10 The hematoxylins and eosin
The hematoxylin and eosin stain (H&E) is the most widely used histological stain. Its popularity is based on its comparative simplicity and ability to demonstrate clearly an enormous number of different tissue structures. Hematoxylin can be prepared in numerous ways and has a widespread applicability to tissues from different sites. Essentially, the hematoxylin component stains the cell nuclei blue-black, showing good intranuclear detail, while the eosin stains cell cytoplasm and most connective tissue fibers in varying shades and intensities of pink, orange, and red. While automated staining instruments and commercially prepared hematoxylin and eosin solutions are more commonly used in today’s laboratories for routine staining, students of histological techniques should have a basic knowledge of the dyes and preparation techniques in order to troubleshoot and/or modify procedures for specialized use. It should be noted that hematoxylin has additional uses beyond just the hematoxylin and eosin combination.
Eosin is the most suitable stain to combine with an alum hematoxylin to demonstrate the general histological architecture of a tissue. Its particular value is its ability, with proper differentiation, to distinguish between the cytoplasm of different types of cell, and between the different types of connective tissue fibers and matrices, by staining them differing shades of red and pink.
The eosins are xanthene dyes and the following types are easily obtainable commercially: eosin Y (eosin yellowish, eosin water-soluble) C.I. No. 45380 (C.I. Acid Red 87); ethyl eosin (eosin S, eosin alcohol-soluble) C.I. No. 45386 (C.I. Solvent Red 45); eosin B (eosin bluish, erythrosin B) C.I. No. 45400 (C.I. Acid Red 91).
Of these, eosin Y is much the most widely used, and despite its synonym it is also satisfactorily soluble in alcohol; it is sometimes sold as ‘water and alcohol soluble’. As a cytoplasmic stain, it is usually used as a 0.5 or 1.0% solution in distilled water, with a crystal of thymol added to inhibit the growth of fungi. The addition of a little acetic acid (0.5 ml to 1000 ml stain) is said to sharpen the staining. Differentiation of the eosin staining occurs in the subsequent tap water wash, and a little further differentiation occurs during the dehydration through the alcohols. The intensity of eosin staining, and the degree of differentiation required, is largely a matter of individual taste. Suitable photomicrographs of H&E-stained tissues are easier to obtain when the eosin staining is intense and the differentiation slight (at least double the routine staining time is advisable). Ethyl eosin and eosin B are now rarely used, although occasional old methods specify their use: for example, the Harris stain for Negri bodies. Alternative red dyes have been suggested as substitutes for eosin, such as phloxine, Biebrich scarlet, etc.; however, although these substitutes often give a more intense red color to the tissues, they are rarely as amenable to subtle differentiation as eosin and are generally less valuable.
Under certain circumstances eosin staining is intense and difficulty may be experienced in obtaining adequate differentiation; this may occur after mercuric fixation. Over-differentiation of the eosin may be continued until only the red blood cells and granules of eosinophil polymorph are stained red. This is, occasionally, used to facilitate the location and identification of eosinophils. Combining eosin Y and phloxine B (10 ml 1% phloxine B, 100 ml 1% eosin Y, 780 ml 95% alcohol, 4 ml glacial acetic acid) produces a cytoplasmic stain, which more dramatically demonstrates various tissue components. Muscle is clearly differentiated from collagen, and red cells stain bright red. According to Luna (1992), eosin dye content should be 88% and not contain sodium sulfate (sometimes used as filler). When using this dye in solution, a fine granular precipitate forms, and the staining of the cytoplasm will be poor.
Hematoxylin is extracted from the heartwood (‘logwood’) of the tree Hematoxylon campechianum that originated in the Mexican State of Campeche, but which is now mainly cultivated in the West Indies. The hematoxylin is extracted from logwood with hot water, and then precipitated out from the aqueous solution using urea (see prior editions). Hematoxylin itself is not a stain. The major oxidization product is hematein, a natural dye that is responsible for the color properties. Hematein can be produced from hematoxylin in two ways.
