Microorganisms Stains

Kinyoun & Ziehl-Neelsen methods
(purpose of stain)

Detection of Mycobacterium tuberculosis and other acid-fast mycobacteria in tissue sections.

Kinyoun & Ziehl-Neelsen methods
(principle of the technique)

The waxy capsule of the acid-fast organism takes up carbol-fuchsin and resists decolorization with dilute mineral acid. Carbol-fuchsin is more soluble in the lipids of the cell wall than in acid-alcohol, but is readily removed from bacteria that lack the waxy capsule. Staining is enhanced by the phenol and alcohol, and both of these chemicals also aid in dissolving the basic fuchsin. Alcoholic rather than aqueous solutions of acid are used because more uniform decolorization is obtained with alcoholic solutions. The Carbol-fuchsin methods provide a specific way of identifying mycobacteria. These organisms are not readily demonstrated by other methods such as the Gram stain. The lipid capsule of mycobacteria is of such high molecular weight that it is waxy at room temperature, and successful penetration by the aqueous-based staining solutions used in the Gram staining procedures is prevented.

Kinyoun & Ziehl-Neelsen methods
(preferred fixative)

Although 10% NBF is preferred, others, with the exception of Carnoy solution may be used. Chloroform in Carnoy solution will make acid-fast organisms non-acid-fast.

Kinyoun & Ziehl-Neelsen methods
(preferred thickness)

Cut paraffin sections at 4 - 5 µm.

Kinyoun & Ziehl-Neelsen methods
(control tissue)

Tissue containing acid-fast organisms must be used for a control.

Kinyoun & Ziehl-Neelsen methods
(major reagents & purpose)

Carbol-fuchsin (basic fuchsin, phenol, ethanol) - demonstration of acid-fast mycobacteria.

Acid alcohol (HCl, ethanol) - differentiate individually until tissue is pale pink. \n \n Running tap water - wash out acid. \n \n Methylene blue - counterstain.

Kinyoun & Ziehl-Neelsen methods
(results)

Acid-fast mycobacteria - bright red (basic fuchsin)

Mast cells, background - light blue (methylene blue)

Kinyoun & Ziehl-Neelsen methods
(sources of error)

Acid-fast organisms have been reported to exist in tap water, so no unfiltered tap water should be used before applying the carbol-fuchsin reagent. The counterstain is critical in this procedure because overcounterstaining with methylene blue will mask any acid-fast organisms present. If the section is overstained, take it back to the acid-alcohol to remove the methylene blue, wash with water and then repeat the counterstaining step. If the acid is not washed out of the tissue before the counterstaining step the tissue will not stain. If the section is allowed to dry after the carbol-fuchsin stain is applied, a compound that is resistant to decolorization will be formed. Repeated attempts to remove this compound will result in complete decolorization of the acid-fast organisms. If the positive control contains too many acid-fast bacteria, there is a possibility of overdecoloriztion without recognizing it, and of cross-contamination through knife or solution metastasis. This method is not satisfactory for the demonstration of Mycobacterium leprae; this organism's lipid capsule is very sensitive to the alcohols and xylene used in routine acid-fast techniques, so special protective measures must be taken during steps requiring alcohol and xylene.

Fite acid-fast stain
(purpose of stain)

Detection of Mycobacterium leprae (causative organism of leprosy) in tissue sections.

Fite acid-fast stain
(principle of the technique)

The lipid capsule of the organism takes up carbol-fuchsin and resists decolorization with dilute mineral acid.

Fite acid-fast stain
(preferred fixative)

Although 10% NBF is preferred, others, with the exception of Carnoy solution may be used. Chloroform in Carnoy solution will make acid-fast organisms non-acid-fast.

Fite acid-fast stain
(preferred thickness)

Cut paraffin sections at 4 - 5 µm.

Fite acid-fast stain
(control tissue)

Tissue containing leprosy organisms must be used for a control.

Fite acid-fast stain
(major reagents & purpose)

Xylene-peanut oil - deparaffinize; peanut oil protects the waxy capsule to enhance acid-fastness.

