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Anatomy Atlases: Atlas of Microscopic Anatomy: Appendix III: Methods of Fixation and Staining Atlas of Microscopic Anatomy

Appendix III: Methods of Fixation and Staining

Ronald A. Bergman, Ph.D., Adel K. Afifi, M.D., Paul M. Heidger, Jr., Ph.D.
Peer Review Status: Externally Peer Reviewed

The following alphabetical listing and brief comments pertain to fixatives and stains that have been used on the cells and tissues photographed for this atlas. Detailed procedures for their use can be found in the books cited in the reference section.

Aceto-Orcein Method

A staining method for sex chromatin bodies in smears of oral mucosa. This technique has two advantages over other methods for nuclear sexing: both the smear and dye are easy to prepare, and the method is just as accurate as other methods but much more rapid. Nuclei stain reddish purple. See Plate 2.

Acetylcholinesterase Histochemical Method (Seligman)

A method useful for demonstrating enzymatic activity by light and electron microscopy using osmiophilic organic compounds as substrates. Enzyme sites appear black. See Plate 73.


See Halmi's Aldehyde-Fuchsin Trichrome Stain.

Alcohol, Ethyl

Among the earliest fixing solutions used by histologists, it is not frequently used as a primary fixative except in Best's carmine method, periodic acid -Schiff reaction, and Ranson's method. Alcohol (70 to 100 per cent) is valuable for the preservation of the cellular polysaccharide glycogen for routine lightmicroscopic study. Alcohol fixation can produce distortions and shrinkage unless used at a low temperature.

Best's Carmine Stain

The rationale for this stain for glycogen is unknown. The carmine is used in an aqueous solution with potassium carbonate and potassium chloride. Glycogen stains bright red. A nuclear stain such as hernatoxylin is commonly used with Best's carmine. The periodic acid-Schiff method is another stain for glycogen and other polysaccharides. See Plate 8.

Bielchowsky's Method

This method consists of silver salt impregnation of blocks of tissue and the subsequent reduction of the silver. As a result, there is silver impregnation of neurofibrils, axons, and dendrites. Other structures, such as connective tissue fibers and neuroglia, may be impregnated. This method is valuable for the study of motor and sensory nerve endings. Nerve fibers appear black. See Plate 120.

Bodian's Silver Method

This is a silver method for the impregnation of nerve fibers in paraffin sections. It was developed by Bodian in 1937. Sections are saturated with an aqueous solution of protargol in the presence of metallic copper. The colloidal silver proteinate is then reduced in hydroquinone, and the silver is replaced by gold chloride (toning with gold). Excess silver is removed by sodium thiosulfate. This method gives uniform, sharp, and specific staining of neural elements, including axons, neurofibrils, and synaptic end feet. Axons appear black or deep blue. See Plates 113 and 118.

Bouin's Fluid

A commonly used fixative composed of a saturated aqueous solution of picric acid (75 ml), formalin (25 ml), and glacial acetic acid (5 ml). Almost any stain can be used after fixation in Bouin's. The picric acid also enhances cellular and tissue staining reactions. This fixative was described by Bouin in 1897.

Brilliant Cresyl Blue Stain

A basic dye of the oxazine group used chiefly for staining blood to demonstrate platelets and reticulocytes. The dye has a strong affinity for nucleic acid. See Plate 55.

Cajal's Gold Sublimate Method

A delicate technique developed by Cajal in 1913 to demonstrate neuroglia astrocytes, using mercuric bichloride and gold chloride. This method is considered to be the first highly selective neuroglial stain to be developed. It is usually used on frozen sections of material fixed in formalin-ammonium bromide mixture. The chemicals and water used must be of exceptional purity for best results. Protoplasmic and fibrous astrocytes appear reddish purple to near black. See Plate 129.


This dye is of great historic interest, having been used in microscopic work in the eighteenth century, considerably before the days of modern histology. It is still of great use for staining embryos, small animals, and large blocks of tissue in toto and as a specific stain for glycogen and mucus. The active principle is carminic acid, which is extracted from cochineal (dried female insects, Coccus cacti). See Plates 8 and 205.