This is a slow process, sometimes taking as long as 3–4 months, but the resultant solution seems to retain its staining ability for a long time. Ehrlich’s and Delafield’s hematoxylin solutions are examples of naturally ripened hematoxylins.
Examples are sodium iodate (e.g. Mayer’s hematoxylin) or mercuric oxide (e.g. Harris’s hematoxylin). The use of chemical oxidizing agents converts the hematoxylin to hematein almost instantaneously, so these hematoxylin solutions are ready for use immediately after preparation. In general, they have a shorter useful life than the naturally oxidized hematoxylins, probably because the continuing oxidation process in air and light eventually destroys much of the hematein, converting it to a colorless compound. Hematein is anionic, having a poor affinity for tissue, and is inadequate as a nuclear stain without the presence of a mordant. The mordant/metal cation confers a net positive charge to the dye-mordant complex and enables it to bind to anionic tissue sites, such as nuclear chromatin. The type of mordant used influences strongly the type of tissue components stained and their final color. The most useful mordants for hematoxylin are salts of aluminum, iron, and tungsten, although hematoxylin solutions using lead as a mordant are occasionally used (for example in the demonstration of argyrophil cells). Most mordants are incorporated into the hematoxylin staining solutions, although certain hematoxylin stains required the tissue section to be pre-treated with the mordant before staining; such as Heidenhain’s iron hematoxylin. Hematoxylin solutions can be arbitrarily classified according to which mordant is used:
This group comprises most of those that are used routinely in the hematoxylin and eosin stain, and produce good nuclear staining. The mordant is aluminum, usually in the form of ‘potash alum’ (aluminum potassium sulfate) or ‘ammonium alum’ (aluminum ammonium sulfate). All stain the nuclei a red color, which is converted to the familiar blue-black when the section is washed in a weak alkali solution. Tap water is usually alkaline enough to produce this color change, but occasionally alkaline solutions such as saturated lithium carbonate, 0.05% ammonia in distilled water, or Scott’s tap water substitute (see Appendix III) are necessary. This procedure is known as ‘blueing’.
The alum hematoxylins can be used regressively, meaning that the section is over-stained and then differentiated in acid alcohol, followed by ‘blueing’, or progressively, i.e. stained for a predetermined time to stain the nuclei adequately but leave the background tissue relatively unstained. The times for hematoxylin staining and for satisfactory differentiation will vary according to the type and age of alum hematoxylin used, the type of tissue, and the personal preference of the pathologist. For routine hematoxylin and eosin staining of tissues, the most commonly used hematoxylins are Ehrlich’s, Mayer’s, Harris’s, Gill’s, Cole’s, and Delafield’s. Carazzi’s hematoxylin is occasionally used, particularly for urgent frozen sections.
Ehrlich’s hematoxylin (Ehrlich 1886)
This is a naturally ripening alum hematoxylin which takes about 2 months to ripen; the ripening time can be shortened somewhat by placing the unstoppered bottle in a warm sunny place such as a window-ledge, and is shorter in the summer than in winter. Once satisfactorily ripened, this hematoxylin solution will last in bulk for years, and retains its staining ability in a Coplin jar for some months. Ehrlich’s hematoxylin, as well as being an excellent nuclear stain, also stains mucins including the mucopolysaccharides of cartilage; it is recommended for the staining of bone and cartilage (Chapter 16).
|Absolute alcohol||100 ml|
|Distilled water||100 ml|
|Glacial acetic acid||10 ml|
|Potassium alum||15 g approx.|
The hematoxylin is dissolved in the alcohol, and the other chemicals are added. Glycerin is added to slow the oxidation process and prolong the hematoxylin shelf life. Natural ripening in sunlight takes about 2 months, but in an emergency the stain can be chemically ripened by the addition of sodium iodate, using 50 mg for every gram of hematoxylin; this will inevitably shorten the bench life of the stain. By definition this chemically oxidized variant is not a true Ehrlich’s hematoxylin and will not have the same longevity as naturally oxidized Ehrlich’s hematoxylin. One should always filter before use.
