2. Introduction:
Hematoxylin and Eosin (H&E) staining is the corner stone of tissue based diagnosis
The process stains thin tissue sections so that pathologists can visualize tissue morphology.
The process uses a Hematoxylin dye to stain cell nuclei (and other parts) Blue and an Eosin dye
to stain other structures pink or red.
Hematoxylin binds strongly to acids and consequently binds to nuclear DNA and stains nuclei
blue.
Properly applied, this technique provides exceptional detail of tissue structure and the makeup
of the cells.
This detail is required for tissue-based diagnosis, particularly in the detection and classification
of infection, cancer or metabolic disease.
3. Routine H&E staining plays a significant role in tissue-based diagnosis.
By coloring otherwise transparent tissue sections, and allowing cell structures including the
cytoplasm, nucleus, and organelles and extra-cellular components to be clearly visible under the
microscope.
In a histology laboratory, all specimens are initially stained with H&E and additional stains are
only ordered if additional information is needed to provide a more detailed analysis.
Staining with H&E is very reliable although it does show some variation and the stain density is
considerably affected by the thickness of the sections – thicker sections take up more stain.
Generally done before any additional staining techniques.
As H&E can confirm the basic tissue type and help to localize the lesion. (Any area of damage,
infection, inflammation, tumor, necrosis or otherwise abnormal tissue.).
4. Since most cell structures are transparent, very little detail of the structure can be seen, unless
the cells are stained.
The same is true of components of the extracellular matrix. Because different parts of the cell
are biochemically different, they take up specific stains to varying degrees.
ROUTINE H&E STAINING in Paraffin Embedded Section (Regressive Staining) Fixation:
Most fixatives can be used except osmic acid solutions which inhibit hematoxylin.
5. Procedure:
1. Clear paraffin embedded sections in first xylene bath for 3 minutes.
2.Transfer to second xylene bath for 2 to 3 minutes.
3.Immerse in first bath of absolute ethyl alcohol for 2 minutes.
4.Transfer to a bath of 95% ethyl alcohol for 1 or 2 minutes.
5.Rinse in running water for 1 minute.
6.Stain with Harris alum hematoxylin for 5 minutes (Ehrlich's hematoxylin requires 15-30 minutes).
6. 7. Wash in running tap water to remove excess stain.
8. Differentiate in 1% acid-alcohol (1 ml concentrated HCl to 99 ml. of 80% ethyl alcohol) for 10-
30 sec. monitoring the changes in color microscopically until only the nuclei are stained.
9. Rinse in tap water.
10. Blue in ammonia water (average of 5 minutes) or 1% aqueous lithium carbonate until the
sections appear blue (about 30 seconds).
11. Wash in running water for 5 minutes.
12. Counterstain with 5% aqueous eosin for 5 minutes. If alcoholic eosin is used, the time can be
reduced to 30 seconds or 1 minute.
12. If aqueous eosin is used, wash and differentiate in tap water under microscope control until the
nuclei appear sharp blue to blue black and the rest of the tissue appear in shades of pink. If
alcoholic solution is used, differentiate with 70% alcohol.
7. Dehydrate, clear and mount.
7. NOTE: -
For tissues fixed with mercuric chloride, the staining time in hematoxylin should
be increased slightly while duration of eosin staining should be reduced.
The mercury should be removed using a 0.5% solution of iodine in 80 to 95%
alcohol and rinsed in water.
The iodine is then removed by placing the slide in 3% sodium thiosulfate solution
for 1 to 5 minutes and washing it well in running water for 3 to 5 minutes.
Alternatively, mercury deposits may be removed after sections are hydrated, by
immersing the sections in Gram's or Lugol's iodine for 5 minutes, followed by
sodium thiosulfate and subsequently washing the section in water prior to
staining.
Staining may be prolonged for chromium and osmium fixed tissues (e.g.
Flemming's fluid), for tissues subjected to long acid decalcification, and after
prolonged storage in acid formalin or 70% alcohol
8. H & E staining of Frozen Sections for Rapid Diagnosis (Progressive Staining)
1. Orient section in the block and freeze with liquid nitrogen.
2. Cut cryostat sections at 5-10 micron.
3. Mount sections on to albuminized slides and dip in 10% formalin to fix.
4. Rinse rapidly in water.
5. Stain with Harris hematoxylin for 30-45 seconds.
6. Rinse in tap water.
7. Blue in ammonia water for 5 seconds.
8. Rinse in tap water.
9. Counterstain with 5% aqueous eosin or 1% alcohol eosin for one minute.
10. Rinse in tap water.
11. Dehydrate in increasing concentrations of alcohol.
12. Clear with xylene.
13. Mount with cover slide.
9. It is somewhat less favored than regressive staining due to the difficulty of producing
sufficiently intense progressive staining of cell structures without staining other parts, thereby
resulting in diffused color and obscured details.
For convenience, reagents for this rapid H&E stain are generally arranged in sequence using a
series of Coplin jars.
This method takes only 5-10 minutes and produces well-differentiated sections that are semi-
permanent and can be stored.
