Oil red o working solution

SVF-differentiated adipocytes were fixed with 4% paraformaldehyde (Sigma-Aldrich, P6148) in PBS and were washed twice with PBS. Completely air-dried at room temperature, cells were then incubated with Oil Red O working solution (Sigma, O0625) at room temperature for 1 h. Cells were washed 3 times with PBS before images were acquired for analysis. For quantification of Oil red O staining, cells were washed with PBS twice and then 200 μL DMSO was added to each well. After incubation for 10 min with gently shaking, samples were measured for OD at 510 nm.
Tissues were immediately frozen in liquid nitrogen and cut using a Leica CM-1850 cryostat microtome (Leica, Wetzlar, Germany). Afterwards, 16 mm-thick sections were fixed in 4% formaldehyde for 10 min and stained with filtered 0.5% Oil Red O, which was dissolved in isopropyl alcohol, for 15 min at room temperature.

Liver tissues were fixed in formalin and then embedded in paraffin. The slide section of liver tissue was stained with hematoxylin–eosin (H&E) and Oil Red O for hepatic histological analysis. Paraffin-embedded livers were cut into sections and stained with primary antibodies for protein localization. Epitope specific antibodies were used for ChREBP, α-SMA, CK18, SIRT1, OGT, F4/80 and SOD1 staining. Chromogenic detection was using with the DAB (DAKO). Then, the slides were dehydrated and mounted in Safemount embedding medium (Labonord, France).
For Oil Red O staining, frozen liver sections (10 μm) were fixed with 10% formaldehyde for 10 min. To measure lipid accumulation, slides containing the cryosections were washed in water, immersed in 100% propylene glycol for 5 min, and then incubated in 0.5% Oil Red O working solution (Sigma O0625) for 30 min at room temperature. The slides were washed with 85% propylene glycol (dilution in distilled water) for 20 min. Slides washed in distilled water for 5 min and then mounted in aqueous mounting medium (mixed gelatin, glycerol, and distilled water).