Finals GNPATH STAINS MICROORGANISMS PDF

SPECIAL STAINS FOR MICROOGRANISMS FINALS 1st SEMESTER | S.Y. 2024-2026 PROCEDURE:...

SPECIAL STAINS FOR MICROOGRANISMS FINALS 1st SEMESTER | S.Y. 2024-2026 PROCEDURE: KINYOUN ACID FAST STAIN 1.​ Deparaffinize and hydrate the sections to Millipore-filtered water. Purpose: Detection of acid fast 2.​ Stain the sections with freshly filtered carbol-fuchsin solution mycobacteria in tissue sections for 30 minutes. 3.​ Wash sections well in running water. Lipoid capsule of acid-fast 4.​ Decolorize with 1% acid alcohol solution until sections are Principle: organisms takes up carbol- pale pink. fuchsin and resists 5.​ Wash thoroughly with tap water, then with distilled water. decolorization with dilute 6.​ Counterstain by dipping one slide at a time in the methylene mineral acid. blue working solution. Secretions should be pale blue. 7.​ Wash with tap water, then with distilled water. Fixative: 10% neutral buffered formalin - 8.​ Dehydrate quickly in 95% and absolute alcohols, 2 changes preferred each. 9.​ Clear with 2 changes of xylene, 2 minutes each 10.​ Mount with synthetic resin. KINYOUN ACID FAST STAIN PROCEDURE RESULT Acid fast bacteria- bright red Background - light blue MICROWAVE ZIEHL- NEELSEN METHOD FOR ACID FAST BACTERIA 1.​ Deparaffinize sections through 2 changes of xylene, hydrate through absolute and 95% alcohols, and rinse in Millipore-filtered (pore size 0.45 μm or smaller) water. 2.​ Stain in Kinyoun carbol-fuchsin solution (freshly filtered) for 1 PROCEDURE: hour at RT or for 30 mins at 56°C. 1.​ Deparaffinize and hydrate the sections to Millipore-filtered 3.​ Wash well in running tap water. water. 4.​ Differentiate in 2 changes of 1% acid alcohol until the tissue 2.​ Place the slides in carbol-fuchsin in a glass coplin jar, and is pale pink. microwave at power level 1 (60W) for 1 ½ minutes. Dip the 5.​ Wash the sections in running tap water. Carry slides through slides up and down several times, and allow them to remain the remainder of the procedure one at a time. in the warm solution for 15 minutes. 6.​ Counterstain in working methylene blue solution for a few 3.​ Wash well in running water to remove excess stain. dips. Do not overstain; sections should be sky blue. 4.​ Decolorize with acid alcohol until sections are pale pink. 7.​ Rinse the sections in tap water. 5.​ Wash in running water for 1 minute, and rinse in 2 changes 8.​ Dehydrate with 2 changes each of 95% and absolute of distilled water. alcohols, clear with 2 or 3 changes of xylene, and mount with 6.​ Counterstain with methylene blue solution for 15 seconds. synthetic resin. 7.​ Rinse with 95% and absolute alcohol, 2 changes each. RESULT 8.​ Clear in 3 or 4 changes of xylene, and mount with synthetic Acid fast bacteria - bright red resin. Background - blue RESULT Acid fast bacilli including Mycobacterium avium intracellulare - red ZIEHL- NEELSEN METHOD FOR ACID FAST BACTERIA Erythrocytes - pink Mast cells - blue PURPOSE: Other tissue elements - pale blue Detection of acid-fast mycobacteria in tissue sections. PRINCIPLE: ​ The lipoid capsule of acid-fast organisms takes up carbol-fuchsin and resists decolorization with dilute mineral acid. ​ Alcoholic, rather than aqueous, solutions of acid are used because more uniform decolorization is obtained with alcoholic solutions. FIXATIVE: Any well-fixed tissue, with the exception of that fixed in Carnoy solution. go future RMT Page 1 of 7  COLOR CODES: HEADINGS | SUBHEADINGS FITE ACID FAST STAIN FOR LEPROSY ORGANISMS (600 W) for 4 minutes. Dip the slides up and down several times, and allow them to remain in the hot solution (80 celcius) for 3 minutes. Discard used solutions. 3.​ Rinse in 3 changes of sterile distilled or millipore filtered water. 4.​ Differentiate sections in 2 changes of acid alcohol, 1 ½ minutes in each change. 5.​ Rinse in 4 changes of distilled water. 6.​ Stain in 0.1% eriochrome black T for 15 seconds. 7.​ Rinse in 3 changes of distilled water. 8.​ Stand slides on end, and thoroughly air dry. PURPOSE: 9.​ Dip in xylene, and mount with synthetic resin. Detection of Mycobacterium leprae in tissue sections 10.