The remaining portion of tissue must be kept for routine processing and are made for
comparison with frozen sections.
10. Precautions in Staining
Stains on the skin should be avoided not only because they are signs of poor technique but because stains
are health hazards per se, being slowly absorbed by the skin and eventually producing side effects.
Stains may be effectively removed from the skin by prompt topical application of 0.5% acid alcohol,
followed by rinsing with tap water.
To remedy the condition, the section is placed in a Coplin jar containing xylol to dissolve the adhesive.
The slide is run back thru the various processes up to the point where the fault was.
A fresh solution is used, and the tissue is re-stained. Stains may be saved and used again for as long as they
have not lost their staining properties.
11. Sections are usually rinsed with distilled water before placing them in used stains.
Formation of precipitate in staining solution and poor staining results signify loss of staining
property and hence, the stain should be discarded and replaced with a fresh solution.
Failure of sections to remain on the slide during staining could have been due to a dirty or oily
slide.
Slides may have been carried thru the first alcohol baths too fast, resulting in a rapid but
incomplete dehydration; or paraffin sections may not have been thoroughly spread on the slide
when mounted
12. Failure of staining may be due to paraffin, fixative, or decalcifying solution that has not been thoroughly washed out and
removed.
Early fixation in alcohol before paraffin embedding may have been incorrect, for which no remedy can be made. Alternatively, the staining
solution may be faulty.
Hematoxylin solutions may not have been properly and sufficiently ripened.
Hematoxylin must not be used too soon after preparation to ensure complete ripening.
Impurities found in the dye or in the water solvent will affect not only the solubility of the dye but even the intensity of the staining
reaction, necessitating purification and filtering of the dye.
Stains that have already been deteriorated should be replaced.
If, after staining, sections are fuzzy and do not appear clear under the microscope, xylol should be replenished.
There may be water in the absolute alcohol, moisture in the coverslip, or too much egg albumin on the slide, thereby obliterating the
image of the stained tissue.
And often, acid-alcohol decolorizer may not have been completely removed, or a film from alkaline alcohol may have been carried along.
13. To remedy the condition, the section is placed in a Coplin jar containing xylol to dissolve
the adhesive.
The slide is run back thru the various processes up to the point where the fault was; a
fresh solution is used, and the tissue is re-stained. Stains may be saved and used again
for as long as they have not lost their staining properties.
Sections are usually rinsed with distilled water before placing them in used stains.
Formation of precipitate in staining solution and poor staining results signify loss of
staining property and hence, the stain should be discarded and replaced with a fresh
solution.
Failure of sections to remain on the slide during staining could have been due to a dirty
or oily slide.
Slides may have been carried thru the first alcohol baths too fast, resulting in a rapid but
incomplete dehydration; or paraffin sections may not have been thoroughly spread on
the slide when mounted
Albumin fixative may be too old, as suggested by the loss of its clear color, or by
emission of an odor. To avoid this, adhesives should be prepared in small amounts
(around 1 ounce) which may last for 2-3 months.
14. COLLODIONIZATION OF SECTIONS
Paraffin ribbons containing air bubbles, torn or inadequately infiltrated sections are likely
to float from the slide when deparaffinized and stained.
They are more firmly attached by coating the slide with dilute (thin) celloidin solutions, a
process known as collodionization, which is also recommended for sections that will be
subjected to strong alkaline or acid solutions and for tissues that contain glycogen for
demonstration.
Procedure:
1. Deparaffinize in xylene.
2. Dehydrate thru absolute alcohol.
3. Dip individual slides in Coplin jar containing dilute ether alcohol solution.
4. Dip in dilute ether solution of celloidin (thin celloidin).
5. Hold slide on one end for 1/2 to 1 minute to drain or until the section begins to whiten
around the edges.
6. Wipe off the back of the slide and place in 80% alcohol for 3-5 minutes to harden the
celloidin.
7. Stain as desired.
15. Sections may be transferred from one solution to another with a bent glass rod (as in
frozen sections), but because they are thicker, they may be handled by means of forceps
instead.
Cellulose nitrate (celloidin) is soluble in absolute alcohol, and will be removed if absolute
alcohol is used in the final dehydration prior to clearing of stained sections.
Instead, sections treated with 95% alcohol may be transferred to a mixture of equal parts
of chloroform, absolute alcohol and xylene (C.A.X,) then treated with xylene and mounted
in Xam.
RE-STAINING OF OLD SECTIONS
Old, bleached or faded sections may be re-stained: the slide is usually immersed in xylene
for 24 hours, or gently heated until the mounting medium begins to bubble.
The coverslip may then be removed by lifting it with a dissecting needle.
The section is placed in xylene for up to 24 hours to remove the remaining balsam and
then brought down to water.
It is placed in a 0.5 potassium permanganate solution for 5-10 minutes, rinsed in tap
water and subsequently immersed in 5% oxalic acid for 5 minutes or until the section is
decolorized.
After washing it again in running tap water for another 5 minutes, the section may then be
re-stained with the appropriate staining technique
