Hematoxylin and Eosin (H&E): Classification, Principle, Procedure, Interpretations

Table of Contents

Hematoxylin and Eosin (H&E) staining is a widely used histological staining technique in pathology and histology. It is used to colourize tissues and cellular structures to observe and study them under a microscope.


Staining is a process by which a colour is imparted to sectioned tissue. Specially manufactured dyes are used for this purpose. These dyes are prepared by adding an auxochrome to a chromophore. An auxochrome is a compound which when added to a chromophore forms a dye. This may be acidic or basic. A chromophore is a compound that, which although coloured, does not have the properties of a dye or stain. The dye stains the tissues by binding with specific sites. Compounds called mordants help in achieving this binding.


All stains are composed of an acid and a basic component. Generally, the stains are classified as:

  • Acid stains
  • Basic stains
  • Neutral stains

Acid Stains:

In an acid stain, the acidic component is coloured and the basic component is colourless, in acid fuchsin, which is composed of sodium and rosaniline trisulphonic acid, the sodium is colourless and rosaniline trisulphonic acid is coloured. Acid dyes stain basic components of tissue e.g., cytoplasmic proteins. The colours imparted are shades of red. The most commonly used acid dye is eosin.

Basic Stains:

In the basic dyes, the basic component is coloured and the acidic component is colourless. The example is basic fuchsin. Basic dyes stain acidic components of tissue e.g., nucleic acids. The colours imparted are shades of blue. The most commonly used basic dye is haematoxylin.

Neutral Stains:

When an acidic dye is combined with a basic dye a neutral dye is formed. As it contains both colouring components it stains all components of tissue but with different colours. This is the basis of Romanowsky stains (e.g., Leishman stain).


Like processing, staining can also be performed manually or mechanically.

Manual Staining:

In a small laboratory where only a few slides are stained, this is the method of choice. It is time-consuming but economical. Reagent containers are placed in a sequence. Slides are placed in a carrier and are then moved from one container to another at specified intervals till the process is complete.

Automated Staining:

The above procedure is performed with the help of a mechanical device similar to the one described for processing. Automated stainers of various kinds are now freely available. In these, the reagent jars are arranged according to a desired sequence. The carrier containing slides is rotated through these at intervals, which are set by the operator (Figure 56.2). These are usually microprocessor-controlled and are programmable.

The advantages are:

  • Reduce manpower requirements
  • Precise control of the timing
  • Large number of slides stained simultaneously
  • Less reagent consumed


Hematoxylin and Eosin (H&E) are commonly used for routine histopathology and diagnostic cytology. Its particular value lies in its ability to impart proper differentiation to distinguish between different types of connective tissue fibres and matrices, by staining them in different shades of red and pink. 


First, the tissue is cleared of all wax and then rehydrated to facilitate the entry of dyes. The tissue sections are then sequentially exposed to a basic dye e.g., Harris’s Haematoxylin and an acid dye e.g., eosin. This stains both the basic and acid components of the tissue.


Harris’s Haematoxylin:

  • Haematoxylin crystals 5.0 g
  • Alcohol 95% 50 ml
  • Ammonium or Potassium Alum 100 g
  • Mercuric oxide 2.5 g
  • Distilled water 1 litre
  • Glacial acetic acid 40 ml

Dissolve separately by heating, haematoxylin in alcohol and alum in water, mix and rapidly boil. Remove from flame and add mercuric oxide. Reheat for 1 minute or until it becomes dark purple. Remove from flame and cool in a basin of cold water. The stain is ready to use. Add 2-4 ml of Glacial acetic acid per 100 ml of solution if desired.

Acid alcohol:

Mix one litre of 70% alcohol with 10 ml of concentrated hydrochloric acid.

Ammonia water:

Mix 2-3 ml of strong ammonia with one litre of tap water.

Alcoholic Eosin solution:

  • Eosin (water soluble) 2 g
  • Distilled water 160 ml
  • Alcohol 95% 640 ml

Other reagents:

Xylol, absolute alcohol, rectified spirit and methylated spirit are also needed.

Staining procedure

  1. Put the sections fixed on a glass slide in xylol for 3 min.
  2. Then transfer to absolute alcohol for 3 min.
  3. Transfer to rectified spirit (80% alcohol) for 2 min.
  4. Place in methylated spirit for 2 min.
  5. Wash the slide in running water for 1 min and put it in Harris haematoxylin for 3-5 min.
  6. Wash in running water for 30 seconds and wash the excess dye in 1% acid alcohol by continuous agitation for 15 seconds.
  7. Wash in running water for 30 seconds.
  8. Give 2-3 dips in ammonia water solution until tissues attain a blue colour.
  9. Wash in running water for 2-3 dips.
  10. Counterstain with eosin for 2-3 min.
  11. Wash in running tap water for 30 seconds.
  12. Dehydrate by keeping in increasing concentrations of alcohol (2-3 dips in 70%, 95% and absolute alcohol).
  13. Clear it in Xylol and mount it with Canada balsam.


  • Nuclei Bright blue
  • Muscle, keratin Bright pink
  • Collagen and cytoplasm Pale pink
  • Erythrocytes orange-red

Notes and Precautions

Other haematoxylins like Mayer’s haematoxylin may also be used. All have different methods of preparation. The reagents must be checked daily for deterioration and changed when needed. In the manual method, the xylol and alcohols must be changed daily, haematoxylin once a week, eosin and acid alcohol twice a week, and ammonia water daily. This regimen may be modified by the amount of usage. In the automatic stainer, xylol, alcohols, eosin and acid alcohol, are changed twice a week. Haematoxylin is changed once in two weeks and ammonia water is changed daily. The quality of alcohol must be checked before use. This can be done by adding 4-5g of copper sulphate crystals to a Coplin jar containing alcohol. If the colour remains unchanged (bluish-white) for 10 min, it is acceptable. If the colour changes to green the quality of alcohol is unsuitable for processing.



Please enter your comment!
Please enter your name here