A 37 per cent solution of formaldehyde. Formaldehyde is a gas soluble to about 55 per cent in water. Solutions used as fixatives are prepared in terms of the percentage of formalin, not formaldehyde. The commonly used 10 per cent formalin (formol) is 10 ml of formalin and 90 ml of water. Formalin is also used with other reagents (as in Helley's, Bouin's, and Regaud's fluids). Because formalin is a strong reducing agent, it is mixed with other ingredients only immediately before use.

Formic Acid

A recommended fixative for Ranvier's gold chloride method, as a substitute for undiluted fresh lemon juice.

Gallocyanin Stain

This method was popularized by Einarson in 1932. It utilizes a basic oxazine dye (gallocyanin) in combination with chrome alum. It was considered by some to be a specific stain for nucleic acids. Nuclear DNA and RNA and cytoplasmic RNA stain blue. Acidic mucopolysaccharides may also be stained. Compared with other basic dyes, gallocyanin binds strongly with nucleic acids. See Plate 90.

Giemsa Stain

This is a widely used stain for blood, spleen, and bone marrow cells, as well as for the identification of protozoan parasites. The stain is composed of methylene blue eosinate, azure A eosinate, azure B eosinate, and methylene blue chloride. The stain was used here because it is technically simple to use and because of its affinity for nuclei and chromosomes. The stained nuclei and chromosomes may vary in color from reddish-blue to purple. See Plate 3.

Glees' Method

This silver method was developed by Glees in 1946 for the study of normal and degenerating synaptic boutons. It also demonstrates neurofibrils. Although Glees' method has produced valuable results in the hands of experienced investigators, it has not come into general use. One reason for this is that this method has so far been effective only in certain regions of the central nervous system. A factor limiting its usefulness is the concomitant impregnation of normal axons and axon terminals, which obscure degenerating axons and terminals. Nerve fibers appear black. See Plates 89 and 91.


This five-carbon dialdehyde of relatively simple structure is the most commonly used fixative in electron microscopy, and it is becoming increasingly useful for light microscopy of thin sections of plasticembedded tissues. It gives superb images of cellular structure and can be used as a fixative in histochemical enzyme localization by light and electron microscopy. It is used as a 1 to 6.5 per cent solution in isotonic buffer at 1 to 4°C. Glutaraldehyde and osmium tetroxide are considered the finest fixatives available to the histologist. One of the advantages of glutaraldehyde compared with osmium is that it does not "stain" or alter the staining characteristics of the tissue during fixation. Glutaraldehyde and osmium tetroxide are sequentially used, in that order, in preparing tissue for electron microscopy. One-micrometer sections of plastic-em bedded tissue permit very high resolution light microscopy. Ultrathin sections of the same material can also be used for electron microscopy.

Golgi-Cox Method

Golgi silver methods for nerve cells depend upon preliminary fixation in a potassium dichromate solution. The silver is selective, tending to impregnate a few nerve cells completely, which then become blackened when the silver is reduced. Although these methods do not reveal details of the internal structure of nerve cells, they do provide, very importantly, a unique view of the entire cell and its processes. They also demonstrate specific details of non-nervous cells and tissue components such as the parietal cells of the stomach and bile canaliculi of the liver. The Golgi-Cox method is one of the simplest of the complex and time-consuming Golgi methods for demonstrating the relations of dendrites and axons to the nerve cell body. Cell bodies and processes are stained black on a light yellowish or colorless background. Blood vessels may also be impregnated. See Plate 87.

Gomori's Aldehyde Fuchsin Stain

This is a valuable stain that is compatible with a number of rather striking staining procedures. Aldehyde fuchsin has a poorly understood affinity for elastic tissue, beta granules in pancreatic islets, neurosecretory material, mast cell granules, and beta cells in the pituitary. The principal ingredients are basic fuchsin and paraldehyde. Several counterstains may be used. A counterstain is one that enhances the appearance of another primary stain but, in actual practice, usually provides additional specific information. Hyaline cartilage, elastic fibers, mucin, mast cell granules, and beta cells stain purple. Other tissues are stained according to the counterstain used. See Plate 47.

