ORO and hematoxylin staining of A549 cells
Oil-red-O: description of the method
Oil red-o, a sensitive stain for neutral lipids, has been proven to be successful in assessing lipid accumulation in tissue histological samples under both physiological and pathological conditions. [Mehlem, A., Hagberg, C., Muhl, L. et al. Imaging of neutral lipids by oil red O for analyzing the metabolic status in health and disease. Nat Protoc 8, 1149–1154 (2013). https://doi.org/10.1038/nprot.2013.055]
Oil red O (ORO) staining is a reliable method to assess the presence of lipid vesicles in various cell types in culture as well, not just in tissue samples.
Using an optical microscope, it is possible to visually analyze ORO-stained lipid vesicles.
A quantitative, morphological, and dimensional evaluation of the lipid vesicles can be achieved by analysing the image with the Image J (or Fiji) program.
The goal of this post is to present a quick method for staining lipid vesicles with ORO in cultured cells.
This method can be combined with an emathoxylin counterstaining, useful for visualizing the nuclei and therefore evaluating the number of cells in culture.
Required reagents
- 4% paraformaldehyde solution
- Oil-red-O powder
- Isopropanol
- HBSS
- PBS
- Double distilled water
If you want to counterstain the nuclei
Required materials
- Cells growth on coverslip (round coverslip, 24 multiwell, cell culture media and reagents)
- Automatic single-channel, mechanical, variable volume pipettes
- beakers and cylinders
- Serological pipettes and pipettor
- falcon tube
- filter paper
- funnel
- syringe filter 0.2 um
- plastic syringe
- laboratory balance
- microscope slide
Protocol
To achieve optimal visualization, cells need to be grown on a coverslip. (a procedure for properly preparing coverslips to use in cultured cells is outlined here).
Plate, let the cells grow and treat them as necessary.
Prepare a 5% ORO solution: Mix 0.5g of oil red O with 10 ml of 100% isopropanol. This solution can be kept tightly closed (isopropanol evaporates quickly) for one month.
Rinse quickly the cells with HBSS (for cell grown in 24 multiwell, use 500 ul of HBSS) at 37°C to remove the cell medium.
Fix cells using 4% paraformaldehyde (here is the protocol to make the paraformadehyde solution) for 10 minutes (for cell grown in 24 multiwell, use 250 ul of paraformaldehyde).
Remove fixative and rinse with phosphate buffered saline (PBS, 500 ul) for 3 times in order to completely remove formaldehyde
Meanwhile, mix the oil-red-o stock solution with ddH2O in a 6:4 ratio and let stand for 10 minutes (ORO working soluion).
This solution is for single use and cannot be stored
Filter with a coffee filter or other fast filter to remove the coarser oil red particles that remain in suspension then use a 0.2 micron syringe filter to remove the finer particles.
If the oil red o is not filtered properly you will have a lot of background staining. (avoid using the syringe filter straight away because it gets blocked by the larger particles, preventing filtration. Don’t just use filter paper because it doesn’t retain the smaller particles which would settle on the cells, “dirtying” the image)
Add the ORO working solution to the cells (for a 24 well, add 250 ul).
Leave the ORO working solution for 10 minutes, then remove and rinse well with 1 ml of dH20 (3 times).
à you can stop here if you want to obtain a staining of the lipid vesicles only, if you want to proceed with the coloring of the nuclei, continue with the following stepsß
Add 200 ul of Mayer’s hematoxylin for 15 minutes (Haematoxylin and Eosin Stain of the Tissue Section | SpringerLink)
The intensity of the blue staining correlates with the amount of DNA in the nuclei and the length of time the sample spent in hematoxylin. Therefore you can change the incubation time if you want nuclei with a more or less intense color
Mayer’s hematoxylin is a water based stain. DO NOT USE an alcohol based hematoxylin (such as Harris hematoxylin) because it will extract lipid and the oil red dye!
Remove hematoxylin (can be reused) and make it bluish with Scott tap water (rinse 2 times with 1000 ul) leave the last rinse 15 minutes to fully develop the blue color of the nuclei
Rinse with water
Mount the coverslip onto a microscope slide using an appropriate mounting medium (90% glycerol in water). Secure the coverslip with nail polish