Blot off excess oil - residual oil helps to prevent shrinkage and injury of the sections. \n \n Ziehl-Neelsen carbol-fuchsin (basic fuchsin, phenol, ethanol) - demonstration of M. leprae. This method uses much less basic fuchsin in the carbol-fuchsin solution compared to the Kinyoun method. \n \n Acid alcohol (HCl, ethanol) - differentiate individually until sections are faint pink. \n \n Wash in tap water - wash out acid. \n \n Methylene blue - counterstain. \n \n Allow sections to air dry completely then mount dry sections with synthetic resin, do not use alcohol and xylene.

Fite acid-fast stain
(results)

M. leprae & other acid-fast bacteria - bright red (basic fuchsin).

Background - light blue (methylene blue).

Fite acid-fast stain
(sources of error)

This method is not as good as the Kinyoun procedure for mycobacteria other than M. leprae. Acid fastness of the leprosy organism is enhanced when the waxy capsule is protected by the mixture of peanut oil and xylene and by the avoidance of dehydrating solutions. Even a short exposure to xylene to remove paraffin has an adverse effect on staining (paradoxically the prolonged exposure to xylene during processing does not have the same effect). The problem is one of resistance to uptake of the stain rather than retention since once the leprosy organism is stained it will resist decolorization tenaciously.

Auramine-rhodamine microwave technique
(purpose of stain)

Detection of Mycobacterium tuberculosis or other acid-fast organisms

Auramine-rhodamine microwave technique
(principle of the technique)

The exact mechanism of this stain is unknown. Both of the dyes used are basic dyes that fluoresce at short wavelengths. Both dyes used in combination yield better staining than either dye used alone.

Auramine-rhodamine microwave technique
(preferred fixative)

10% NBF is preferred. Heavy metals such as zinc may quench primary fluorescence of the specimen thus fluorescence microscopy is unsatisfactory if zinc formalin is used for fixation.

Auramine-rhodamine microwave technique
(preferred thickness)

Cut paraffin sections at 4 - 5 µm.

Auramine-rhodamine microwave technique
(microscope used)

Fluorescence microscope with a high-dry objective, a UG 1 or UG 2 exciter filter, and a colorless UV barrier filter.

Auramine-rhodamine microwave technique
(control tissue)

Tissue containing acid-fast mycobacteria must be used for control.

Auramine-rhodamine microwave technique
(major reagents & purpose)

Auramine O-Rhodamine B (80C in microwave) - demonstration of acid-fast organisms.

Acid alcohol (HCl, ethanol) - differentiate. \n \n Eriochrome black T - counterstain. \n \n Allow sections to air dry completely then mount dry sections with synthetic resin, do not use alcohol and xylene.

Auramine-rhodamine microwave technique
(results)

Acid-fast organisms - reddish-yellow fluorescence (auramine-rhodamine).

Background - black (eriochrome black).

Auramine-rhodamine microwave technique
(sources of error)

This is an extremely sensitive and highly specific method for mycobacteria; however there is an increased possibility of false positives. Slides stained with the auramine-rhodamine method can be restained with carbol-fuchsin for confirmation if the results are questionable; however, carbol-fuchsin stained slides cannot be restained with auramine-rhodamine. Although rhodamine is a fluorochrome, it can act to quench fluorescence. Therefore, reducing the concentration of rhodamine greatly intensifies the fluorescence of mycobacteria. However, a more intense fluorescence is obtained with the addition of even a small amount of rhodamine compared to auramine alone.

Brown-Hopps modification of the Gram stain
(purpose of stain)

Demonstration of Gram-negative and Gram-positive bacteria in tissue.