Gomori's Chrome Alum Hematoxylin Stain

Primary fixation should be with Bouin's or Helly's fluids, although secondary treatment by these fluids of formalin-fixed material is satisfactory. This method uses chrome alum hematoxylin stain with phloxine as a counterstain. Basophils stain blue and acidophils appear red. In the posterior pituitary gland, neurosecretory substances stain deep blue. See Plate 116.

Halmi's Aldehyde-Fuchsin Trichrome Stain

Proposed originally by Halmi (1952) for use in differentiating subpopulations of basophils in the hypophysis, this method utilizes the sequential staining of Bouin's or SUSA-fixed tissue (Romeis, 1948) by (a) immersion in aldehyde fuchsin, (b) nuclear staining with Ehrlich's hematoxylin, and (c) a rapid, single-step counterstain using light green, orange G, and Chromotrope 2R (optional) dissolved in a phosphotungstic-acetic acid mixture. The stain has subsequently found application as a general-purpose "tri-chrome" stain in a variety of tissues. Collagenous fibers stain green; nuclei, blue; nucleoli, red; erythrocytes, orange; colloid, pink; and cytoplasm, grey. See Plate 280.

Helly's Fluid

Also known as Zenker-formol, this is a modification of Zenker's fluid in which formalin is substituted for acetic acid. The concentration of formalin used varies between 5 and 10 per cent. This is an excellent cytological and tissue fixative.

H. & E. (Hematoxylin and Eosin Stain)

This is the most widely used and important general purpose stain combination. It may be used after any fixation except osmium tetroxide. Nuclear stain is the basic hematoxylin, whereas the cytoplasmic counterstain is eosin. Nuclear heterochromatin stains blue and the cytoplasm of cells rich in ribonucleoprotein also stains blue. The cytoplasm of cells with minimal amounts of ribonucleoprotein tends to be lavender in color, whereas the mature red blood cell and muscle contractile protein, which are devoid of RNA, stain red. Although it is an esthetically pleasing combination and is widely used, it is limited in its ability to differentiate cytoplasmic organelles and many other tissue components. See Plate 14.

lmmunocytochemistry, ABC Technique for VIP

Following perfusion with a picric acid- paraformaidehyde fixative, tissues are removed ' washed in sucrose-P04 buffer solution, and sectioned with a freezing microtome.. Tissue sections are processed for vasoactive intestinal polypeptide (VIP) immunohistochemistry using an adaptation by Jean Y. Jew, M.D., of the ABC method (Hsu, et al, J Histochem Cytochem 29: 577-588, 1981). Briefly, tissues were sequentially incubated in the following solutions, with washes in PO,-buffered saline between incubations: VIP antiserum, diluted 1:1000; biotinylated goat anti-rabbit IgG; avid in-bioti nylated peroxidase complex; and substrate-containing diaminobenzidine (DAB) and H201 in Tris buffer. After the DAB visualization step and wash in Tris buffer, sections are mounted on chrome alum gelatin-coated slides, dried, dehydrated through a graded ethanol and xylene series, and coverslipped with Permount. See Plate 111.

Iron Hematoxylin (Heidenhain)

This is one of the standard stains capable of excellent results after any good fixation. Tissues are stained in aqueous hematoxylin after mordanting in iron ammonium sulfate (iron alum). Many counterstains can be used. It is not specific for any structure but is particularly useful for demonstrating cell membranes, terminal bars (tight junctions or junctional complexes), secretory granules, nuclear heterochromatin, mitochondria (if preserved), and the cross striations of voluntary muscle. All these structures stain black. See Plate 16.

Kopsch's Method

This is one of several methods used to demonstrate the Golgi apparatus. Small pieces of tissue are immersed in 2 per cent osmic acid for 8 to 16 days. The Golgi apparatus stains black. If stained, mitochondria and ground substance appear reddish brown. Several modifications of this method have been developed. See Plate 7.

Luxol Fast Blue-PAS Method

This method, utilizing luxol fast blue, has become popular for staining myelin, because it can be used with other stains, allowing combinations that are not possible with the older hernatoxylin methods. Myelin nerve sheaths stain blue-green. PAS-positive substances stain pink to violet. The method consists of staining in 0.1 per cent alcoholic solution of luxol fast blue, differentiation in lithium carbonate, and counterstaining with periodic acid-Schiff. The method was introduced by Klüver and Barrera in 1953.