​ Examine sections with a high dry objective, a UG1 or UG2 exciter filter, and a colorless UV barrier filter. PRINCIPLE: RESULT: The lipoid capsule of the organism takes up carbol-fuchsin and resists decolorization with dilute mineral acid. Acid fast organisms- reddish yellow fluorescence Background- black FIXATIVE: 10% Neutral buffered formalin - preferred Other fixative, except Carnoy's solution may be used BROWN- HOPPS MODIFICATION OF GRAM STAIN PROCEDURE: 1.​ Deparaffinize sections with two 12- minute changes of xylene- peanut oil mixture. 2.​ Drain sections, wipe off excess oil, and blot to opacity. The residual oil helps prevent shrinkage and injury of the sections. 3.​ Stain sections in freshly filtered Ziehl Neelsen carbol- fuchsin solution for 20-30 minutes at room temperature. This solution may be saved for reuse. 4.​ Wash sections in running tap water. 5.​ Differentiate slides individually with 1% acid alcohol until the sections are faint pink. PURPOSE: 6.​ Wash in tap water. Demonstration of gram negative and gram positive bacteria in tissue. 7.​ Counterstain sections lightly with working methylene blue 10% Neutral buffered formalin solution. Do not overstain; the sections should look sky blue. PROCEDURE: 8.​ Rinse off excess methylene blue in tap water. 1.​ Deparaffinize and hydrate sections to distilled water 9.​ Blot sections and let stand for a few minutes to air dry 2.​ Stain sections with crystal violet for 2 minutes. completely. 3.​ Rinse slides in distilled water. 10.​ Mount in air dried sections with synthetic resin. Do not use 4.​ Stain slides with gram iodine for 5 minutes. alcohol and xylene. 5.​ Rinse slides in distilled water to remove excess iodine. RESULT: 6.​ Blot 1 slide at a time with slightly damp filter paper, and M. leprae and other acid fast bacteria - bright red decolorize quickly in acetone. Background - light blue 7.​ Rinse slides quickly but thoroughly in distilled water. 8.​ Stain sections with working basic fuchsin for 5 minutes. MICROWAVE AURAMINE- RHODAMINE FLUORESCENCE 9.​ Rinse slides in distilled water. TECHNIQUE 10.​ Differentiate sections with gallego solution for 5 minutes. 11.​ Rinse slides in distilled water and blot sections, but do not blot to dryness. 12.​ Quickly dip slides in acetone 3 times. 13.​ Quickly dip slides in picric acid acetone 3 times 14.​ Quickly dip slides in acetone 3 times. 15.​ Pass slides through acetone xylene mixture (1:2) for 5 quick dips, and then clear with 2 changes of xylene. 16.​ Mount with synthetic resin. RESULT: Gram positive bacteria - blue Gram negative bacteria - red Background tissue - yellow PURPOSE: Nuclei- light red Detection of Mycobacterium tuberculosis or other acid fast organisms. PRINCIPLE: 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. FIXATIVE: 10% Neutral buffered formalin PROCEDURE: 1.​ Deparaffinize and hydrate sections to sterile distilled or Millipore- filtered water. MODIFIED DIFF QUICK GIEMSA STAIN FOR HELICOBACTER 2.​ Place the slides in 45 ml of the auramine O-rhodamine B PYLORI solution in a glass coplin jar, and microwave at power level 1 Good luck! Page 2 of 7  COLOR CODES: HEADINGS | SUBHEADINGS 7.​ Wash well with water 8.​ Stain with toluidine blue solution for 3 minutes 9.​ Wash well with water 10.​ Blot sections dry 11.​ Dehydrate, clear, and mount with synthetic resin RESULT: H. pylori oranisms - blue Mucin - yellow Background - blue GRIDLEY FUNGUS STAIN PURPOSE: Identification of H. pylori in tissue sections PRINCIPLE: The romanowsky stain, neutral dyes combining the basic dye methylene blue and the acid dye eosin, give a wide color range when staining tissues and blood smears. FIXATIVE: 10% neutral buffered formalin PROCEDURE: 1.​ Deparaffinize sections in xylene and hydrate with 2 changes of absolute alcohol, 2 changes of 95% alcohol, and one change of 70% alcohol to distilled water. Carry slides one at a time through the remainder of the procedure. 2.​ Dip the slide in Diff-Quik Solution I, 25 dips. 3.​ Dip slide in Diff-Quik Solution II, 25 dips. 4.​ Rinse quickly in distilled water. 5.​ Differentiate in 2 changes of acetic water, 5 dips in each. 6.​ Rinse quickly in distilled water. Check microscopically, H. pylori and nuclei should be dark blue, cytoplasm should be pink. If greater enhancement of the stain is desired, steps 2-6 can be repeated. 7.​ Dehydrate in 1 change of 65% alcohol, 15 quick dips. 8.​ Continue dehydration with 1 change of absolute alcohol, 15 PURPOSE: quick dips. Demonstrate of fungi in tissues 9.