Brown-Hopps modification of the Gram stain
(principle of the technique)

Crystal violet is applied first, followed by an iodine mordant forming a dye lake. At this point, both Gram-negative and Gram-positive organisms are stained. The cell walls of Gram-positive bacteria are thicker than those of Gram-negative organisms; this accounts for differences in the way that bacteria will decolorize in the next procedural step. The large crystal violet-iodine molecular complex cannot easily be washed out of the intact peptidoglycan layers of Gram-positive cells; however, it is easily removed from Gram-negative bacteria, because alcohol or acetone disrupts the outer lipoprotein layer, and the remaining thin peptidoglycan cell wall cannot retain the complex. Gram-positive cell walls will retain the crystal violet-iodine complex unless the cell walls have been damaged or disrupted for some other reason (then the organism will stain Gram-negative). The decolorization step is a relative one, and sections can be overdecolorized, removing stain from both Gram-negative and Gram-Positive organisms. After decolorization, a counterstain is applied to color the Gram-negative organisms.

Brown-Hopps modification of the Gram stain
(preferred fixative)

10% NBF

Brown-Hopps modification of the Gram stain
(preferred thickness)

Cut paraffin sections at 4 - 5 µm.

Brown-Hopps modification of the Gram stain
(control tissue)

Sections containing both Gram-positive and Gram-negative organisms should be used.

Brown-Hopps modification of the Gram stain
(major reagents & purpose)

Crystal violet - demonstrate Gram-positive bacteria

Gram iodine - mordant forms a dye lake. \n \n Acetone - decolorize crystal violet which is not adherent to bacterial cell walls. \n \n Basic fuchsin - counterstain for Gram-negative bacteria. \n \n Gallego solution (37% formalin, glacial acetic acid) - differentiates and fixes basic fuchsin. \n \n Rinse and blot, but not to dryness. \n \n Acetone - decolorize basic fuchsin. \n \n Picric acid-acetone - decolorize background tissue & counterstain. \n \n Acetone - decolorize picric acid. \n \n Acetone-xylene - decolorize & clear..

Brown-Hopps modification of the Gram stain
(results)

Gram-positive bacteria - blue (crystal violet).

Gram-negative bacteria - red (basic fuchsin). \n \n Background tissue - yellow (picric acid). \n \n Nuclei - light red (basic fuchsin).

Brown-Hopps modification of the Gram stain
(sources of error)

Brown-Hopps is the preferred modification for Gram-negative bacteria and rickettsiae, but Brown-Brenn is preferred for demonstrating Gram-positive bacteria (B&B skips the Gallego solution). The depth staining of weakly Gram-negative organisms can be intensified by increasing the concentration of basic fuchsin. The picric acid-acetone decolorizes sections better if the picric acid is nearly anhydrous. Sections should not be allowed to dry at any stage of the procedure, because drying leads to the formation of insoluble compounds that are difficult or impossible to decolorize with picric acid-acetone.

Modified diff-quik Giemsa stain
(purpose of stain)

Identification of H. pylori, a gram negative bacterium that has been shown to be the causative organism in some gastric and duodenal ulcers.

Modified diff-quik Giemsa stain
(principle of the technique)

Giemsa is the most common example of a Romanowsky type stain i.e. "neutral" dyes combining the basic dye methylene blue and the acid dye eosin. These give a wide color range when staining tissues and blood smears because of impurities present in the actual dye solution. On standing in solution (particularly at an alkaline pH) methylene blue gives rise to new substances that are metachromatic (azure A & B); however, today most commercial solutions are prepared with weighed amounts of the azures, and most omit methyl violet.

Modified diff-quik Giemsa stain
(preferred fixative)

10% NBF

Modified diff-quik Giemsa stain
(preferred thickness)

Cut paraffin sections at 4 - 5 µm.

Modified diff-quik Giemsa stain
(control tissue)

Sections containing H. pylori.

Modified diff-quik Giemsa stain
(major reagents & purpose)

Diff-Quik solution I (anionic xanthene dye buffered solution of eosin Y) - stains the cytoplasmic elements pink.

Diff-Quik solution II (cationic thiazine dye mixture of azure A and methylene blue) - stains the nuclei and bacteria blue. \n \n Acetic acid water - differentiate. \n \n Rinse quickly in distilled water & dehydrate quickly through ethanols.