Mallory's Connective Tissue Stain

This is one of the most beautiful and widely used of all stains. Several modifications of the original method have been developed. Basic ingredients are acid fuchsin, aniline blue, orange G, and phosphotungstic acid. Collagen and reticular fibers stain blue; elastic fibers, yellow or pink; nuclei, fibrin, and neuroglial fibrils, red. Heidenhain's azan stain (follows) uses azocarmine instead of acid fuchsin and is a widely used substitute. See Plate 18.

Mallory-Azan (Heidenhain's Azan) Stain

This connective tissue stain is a modification of Mallory's original connective tissue stain, in which azocarmine is used along with the aniline blue-orange G mixture. Collagen and basophil granules stain 502 blue; muscle and acidophil granules, orange to red; and nuclei and cytoplasm, red. Elastic fibers are unstained or, if stained, yellow or pink. Mallory-azan is an extremely useful and beautiful stain combination. Heidenhain was the first histologist to use the azan modification of Mallory's stain. See Plate 295.

Marchi's Method and Modifications (Swank and Davenport)

Normal adult myelin contains no hydrophobic neutral lipids (triglycerides and cholesterol esters), whereas degenerating myelin does. All ethylenic (double) bonds, such as those found in fatty acids of all lipids, will reduce osmium tetroxide to the lower oxides and black metallic osmium. They cannot do this, however, if previously or simultaneously oxidized. Thus, if degenerating (hydrophobic) and normal (hydrophilic) myelin lipids are exposed to aqueous potassium chlorate, only normal myelin ethylene bonds will be attacked. Unaffected bonds of degenerating myelin are thus the only ones still free to reduce osmium tetroxide. Nervous tissue in which degenerating myelin is present is fixed for 2 to 3 days in either formalin or a magnesium sulfate-potassium clichromate mixture followed by formalin. The tissue is then placed for 8 to 10 days in a solution containing potassium chlorate, osmium tetroxide, formaldehyde, and glacial acetic acid. After a thorough wash, the tissue is embedded in celloidin (nitrocellulose), sectioned, and mounted. Degenerating myelin and neutral lipids elsewhere will be black. Normal myelin is unstained. See Plate 327.

Masson's Trichrome Method

This method was first described by Pierre Masson in 1951. Although Bouin's is the recommended fixative, Orth's (formalin-Müller), Zenker's, or 10 per cent formalin in alcohol may be used. Dyes and solutions used are hematoxylin, acid fuchsin, phosphotungstic acid, and light green. Several modifications of this method are available. Nuclei stain dark blue; cytoplasm and neuroglial fibers, red; collagen, green. See Plate 282.

Methylene Blue and Erythrocin

This is a frequently used combination to stain tissues. Methylene blue stains basophilic constituents of the tissue, whereas erythrocin stains the acidophilic elements. See Plate 1.

Modified Aldehyde Fuchsin Stain

Halmi's modification of Gomori's method utilizes aldehyde fuchsin with light green or orange G as the counterstain. Nuclei can be stained with celestine blue or haemalum (alum hematoxylin). Depending upon the concentration of organelles found in different cell types, they may appear yellow, purple, or green. Collagen and the basement membrane are green, and mast cells, elastic fibers, and goblet cell mucus are purple. See Plate 20.

Müller's Fluid

Potassium clichromate (2.0 to 2.5 per cent)-sodium sulfate (1 per cent) mixture in distilled water. Formerly, this was much used for prolonged fixation and mordanting of nervous tissue. It has now been largely replaced by Orth's fluid, which is composed of potassium clichromate (2.0 to 2.5 per cent) and formalin (10 per cent) in distilled water. The sodium sulfate is omitted. Fixatives of this type do not store well and must be prepared immediately before use.

