 days and saw lipid droplets in cells, suggesting adipogenesis had occurred. For staining I removed media, washed cells with PBS, added 4 % paraformaldehyde for 30 minutes at room temperature, removed and washed cells with diH2O twice. Added 60 % IPA to cells for 5 minutes, removed then added my stain. I'm using a solution of Oil Red O from sigma (0.5 % ORO in IPA). Initially I used the stain as-is for 15 minutes, then removed and washed cells with diH2O 5 times. The stain was not present in any of the cells. Could this be due to the 60 % IPA treatment for 5 minutes? Any advice would be so helpful! Thanks!

My first thought is that it is the pre-treatment with the isopropanol that is dissolving the lipid. When I do my staining the 0.5% oro in IPA is diluted to 60% and then filtered before use.

Hi Kathryn Murray , did you get a good Oil Red O staining avoiding the incubation with 60% IPA? I have been reading some protocols and this step is very confusing for me.

Hi Cl Pe - in short, yes I got good staining when I did the pre-incubation step with 60 % IPA.

I repeated the experiment and it looks like I needed to dilute the dye to 60% (i.e. take the 0.5 % Oil red O solution in 100 % IPA and dilute 6:4 in diH2O to get 60 % Oil red O, 40 % H2O). This gave good staining of the lipid droplets. I also checked whether this worked when I did the pre-incubation step with 60 % IPA and it did (so you don't need to avoid the pre-incubation so long as it's with 60 % IPA too). My updated method is now:

Removed media and washed cells with PBS, then added 4 % paraformaldehyde for 30 minutes at room temperature, removed and washed cells with diH2O twice. Added 60 % IPA to cells for 5 minutes and removed. Diluted Oil Red O stock solution (0.5 % Oil Red O in 100 % IPA) 6:4 in diH2O to get a working solution of oil red O in 60 % IPA, 40 % H2O. I let the working solution stand for 10 minutes then filtered it directly before use. Added stain to cells for 15 minutes, removed and washed wells 5 times with diH2O. Removed H2O and added PBS then images wells using a light microscope.

I think the reason my staining didn't work last time is that the Oil Red O stain was in 100 % IPA. When I added this to the cells, the stain remained in the IPA solution, rather than transferring to the lipid droplets. When the stain is in the 60 % IPA solution, the dye is more soluble in the lipid droplets than 60 % IPA and so it transfer into the droplets. This is a very basic explanation! The pre-incubation step with 60 % IPA removes background lipids from the sample, reducing background staining.

Hope that helps and this works for you too.

It happened to me once that I apply 0,2% ORO working solution and I didn't get any staining. When I checked back the steps of my protocol I realized that instead of dissolving my 0.5% w/v ORO stock (dissolved in IPA) in water by adding 2 parts of my stock to 3 parts of distilled water I used IPA again. When then repeated the staining this time properly dissolving the working solution in water and we got beautifully stained adipocytes. So, probably to 60% IPA preincubation dissolved all the lipids in your cells