Modified diff-quik Giemsa stain
(results)

H. pylori & other bacteria - dark blue (Diff-Quik solution II)

Nuclei - dark blue (Diff-Quik solution II) \n \n Cytoplasm - pink (Diff-Quik solution I)

Modified diff-quik Giemsa stain
(sources of error)

Check microscopically. Do not prolong the time in the last distilled water rinse or in the dehydrating alcohols, or it will lead to excess decolorization.

Alcian yellow-toluidine blue method
(purpose of stain)

Detection of H. pylori, a gram negative bacterium that has been shown to be the causative organism in some gastric and duodenal ulcers.

Alcian yellow-toluidine blue method
(principle of the technique)

Mucus, in and near the surface of the stomach lining, is neutral and will not stain directly with basic dyes. In both techniques, periodic acid oxidizes the hydroxyls in carbohydrates (mucus) to aldehydes. Sodium metabisulfite then attacks the aldehydes to create sulfonic acids. Mucus is now highly acidic and stains readily and nearly irreversibly with Alcian yellow. Toluidine blue at high pH then stains bacteria and most tissue elements except yellow mucus. Because bacteria are restricted to yellow mucus within and upon the surface of mucous cap cells, they stand out clearly. Alcian yellow has blocked the acidic sites in the mucus, and cannot be displaced by toluidine blue. Further, Alcian yellow is insoluble in higher alcohol, so it is not removed during dehydration; toluidine blue (and nearly all other basic dyes) are extracted in higher alcohol solutions. In summary, alcian yellow is a monoazo dye that reacts similar to alcian blue, staining acid mucosubstances yellow. Toluidine blue is a basic dye and metachromatic stain that stains the H. pylori bacteria and nuclei blue.

Alcian yellow-toluidine blue method
(preferred fixative)

10% NBF

Alcian yellow-toluidine blue method
(preferred thickness)

Cut paraffin sections at 4 - 5 µm.

Alcian yellow-toluidine blue method
(control tissue)

Sections containing H. pylori.

Alcian yellow-toluidine blue method
(major reagents & purpose)

Periodic acid - oxidizes the hydroxyls in mucus to aldehydes.

Sodium metabisulfite - attacks the aldehydes to create sulfonic acids. \n \n Alcian yellow acetic acid ethanol solution - demonstration of acidic mucosubstances. \n \n Toluidine blue sodium hydroxide solution - demonstration of H. pylori.

Alcian yellow-toluidine blue method
(results)

H. pylori, nuclei - blue (toluidine blue).

Mucin - yellow (alcian yellow). \n \n Background - pale blue.

Hotchkiss-McManus PAS reaction
(purpose of stain)

Demonstration of fungi in tissue.

Hotchkiss-McManus PAS reaction
(principle of the technique)

Polysaccharides present in the fungal cell walls are oxidized by periodic acid to aldehydes. The aldehydes react with Schiff reagent to yield rose-colored fungi.

Hotchkiss-McManus PAS reaction
(preferred fixative)

10% NBF, Bouin solution or Zenker solution.

Hotchkiss-McManus PAS reaction
(preferred thickness)

Cut paraffin sections at 4 - 5 µm.

Hotchkiss-McManus PAS reaction
(control tissue)

A section containing fungi must be used for a control.

Hotchkiss-McManus PAS reaction
(major reagents & purpose)

Periodic acid - oxidation of polysaccharides present in the fungal cell walls.

Schiff reagent (basic fuchsin, sodium metabisulfite) - demonstrates aldehydes generated by previous oxidation step. \n \n Sulfurous acid rinse (HCl, sodium metabisulfite) - washes away unbound leucofuchsin to prevent false positive result. \n \n Wash in running tap water - to develop full color. \n \n Fast green - counterstain.

Hotchkiss-McManus PAS reaction
(results)

Fungi - rose (PAS).

Background - green (fast green).

Hotchkiss-McManus PAS reaction
(sources of error)

When staining for fungi, the green counterstain provides a better contrast, without masking organisms, than does hematoxylin. Light green may also be used as a counterstain. In all of the methods for fungi, it might be helpful to use diastase digestion on sections containing glycogen (e.g. liver). The oxidizing agent and the Schiff reagent must not be overused or poor stains will result. It is best if fresh oxidizing agent is used each time.