Nassar-Shanklin Method

A good silver method for staining neuroglia in formalin sections. Formalin-fixed paraffin sections are impregnated with silver diaminohydroxicle or with strong Hortega silver carbonate dissolved with strong ammonia after sensitizing with sodium sulfite. Microglia, oligodendroglia, and fibrous and protoplasmic astrocytes can be successfully impregnated by this method. Sensitization with sodium sulfite is essential for good results. See Plates 128 and 130.

Osmium Tetroxide (OsO4)

Commonly but incorrectly referred to as osmic acid, this is considered an excellent fixative for most cytological morphology. It is used either as a primary fixative or secondarily after alclehyde fixation. it cannot be used for histochemical studies. Osmium blackens lipids and stains the Golgi apparatus in light microscopic preparations. A severe limitation of osmium-fixed material is that most routine and special stains cannot be used. Its primary use is in electron microscopy. See Plate 9.

Pal-Weigert Method

First described in 1887, this is a modification of the Weigert method for myelin sheaths in which the differentiation between the myelinated fibers and the surrounding tissues may be carried to a greater degree than in the original method. Originally Müller's fluid was used as the fixative; later, formalin was used. Several modifications of the original Pal-Weigert method are available. Normal myelin sheaths are stained deep blue. Degenerative myelin does not stain (see also Weigert's method). See Plate 324.

Periodic Acid-Schiff (PAS) Method

This method is principally used to demonstrate structures rich in polysaccharides (glycogen), mucopolysaccharides (e.g., ground substance of connective tissues, basement membrane, and mucus), glycoproteins (thyroglobulin), and glycolipids. This method depends upon the selective oxidation by periodic acid of 1,2-glycols and 2,2-amino alcohols to aldehydes. The aldehydes are then detected by the Schiff reagent, which stains them reddish purple. Schiff reagent is formed by the reaction of basic fuchsin with sulfurous acid. See Plate 78.

Phosphotungstic Acid Hematoxylin (PTAH)

This stain was developed by Mallory. it is an ideal stain for the demonstration of astroglial fibers, which stain blue as do striated muscle fibers and mitochondria. Collagen, reticular fibers, and ground substance of bone and cartilage stain in varying shades of yellow to brownish red. Coarse elastic fibers stain purple. Nuclei are blue. See Plate 66.

Pinkus'Acid Orcein-Giemsa Method

This is a modification of the original Unna-Taenzer procedure by Pinkus in 1944, which has been further modified and simplified by Pinkus and Hunter. A good method to demonstrate connective tissue elements. Material is fixed in 10 per cent formalin, formol-alcohol, or absolute alcohol. Paraffin sections are used. Nuclei stain deep blue; cytoplasm, light blue; collagen, rose pink; and elastic fibers, dark brown. See Plate 133.

Ranson's Method

This is a pyridine silver method described by Ranson in 1911 for impregnating unmyelinated nerve fibers. The largest myelinated axis cylinders are usually yellow, surrounded by a colorless ring of myelin, and there is a gradation to black in the small myelinated fibers. Unmyelinated fibers are typically dark brown or black. Nerve cells are yellow to brown, with dark brown to black neurofibrils. End bulbs, when stained, appear black or as small black rings. See Plate 95.

Ranvier's Gold Chloride Method

This method was described by Ranvier in 1880. It demonstrates nerve endings in muscle. Nerve fibers are variably stained red to purple-black. The tissue is fixed in formic acid or undiluted fresh lemon juice before immersion in an aqueous solution of gold chloride. See Plate 119.

Regaud's Method

Described in 1910 by Regaud, this is a modification of Heidenhain's iron hematoxylin method for mitochondria, which consists of prolonged mordanting of tissues in potassium dichromate. It stains mitochondria blue-black. It is the most permanent and the simplest of all mitochondrial stains. See Plate 6.

Rossman's Fluid

A fixative capable of preserving cellular glycogen. This fixative consists of absolute ethyl alcohol (90 ml) saturated with picric acid (approximately 9 per cent) and neutral formalin (10 ml). After overnight fixation, the excess picric acid is removed by rinsing in 95 per cent alcohol for several days.

Silver Diaminohydroxide Method

See Nassar-Shanklin method.