Ehrlich’s hematoxylin, being a strong hematoxylin solution, stains nuclei intensely and crisply, and stained sections fade much more slowly than those stained with other alum hematoxylins. It is particularly useful for staining sections from tissues that have been exposed to acid. It is suitable for tissues that have been subjected to acid decalcification or, more valuably, tissues that have been stored for a long period in formalin fixatives which have gradually become acidic over the storage period, or in acid fixatives such as Bouin’s fixative. Ehrlich’s hematoxylin is not ideal for frozen sections.
Delafield’s hematoxylin (Delafield 1885)
|95% alcohol||125 ml|
|Saturated aqueous ammonium alum (15 g/100 ml)||400 ml|
The hematoxylin is dissolved in 25 ml of alcohol, and then added to the alum solution. This mixture is allowed to stand in light and air for 5 days, then filtered, and to it are added the glycerin and a further 100 ml of 95% alcohol. The stain is allowed to stand exposed to light and air for about 3–4 months or until sufficiently dark in color, then is filtered and stored. Filter before use.
Mayer’s hematoxylin (Mayer 1903)
This alum hematoxylin is chemically ripened with sodium iodate. It can be used as a regressive stain like any alum hematoxylin. However, it is also useful as a progressive stain, particularly in situations where a nuclear counterstain is needed to emphasize a cytoplasmic component which has been demonstrated, by a special stain, and where the acid-alcohol differentiation might destroy or de-color the stained cytoplasmic component. It is used as a nuclear counterstain in the demonstration of glycogen, in various enzyme histochemical techniques, and in many others. The stain is applied for a short time (usually 5–10 minutes), until the nuclei are stained, and is then ‘blued’ without any differentiation.
|Distilled water||1000 ml|
|Potassium or ammonium alum||50 g|
|Sodium iodate||0.2 g|
|Citric acid||1 g|
|Chloral hydrate SLR||50 g or|
|Chloral hydrate AR||30 g|
The hematoxylin, potassium alum, and sodium iodate are dissolved in the distilled water by warming and stirring, or by allowing to stand at room temperature overnight. The chloral hydrate and citric acid are added, and the mixture is boiled for 5 minutes, then cooled and filtered. If higher-purity chloral hydrate AR grade is used, the amount may be reduced, as shown above. The stain is ready for use immediately. Filter before use.
Harris’s hematoxylin (Harris 1900)
This alum hematoxylin was traditionally chemically ripened with mercuric oxide. As mercuric oxide is highly toxic, environmentally unfriendly, and has detrimental and corrosive long-term effects on some automated staining machines, sodium or potassium iodate is frequently used as a substitute for oxidation. Harris is a useful general-purpose hematoxylin and gives particularly clear nuclear staining, and for this reason has been used, as a progressive stain, in diagnostic exfoliative cytology. In routine histological practice, it is generally used regressively, but can be useful when used progressively. When using Harris’s hematoxylin as a progressive stain, an acetic acid-alcohol rinse provides a more controllable method in removing excess stain from tissue components and the glass slide. The traditional hydrochloric acid-alcohol acts quickly and indiscriminately, is more difficult to control, and can result in a light nuclear stain. A 5–10% solution of acetic acid, in 70–95% alcohol, detaches dye molecules from the cytoplasm/nucleoplasm while keeping nucleic acid complexes intact (Feldman & Dapson 1985).
|Absolute alcohol||25 ml|
|Potassium alum||50 g|
|Distilled water||500 ml|
|Mercuric oxide||1.25 g or|
|Sodium iodate||0.5 g|
|Glacial acetic acid||20 ml|
The hematoxylin is dissolved in the absolute alcohol, and is then added to the alum, which has previously been dissolved in the warm distilled water in a 2-liter flask. The mixture is rapidly brought to the boil and the mercuric oxide or sodium iodate is then slowly and carefully added. Plunging the flask into cold water or into a sink containing chipped ice rapidly cools the stain. When the solution is cold, the acetic acid is added, and the stain is ready for immediate use. The glacial acetic acid is optional but its inclusion gives more precise and selective staining of nuclei.