Chromic acid-Schiff stain
(purpose of stain)

Identification of fungi in tissue.

Chromic acid-Schiff stain
(principle of the technique)

Chromic acid oxidizes the carbohydrates present in the fungal cell walls to aldehydes. Chromic acid is a strong oxidizer, and when given time it will eliminate reactive aldehydes in all but the structures with the greatest concentration of carbohydrates; this includes mucin, glycogen, and fungal cell walls. Because nonspecific staining is reduced, a cleaner background is provided with chromic acid oxidation (CAS) than with periodic acid oxidation (PAS).

Chromic acid-Schiff stain
(preferred fixative)

10% NBF

Chromic acid-Schiff stain
(preferred thickness)

Cut paraffin sections at 4 - 5 µm.

Chromic acid-Schiff stain
(control tissue)

A section containing fungi must be used for a control.

Chromic acid-Schiff stain
(major reagents & purpose)

Chromic acid (60C) - oxidation of carbohydrates in fungal cell wall to aldehydes.

Schiff reagent (basic fuchsin, sodium metabisulfite) - demonstrates aldehydes generated by previous oxidation step. \n \n Sulfurous acid rinse (HCl, sodium metabisulfite) - washes away unbound leucofuchsin to prevent false positive result. \n \n Wash in running tap water - to develop full color. \n \n Harris hematoxylin or fast green - counterstain. \n \n Ammonia water - to blue hematoxylin.

Schiff reagent (basic fuchsin, sodium metabisulfite) - demonstrates aldehydes generated by previous oxidation step.

Chromic acid-Schiff stain
(results)

Fungi - deep rose to purple (CAS)

Nuclei - blue (hematoxylin, if used) \n \n Background - green (fast green, if used)

Chromic acid-Schiff stain
(sources of error)

Be sure that the Schiff reagent comes to room temperature before use. Insufficient oxidation with chromic acid may lead to increased background staining; prolonged oxidation may cause reduced staining of fungal organisms, so care must be taken to carefully control the oxidation step. Be sure that the chromic acid has not darkened because of reduction with alcohol remaining from the rehydration step; wash slides thoroughly after the alcohol before placing in chromic acid.

Gridley stain
(purpose of stain)

Demonstration of fungi in tissue.

Gridley stain
(principle of the technique)

Chromic acid oxidizes adjacent glycol groups to aldehydes, which are then reacted with Schiff reagent. Because chromic acid is a stronger oxidizing agent than periodic acid, it further attacks and destroys aldehydes, so fewer reactive groups are left to react with the Schiff reagent. A less intense reaction is obtained than with the PAS technique, but background staining is also decreased. The aldehyde fuchsin acts as an aldehyde and occupies uninvolved linkages of the Schiff reagent, thus reinforcing the depth of the stain. Both Schiff reagent and aldehyde fuchsin serve as primary stains, producing a pink to purple result against a yellow background of metanil yellow.

Gridley stain
(preferred fixative)

10% NBF

Gridley stain
(preferred thickness)

Cut paraffin sections at 4 - 5 µm.

Gridley stain
(control tissue)

A section containing fungi must be used for a control.

Gridley stain
(major reagents & purpose)

Chromic acid - oxidation of adjacent glycol groups in fungal cell wall to aldehydes.

Schiff reagent (basic fuchsin, sodium metabisulfite) - demonstrates aldehydes generated by previous oxidation step. \n \n Sulfurous acid rinse (HCl, sodium metabisulfite) - washes away unbound leucofuchsin to prevent false positive result. \n \n Rinse in several changes of 70% ethanol - prepares sections for aldehyde fuchsin solution. \n \n Aldehyde fuchsin solution (pararosaniline, 70% EtOH, HCl, paraldehyde) - to achieve more intense staining. \n \n Metanil yellow - counterstain.

Gridley stain
(results)

Mycelia, conidia, elastic fibers and mucin - deep purple (CAS & aldehyde fuchsin).

