Oil Red O Staining Protocol

I. Principle

Oil Red O is a lipid-soluble azo dye with strong hydrophobicity: it dissolves well in organic solvents but poorly in aqueous media. When tissue sections or cells are exposed to Oil Red O working solution, the dye preferentially dissolves into and accumulates within intracellular neutral lipids (e.g., triacylglycerols, cholesteryl esters), producing orange-to-red lipid droplet signals, while the aqueous phase and most hydrophilic tissue components retain little dye, enabling relatively selective lipid staining.

II. Preparation

1.Materials

Fresh tissue (e.g., adipose, liver), 4% paraformaldehyde fixative, 60% isopropanol, Oil Red O stock (0.5 g Oil Red O in 100 mL isopropanol), Oil Red O working solution (mix stock with distilled water at 3:2 immediately before use, mix well and filter), hematoxylin, 1% acid alcohol differentiation solution, eosin, neutral mounting medium, slides, coverslips, staining jars, light microscope.

2.Equipment

Microtome, incubator, micropipettes, centrifuge, balance, and other standard histology equipment.

III. Sample Handling

1.Fixation

Place fresh tissue promptly into 4% paraformaldehyde for ~4–24 h to stabilize morphology and structure. Adjust time to tissue size and type.

2.Dehydration and embedding

After fixation, pass through graded ethanol (70%, 80%, 90%, 95%, 100%), 1–2 h each. Clear in xylene, then infiltrate with molten paraffin and embed to form blocks.

IV. Sectioning and Dewaxing

1.Sectioning

Cut paraffin blocks into ~4–6 μm serial sections, mount on slides, and air-dry.

2.Dewaxing and rehydration

Place slides in xylene I and II for 10–15 min each to remove paraffin. Rehydrate through 100%, 95%, 90%, 80%, 70% ethanol, ~5 min each, then rinse in distilled water.

V. Oil Red O Staining

1.Staining

Immerse fully rehydrated sections in freshly prepared, filtered Oil Red O working solution at 37°C for ~10–20 min. Fine-tune time by tissue type and lipid content; stop when lipid droplets show uniform orange-red.

2.Differentiation

Briefly differentiate in 60% isopropanol for a few to ~10 seconds to remove background while retaining lipid-specific signal; rinse gently with distilled water.

3.Counterstain (nuclei)

Immerse in hematoxylin for ~3–5 min to stain nuclei. Rinse with tap water or gently running water to remove excess dye.

4.Differentiation and bluing

Dip briefly in 1% acid alcohol differentiation solution for a few seconds to lightly differentiate nuclei, then rinse under running water ~10–15 min to blue nuclei to a clear blue.

VI. Mounting and Microscopy

1.Optional cytoplasmic counterstain

If needed, immerse in eosin for ~1–2 min after bluing to give cytoplasm a pale pink/red. Rinse quickly with distilled water.

2.Dehydration and clearing

If mounting with neutral resin, dehydrate again through 70%, 80%, 90%, 95%, 100% ethanol, ~3–5 min each, then clear in xylene I and II for ~5–10 min each. Avoid drying or curling of sections.

3.Mounting

Apply neutral mounting medium, place coverslip carefully, minimizing bubbles. Air-dry at room temperature or cure at low temperature in an oven until fully set.

4.Observation and documentation

After curing, observe under a light microscope. Scan with low power for overall architecture and lipid distribution, then switch to high power to assess droplet size, morphology, and staining intensity. Capture images and record as needed.

VII. FAQs

Q1: Background is heavy and the entire section looks red—lipid droplets are unclear.

A: Common causes: insufficient filtration of working solution; over-staining; inadequate differentiation with 60% isopropanol; drying or precipitate formation during staining. Improve by filtering before each use, shortening staining time, slightly extending differentiation, and keeping sections submerged to prevent drying. If precipitates persist, prepare fresh working solution.

Q2: Can paraffin and frozen sections be compared directly?

A: They differ methodologically in lipid preservation. Paraffin processing removes some lipids during dehydration/clearing, often yielding weaker signals—suited to qualitative/semi-quantitative observation. Frozen sections preserve lipids better and are preferable for comparative and quantitative droplet analyses. For strict comparisons, use the same sectioning method and unified pretreatments; avoid mixing paraffin and frozen results.

Q3: Does fixation time need to be exact? What if it’s too long?

A: It need not be precise to the minute but should stay within a reasonable window. The common 4–24 h range works for most cases. Too short risks poor structural integrity and section quality; too long risks over-crosslinking, hindering dye penetration and affecting droplet morphology/distribution. For small/thin tissues, shorten appropriately; for larger/harder tissues, modestly extend, but avoid soaking for days.

Aladdin: https://www.aladdinsci.com/