Susa Fixative

Susa fixative, developed by Heidenhain (a nineteenth-century German histologist), derives its name from the initial letters of sublimate and saure, key ingredients of the fixative. The solution consists of mercuric chloride (sublimate) saturated in 0.6 per cent NaCl; formalin; and two acids (saure), namely, trichloracetic and glacial acetic acid, all in aqueous solution. It is considered a good general- purpose fixative, with rapid penetration.

Tetrazolium Method

Tetrazoliurn salts have low oxidation-reduction potentials and are thus capable of intercepting electrons in many biological oxidation-reduction reactions, including those facilitated enzymatically by dehydrogenases. The reduced, colored end product (blue diformazan) of these reactions is insoluble and thereby demonstrates the sites of such activity. Localization of the site of enzymatic activity is an important contribution to our understanding of cellular function. The methods of Nachlas, Walker, and Seligman are outstanding examples of this technique. See Plate 72.

Toluldine Blue Stain

Toluidine blue 0 is a basic dye of the thiazine series closely related to methylene blue. In routine preparations, toluidine blue stains nucleic and cytoplasmic ribonucleic acid ortho- or normochromatically (blue) and cartilage matrix and mast cell granules metachromatically (reddish purple). This is thus a metachromatic stain and is very useful for 1 µn sections of plastic-embedded tissue. See Plates 28 and 137.

Van Gehuchten's Fluid

This is a modified mixture of Carnoy's fluid containing absolute alcohol, chloroform, and glacial acetic acid. A very fast-acting fixative.

Van Gieson's Stain

This is a good stain for connective tissue. It consists of staining by acid fuchsin and picric acid. The stain fades in time and is therefore not used as frequently as the Mallory connective tissue stain. Collagen stains bright red; muscle and cytoplasm, yellow; and nuclei, blue to black. Delicate collagen fibrils and reticulum stain very faintly or not at all. It is valuable as a counterstain in techniques that stain elastic fibers a contrasting color (e.g., the methods of Verhoeff and Weigert). Combined with Verhoeff's stain, it is one of the most valuable stains for the study of blood vessels. See Plate 67.

Verhoeff and Van Gieson's Stain

See Van Gieson's stain.

Weigert's Elastic Fiber Stain

This technique is an excellent method for staining elastic fibers. The procedure consists of staining with a ferric chloride-hematoxylin mixture. Several counterstains may be used. Elastic fibers stain blue-black to black. Other tissue elements depend upon the counterstain used. Commonly used counterstains are phloxine or van Gieson's. See Plate 152.

Weigert's Method

This method was described by Weigert in 1885 for normal myelin sheaths. It is used principally on formol-fixed tissue of the central nervous system to demonstrate fiber tracts and to show the arrangement of gray and white matter. The basic procedure requires the use of a mordant, potassium dichromate, a hematoxylin stain, and a differentiating fluid. Normal but not degenerating or degenerated myelin sheaths stain black. See Plate 319.

Weil's Method

Weil's hernatoxylin method for myelin does not depend upon a specific method of fixation, although morclanting of formalin-fixed material by Weigert's potassium dichromate and chromium fluoride is advisable. The stain is composed of equal parts of iron alum (4 per cent) and hernatoxylin (1 per cent). Myelin sheaths stain black or dark gray.

Wilder's Method

This is a silver impregnation method for reticular fibers. Reticular fibers stain black; collagen, rose color; and other tissue elements depend upon the counterstain used. See Plate 37.

Wright's Stain

This is a differential stain for blood cells named after the American pathologist James Wright (11869-1928), who described this method in 1902. The main constituents of his stain are methylene blue and eosin. Erythrocytes bind eosin and appear red or pink; nuclei, deep blue or purple; basophilic granules, deep purple; eosinophilic granules, red to red-orange; neutrophilic granules, reddish-brown to lilac; platelets, violet to purple; and lymphocyte cytoplasm, pale blue. See Plates 52, 53, and 54.

Zenker's Fluid

This excellent fixative is a mixture of potassium clichromate and bichloride of mercury to which glacial acetic acid is added. A mercury precipitate is left in the tissues and is troublesome to the inexperienced microscopist unless removed by an iodine solution.

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