Background - yellow (metanil yellow).

Gridley stain
(sources of error)

Chromic acid is a stronger oxidizing agent than periodic acid. It oxidizes all 1,2 glycols to aldehydes and then oxidizes some of the aldehydes further to acids, especially where trace amounts of polysaccharides are present. While chromic acid continues further oxidation of all polysaccharides, complete conversion to acids in the areas of heaviest concentration takes longer, so that these remain reactive with Schiff reagent. Tissue components with heavy polysaccharide concentrations (mucin, glycogen, fungi) will continue to show a positive Schiff reaction long after basement membranes and connective tissue become nonreactive. Unlike chromic acid, periodic acid does not oxidize aldehydes to acids so that many more tissue components remain reactive with Schiff reagent (PAS) and with silver methods that detect aldehyde groups.

Grocott methenamine-silver nitrate stain
(purpose of stain)

Demonstration of fungal organisms in tissue sections. In addition to fungi, the procedure will also demonstrate Actinomyces (Gram-positive bacteria with hyphae), Nocardia asteroids (Gram-positive bacteria causing pulmonary infection) and certain encapsulated bacteria.

Grocott methenamine-silver nitrate stain
(principle of the technique)

Polysaccharides in the fungal cell wall are oxidized to aldehydes by chromic acid. Chromic acid is a strong oxidant, further oxidizing many of the newly formed aldehyde groups to break down products that will not react; this helps suppress the weaker background reactions of collagen fibers and basement membranes. Only substances that possess large quantities of polysaccharides, such as fungal cell walls, glycogen, and mucins, will remain reactive with the methenamine-silver, reducing it to visible metallic silver. Methenamine gives the solution the alkaline properties necessary for proper reaction, and sodium borate acts as a buffer. Gold chloride is a toning solution and sodium thiosulfate removes any unreduced silver.

Grocott methenamine-silver nitrate stain
(preferred fixative)

10% NBF is preferred. Glutaraldehyde fixative should be avoided, because the free aldehyde groups can reduce silver resulting in nonspecific staining.

Grocott methenamine-silver nitrate stain
(preferred thickness)

Cut paraffin sections at 4 - 5 µm or frozen sections at 6 µm

Grocott methenamine-silver nitrate stain
(control tissue)

A section containing fungi must be used. If staining for Pneumocystis jirovecii, use a Pneumocystis control, because the timing in the methenamine-silver solution is different.

Grocott methenamine-silver nitrate stain
(major reagents & purpose)

Chromic acid - oxidize polysaccharides in fungal cell wall to aldehydes (solution may be reused).

Sodium bisulfite - to remove any residual chromic acid. \n \n Methenamine silver (borax, methenamine, silver nitrate, 58C) - silver impregnation, check microscopically (use nonmetallic forceps). \n \n Gold chloride - toner (poor toning results in marked nonspecific staining). \n \n Sodium thiosulfate - removes unreduced silver. \n \n Light green - counterstain.

Grocott methenamine-silver nitrate stain
(results)

Fungi - cell walls should be black, and the internal structures should be visible. If overstained the morphology of the fungi is obscured and identification of the organisms is very difficult (silver).

Mucin - taupe to dark gray (silver). \n \n Background - green (light green).

Grocott methenamine-silver nitrate stain
(sources of error)

Failure to adequately remove the alcohol used during deparaffinization and hydration will result in reduction of the chromic acid solution; this will cause the color of the solution to change from orange to brown. The solution should be discarded when a color change is noted. Chemically clean glassware and nonmetallic forceps must be used. The silver should not be overheated, because it will cause breakdown of the solution yielding formaldehyde which nonselectively reduces silver thus causing nonspecific staining. Substituting periodic acid for chromic acid potentially results in false negatives due to inadequate oxidation. Because chromic acid is a stronger oxidizer than periodic acid it decreases staining of connective tissue thus producing a cleaner background. Conversely, inadequate oxidation by periodic acid will cause reticulin fibers and basement membranes to stain strongly, and can mask the fungal organisms when they are few in number (i.e. false negative).

