H&E Staining of Paraffin-Embedded Tissue Sections: A Differential Staining Technique to Visualize Liver Tissue Sections Using Combination of Hematoxylin and Eosin Dyes

Abstract

Source: Ellis, J. L et al., Histological Analyses of Acute Alcoholic Liver Injury in Zebrafish. J. Vis. Exp. (2017).

In this video, we describe a technique of hematoxylin and eosin staining of paraffin-embedded tissue sections to study their histology. The procedure includes deparaffinization of the tissue followed by staining the cells in the tissue. Upon staining, the nucleus appears distinctly blue, and the proteins in the cytoplasm display pink coloration.

Protocol

1. Preparation of staining solutions

1. Filter Harris hematoxylin stock solution to remove any solids that have precipitated.
2. Fold a coffee filter in half so that the filter creates a cone. Open the side panel so that the liquid runs through the filter, allowing any debris to be caught.
3. Place the filter inside a funnel and the funnel into a glass media bottle. Gradually pour the stock hematoxylin through the filter.
   CAUTION: Hematoxylin is hazardous in case of eye contact, ingestion, and inhalation. Always wear a lab coat and gloves and handle the stock solution in a chemical hood.
4. Prepare 0.05% HCl by adding 125 µL of 12 N HCl to 250 mL of ddH₂O.
5. Prepare eosin Y-phloxine B solution mixture by combining 25 mL of 1% eosin Y (aq), 2.5 mL of 1% phloxine B (aq), 195 mL of 95% ethanol, and 1 mL of glacial acetic acid in a glass bottle. Mix well by swirling and store at RT
   CAUTION: Eosin Y is hazardous in the case of eye contact, ingestion, and inhalation. Always wear a lab coat and gloves and handle the stock solution in a chemical hood.
6. Prepare 2% ethanol by adding 2 mL of pure ethyl alcohol to 98 mL of egg water.
   CAUTION: Ethyl alcohol is an eye irritant. Always wear proper protective equipment when handling ethyl alcohol. It is also flammable and should be handled and stored accordingly.
   NOTE: Prepare fresh 2% ethanol each time before conducting the acute ethanol treatment.

2. Hematoxylin and Eosin Staining of Paraffin Sections

1. Deparaffinize the slides by dipping them in 100% xylene for 15 min; change it to fresh 100% xylene for another 15 min.
2. Rehydrate the slides by dipping them through the following series of graded ethanol until the liquid runs cleanly off the slides. For each solution, dip 8-10 times, 2 s per dip: 100% ethanol, 100% ethanol, 95% ethanol, 95% ethanol, 70% ethanol, 50% ethanol, 30% ethanol, and deionized water.
   NOTE: The protocol can be stopped here, and the slides can be left at RT in water for several hours. If necessary, the slides can be stored at 4 °C in water O/N.
3. Place the slides in 100% filtered Harris hematoxylin for 4 min. Immediately transfer them back to the container with 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.
   NOTE: Do not let the water run directly onto the slides, as the sections may come off the slides.
4. Quickly check the hematoxylin intensity on a dissecting microscope using gooseneck lights; do not allow the slides to dry. If the stain has reached the desired intensity, continue to the next step. If the color is not dark enough, place the slides in 100% hematoxylin for 1 min and repeat the water washes before checking again.
   NOTE: The hematoxylin needs to be dark enough so that the color is not lost during the eosin staining. However, if hematoxylin staining is seen in the cytoplasm, the sections are overstained.
5. Dip the slides twice in 0.05% HCl and immediately transfer them back to the container with clean deionized water. Empty the water and refill the container with water twice.
6. Transfer the slides to 95% ethanol for 30 s and then transfer them to a new container with 95% ethanol for 30 s.
7. Place the slides into the eosin Y-Phloxine B solution for 2 min. Transfer the slides back to the previous 95% ethanol container and quickly check the intensity of the color under the dissecting microscope. If the staining is sufficient, proceed to the next step. If not, return to the eosin solution for 30 s, check again, and repeat as necessary.
   NOTE: Sufficient eosin staining is bright pink and appears in distinct contrast to the hematoxylin stain. Be sure to wipe any solution off the back of the slides when checking under the microscope so that no false color is observed. Because eosin is made up of 95% ethanol, prolonged periods of time in 95% ethanol while checking color intensity may leach out some of the color.
8. Transfer the slides to 100% isopropanol for 15 s. Replace with fresh 100% isopropanol and place the slides back in the isopropanol for another 15 s. Repeat this process for a total of 6 isopropanol washes.
   CAUTION: Isopropanol is an eye and respiratory irritant. Always be sure to wear proper protective equipment and avoid splashing. It is also highly flammable and should be stored and handled accordingly.
   NOTE: To save on reagents, discard only the first (pinkest) isopropanol wash. Keep the other five wash solutions in bottles numbered 1 through 5, to be reused for the next staining. When reusing washes, start with the bottle numbered "1" and discard after use. Once wash "2" has been used, pour it into bottle "1," and so on. The final wash should always be fresh isopropanol.
9. Place the slides in 100% xylenes for 3 min. Remove one slide at a time and place a coverslip.
   1. Add sufficient mounting medium to cover the sections and dip them in 100% xylene.
      NOTE: This step will smooth the surface of the mounting medium and remove any bubbles.
   2. Apply a coverslip to the bottom of the slide.
      NOTE: The mounting medium should pull the slide into position. If the coverslip is not straight or completely seated, gently tap it into place.
   3. Blot any excess mounting medium on a paper towel until only a thin line is seen. Dip a tissue wipe 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. Coverslip all the remaining slides in the same fashion. Allow the xylene to evaporate in the hood for 10 min.
      NOTE: Any materials contaminated with xylene should be removed and discarded in a sealed container in the hood to prevent any vapors from escaping.
10. Allow the mounting medium to harden at RT O/N.

3. Imaging and Storage of Stained Slides

1. Image sections on a compound inverted microscope.
   NOTE: Slides can be kept at room temperature indefinitely.

Materials

95% ethanol (EtOH)	Decon Labs, Inc.	2801	Flammable
Charged slides	Fisher Scientific	12-550-16
Dissecting microscope	Leica Biosystems	Leica Mz 95
Eosin-Phloxine stain set	Newcomer Supply	1082A
Ethyl alcohol	Sigma-Aldrich	E7023	Flammable
Harris hematoxylin	Poly Scientific R&D Corp.	s212	Irritant
Hydrochloric acid (HCl)	Fisher Scientific	A144	Irritant
Isopropanol	Newcomer Supply	12094E	Flammable
Xylenes	Fisher Scientific	X3S-4	Irritant