As with most of the chemically ripened alum hematoxylins, the quality of the nuclear staining begins to deteriorate after a few months. This deterioration is marked by the formation of a precipitate in the stored stain. At this stage, the stain should be filtered before use, and the staining time may need to be increased. For the best results, it is wise to prepare a fresh batch of stain every month, although this may be uneconomical unless only small quantities are prepared each time.
Cole’s hematoxylin (Cole 1943)
|Saturated aqueous potassium alum||700 ml|
|1% iodine in 95% alcohol||50 ml|
|Distilled water||250 ml|
The hematoxylin is dissolved in warm distilled water and mixed with the iodine solution. The alum solution is added, and the mixture brought to the boil, then cooled quickly and filtered. The solution is ready for immediate use, but may need filtering after storage, for the same reason as described above for Harris’s hematoxylin. Filter before use.
Carazzi’s hematoxylin (Carazzi 1911)
|Preparation of solution|
|Potassium alum||25 g|
|Distilled water||400 ml|
|Potassium iodate||0.1 g|
The hematoxylin is dissolved in the glycerol, and the alum is dissolved in most of the water overnight. The alum solution is added slowly to the hematoxylin solution, mixing well after each addition. The potassium iodate is dissolved in the rest of the water with gentle warming and is then added to the hematoxylin-alum-glycerol mixture. The final staining solution is mixed well and is then ready for immediate use; it remains usable for about 6 months. Care must be taken in preparing the hematoxylin to avoid over-oxidation; it is safer if heat is not used to dissolve the reagents. Filter before use.
Like Mayer’s hematoxylin, Carazzi’s hematoxylin may be used as a progressive nuclear counterstain using a short staining time, followed by blueing in tap water. It is particularly suitable, since it is a pale and precise nuclear stain and does not stain any of the cytoplasmic components.
Gill’s hematoxylin (Gill et al. 1974 modified)
|Sodium iodate||0.2 g|
|Aluminum sulfate||17.6 g|
|Distilled water||750 ml|
|Ethylene glycol (ethandiol)||250 ml|
|Glacial acetic acid||20 ml|
The distilled water and ethylene glycol are mixed, and then the hematoxylin is added and dissolved. The ethylene glycol is an excellent solvent for hematoxylin and it prevents the formation of surface precipitates (Carson 1997). Sodium iodate is added for oxidation, and the aluminum sulfate mordant is then added and dissolved. Finally, the glacial acetic acid is added and stirred for 1 hour. The solution is filtered before use. Carson reported that, although the stain can be used immediately, it provides a better intensity if allowed to ripen for 1 week in a 37°C incubator. It should be noted that the popularity of Gill’s solution has made it one of the more commercially successful formulas.
Double or triple hematoxylin concentrations may be used as preferred. These are usually referred to as Gill’s I (normal), Gill’s II (double), and Gill’s III (triple), with the Gill III being the most concentrated. Gill’s hematoxylin is more frequently used for routine H&E staining than Mayer’s hematoxylin, and is more stable than Harris’s hematoxylin, as auto-oxidation is inhibited to the extent that no measurable changes occur over many months. Disadvantages associated with Gill’s hematoxylin include staining of gelatin adhesive and even the glass itself. Some mucus may also stain darkly, as compared to Harris’s, where mucus generally remains unstained, and the glass usually fails to attract the stain. Feldman and Dapson (1987) theorized that the aluminum sulfate mordant is responsible. Certain charged sites in the tissue, in the adhesive, and on the glass are masked by the Harris mordant, leaving them unavailable for staining. Gill’s mordant system fails to do that, and the sites attract the dye-mordant complex.