Warthin-Starry technique
(purpose of stain)

Demonstration of spirochetes in tissue sections

Warthin-Starry technique
(principle of the technique)

This is an argyrophil method; i.e. spirochetes have the ability to bind silver ions from a solution, but they do not have the ability to reduce the silver to a visible metallic form. A chemical reducer, hydroquinone, is used for that purpose (note there is no oxidation step).

Warthin-Starry technique
(preferred fixative)

10% NBF.

Warthin-Starry technique
(preferred thickness)

Cut paraffin sections at 4 - 5 µm

Warthin-Starry technique
(control tissue)

The tissue must contain spirochetes.

Warthin-Starry technique
(major reagents & purpose)

Acidulated water (citric acid pH 4.0) - acidifies sections to promote impregnation.

Impregnation solution (silver nitrate, acidulated water at 43C) - spirochetes adsorb silver ions. \n \n Developer solution (silver nitrate, acidulated water, gelatin, hydroquinone at 54C) - reduce silver to visible metallic form

Warthin-Starry technique
(results)

Spirochetes and other bacteria (A. felis, L. pneumophila, N. asteroids, H. pylori) - black (silver)

Background - pale yellow to light brown

Warthin-Starry technique
(sources of error)

If the sections have been overdeveloped, they may be treated with iodine and sodium thiosulfate for color removal and then restained. All bacteria are nonselectively blackened by silver impregnation methods, which best demonstrate small, weakly Gram negative bacteria. When compared with Gram stains, silver impregnation procedures provide much greater sensitivity when screening for small numbers of both Gram-positive and Gram-negative bacteria. Any reducing substances present in the tissue (eg, formalin pigment) will also give a positive reaction. New plastic centrifuge tubes and chemically clean glassware must be used.

Churukian & Schenk modified Warthin-Starry
(purpose of stain)

Demonstration of bacteria in tissue sections

Churukian & Schenk modified Warthin-Starry
(principle of the technique)

This is an argyrophil method; i.e. spirochetes have the ability to bind silver ions from a solution, but they do not have the ability to reduce the silver to a visible metallic form. A chemical reducer, hydroquinone, is used for that purpose. Note there is no oxidation step.

Churukian & Schenk modified Warthin-Starry
(preferred fixative)

10% NBF.

Churukian & Schenk modified Warthin-Starry
(preferred thickness)

Cut paraffin sections at 4 - 5 µm

Churukian & Schenk modified Warthin-Starry
(control tissue)

The tissue must contain bacteria.

Churukian & Schenk modified Warthin-Starry
(major reagents & purpose)

Glycine acetic acid (pH 4.2) - acidifies sections to promote impregnation.

Impregnation solution (silver nitrate, glycine, acetic acid at 80C) - bacteria adsorb silver ions. \n \n Developer solution (silver nitrate, gelatin, hydroquinone at 80C) - reduce silver to visible metallic form.

Churukian & Schenk modified Warthin-Starry
(results)

Alipia felis, Legionella pneumophila, N. asteroids, H. pylori - black (silver nitrate).

Nuclei - brown.

\n Erythrocytes - brown.

Churukian & Schenk modified Warthin-Starry
(sources of error)

The type of gelatin used in the developer solution is important, to give less background staining. The results are more consistent and reliable when staining and developing are done at the lowest power levels because there is a more uniform distribution of heat in the solutions at the lower power levels. Spirochetes are not well demonstrated by this method. To demonstrate spirochetes, first incubate sections in uranyl nitrate then place in developer solution. Uranyl nitrate acts as a sensitizer which is then replaced by silver.

Dieterle method
(purpose of stain)

Demonstration of spirochetes or the causative organism of legionellosis.

Dieterle method
(principle of the technique)

Spirochetes are argyrophilic; i.e. they will adsorb silver from a silver solution but the adsorbed silver must be chemically reduced to the visible metallic form. Hydroquinone is the reducing agent or developer. Note there is no oxidation step.

Dieterle method
(preferred fixative)

10% NBF