This article details the process of Hematoxylin and Eosin staining, a crucial technique in histology for visualizing tissue structures. The method enhances the contrast between cellular components, allowing for detailed examination under a microscope.
Begin Hematoxylin and Eosin staining by taking a glass slide containing the paraffin-embedded tissue section. Dip the slide in a deparaffinization solution containing xylene - an organic solvent that dissolves the paraffin layer around the tissue, making it accessible to stains in subsequent steps.
Treat the tissue section with absolute alcohol to remove the xylene. Immerse the tissue in decreasing concentrations of alcohol. This step rehydrates the tissue by replacing the alcohol with water for better compatibility with water-soluble dyes.
Stain the tissue with basic hematoxylin, which in its oxidized form binds to mordant - a metal cation. This complex electrostatically binds to the nucleic acids in the genome, staining the nucleus purple.
Submerge the stained section in an acidic differentiation solution to remove excess hematoxylin from the cytoplasm, making the dye-retaining nucleus appear distinct. Treat the tissue with an alkaline bluing agent to neutralize the free acids. This regresses the purple dye color to blue, enhancing the nuclear details.
Immerse the slide in acidic eosin solution to stain the basic protein components in the cytoplasm and the connective tissue fibers in shades of red to pink. Treat the tissue with alcohol to eliminate excess eosin. Use increasing concentrations of ethanol to dehydrate the tissue. Dip the slide in xylene to clear any remaining reagents.
Upon imaging, the cells in the tissue section show clear blue nuclei and pink cytoplasmic regions.
Deparaffinize the slides by immersing them in 100% xylene, twice, for 15 minutes each time. Then, rehydrate the slides through an ethanol gradient. For each solution, dip 8 to 10 times, for two seconds per dip, until the liquid runs cleanly off the slides. Place the slides in 100% filtered Harris hematoxylin for four minutes.
After four minutes, quickly transfer them back to the container of deionized water. Run deionized water into the back corner of the container farthest away from the sections. Empty the container periodically until the water is no longer purple.
Quickly check the hematoxylin intensity on a dissecting microscope using gooseneck lights. Return the slides to 100% hematoxylin for one minute, if further staining is required.
Once the desired hematoxylin staining has been obtained, dip the slides twice in 0.05% hydrochloric acid, then, immediately transfer them back to the container with clean deionized water. Empty and refill the container with water twice.
Transfer the slides to two containers of 95% ethanol for 30 seconds each. Place the slides into the eosin Y phloxine-B solution for two minutes.
After the two minutes, transfer the slides back to the previous 95% ethanol container, then check the intensity of the color under the dissecting microscope. If the staining is not sufficient, return to the eosin solution for 30 seconds. Repeat as necessary.
When staining of the desired intensity has been obtained, transfer the slides to 100% isopropanol for 15 seconds. Replace with fresh 100% isopropanol, and place the slides back in the isopropanol for another 15 seconds. Repeat this process for a total of six isopropanol washes.
After immersing in 100% xylene for three minutes, remove one slide, add sufficient mounting medium to cover the sections, and then, dip the slide in 100% xylene.
Apply a coverslip to the slide and blot any excess mounting medium on a paper towel until only a thin line is seen. Then, dip a tissue into the xylene and wipe the back of the slide to remove any medium that has dripped.
Place the slide flat on a sturdy but mobile surface, like a piece of cardboard, and allow the xylene to evaporate in the hood for 10 minutes. Leave the slides at room temperature overnight to allow the mounting medium to harden before imaging.
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