​ Clear in xylene, and mount with synthetic resin. PRINCIPLE: RESULT: This is a modification of the Baurer technique, which uses chromic H. pylori - dark blue acid to oxidize adjacent glycol groups to aldehydes. The aldehydes Other bacteria - blue are then reacted wtih Schiff reagent. Nuclei - dark blue FIXATIVE: Cytoplasm - pink 10% Neutrabl buffered formalin ALCIAN YELLOW - TOLUIDINE BLUE METHOD FOR H. PYLORI PROCEDURE: 1.​ Deparaffinize the sections, and hydrate to distilled water. 2.​ Oxidize sections in 4% chromic acid for 1 hour. 3.​ Wash slides in running water for 5 minutes. 4.​ Stain sections in schiff reagent for 15 mins 5.​ Wash slides in running for 15 mins 6.​ Rinse in several changes of 70% alcohol 7.​ Stain sections in aldehyde fuchsin for 30 mins 8.​ Rinse off excess stain with 95% alcohol 9.​ Rinse slides in distilled water 10.​ Counterstain sections with metanil yellow solutions for 30 secs to 1 min. Do not ovestain. PURPOSE: 11.​ Rinse slides in distilled water Detection of H. pylori in tissue sections 12.​ Dehydrate in 2 changes each of 95% and absolute alcohol, PRINCIPLE: clear in xylene, and mount in synthetic resin. Alcian yellow is a monoazo dye that reacts similarly to alcian blue, RESULT: staining mucin yellow Mycelia - deep purple Toluidine blue is a basic dye and metachromatic stain that stains the Conidia - deep rose to purple H. pylori organisms and nuclei blue. Background- yellow FIXATIVE: Elastic fibers and mucin - deep purple 10% Neutral buffered formalin PROCEDURE: 1.​ Deparaffinize and hydrate sections to distilled water. 2.​ Oxidize in 1% periodic acid for 10 minutes. 3.​ Wash well with water GROCOTT METHENAMINE SILVER NITRATE FUNGUS STAIN 4.​ Place sections in sodium metabisulfite solution for 5 minutes. 5.​ Wash in running water for 2 minutes 6.​ Stain with alcian yellow for 5 minutes Good luck! Page 3 of 7  COLOR CODES: HEADINGS | SUBHEADINGS PURPOSE: PURPOSE: Demonstration of spirochetes in tissue secretions Demonstration of fungal organisms in tissue sections PRINCIPLE: PRINCIPLE: This is an argyrophil method, that is; the spirochetes have the ability Polysaccharides in the fungal cell wall are oxidized to aldehydes by to bind silver ions from a solution, but they do not have the ability to chromic acid. reudce the silver to a visible metallic form. A chemical reducer, Chromic acid is a strong oxidant, further oxidizing many of the newly hydroquinone, is used for that purpose. released aldehyde groups; this helps suppress the weaker FIXATIVE: background reactions of collagen fibers and basement membranes. 10% NBF Substance that possess large quantities of polysacchardies, such as fungal cell walls, glycogen and mucins, will remain reactive with PROCEDURE: methenamine silver, reducing it to visible metallic silver. 1.​ Place the 2% silver nitrate, 5% gelatin, and hydroquinone Sodium borate acts as a buffer, Gold chloride is a toning solution and solutions in separate 50 ml plastic centrifuge tubes. Heat in a the sodium thiosulfate removes any unreduced silver. water bath at 540 for at least 1 hour. 2.​ Place a 100ml graduated cylinder and a chemically clean coplin jar in the oven for at least 1 hour FIXATIVE: 3.​ Deparaffinize and hydrate sections acidulated water 10% NBF 4.​ Place slides in 1% silver nitrate impregnating solution in a PROCEDURE: water bath at 43C for 30 mins. Do not preheat solation. 1.​ Deparaffinize sections, and hydrate to distilled water. 5.​ Just before the slides are due out of the impregnating 2.​ Oxidize sections in chromic acid solution for 1 hour at room solution, prepare the developer and place in the 54C water temperature or for 5-10 minutes in a solution preheated to both 60°C. Begin preheating the silver solution about 20 minutes 6.​ Put slides in the developer for 3-4 mins. Check ofter 2 mins before needed. The chromic acid solution may be reused and continue checking frequently until they are ready until it turns dark. 7.​ Wash slides quickly and thoroughly in distilled water. 3.​ Wash slides in running tap water for a few seconds. 8.​ Dehydrate sections in 95% and absolute alcohols, and clear 4.​ Rinse in 1% sodium bisulfite for 1 minute to remove any in xylene (2 changes each) residual chromic acid 9.​ Mount sections with synthetic resin 5.​ Wash in tap water for 5-10 minutes. 6.​ Wash with 3 or 4 changes of distilled water. RESULT: 7.​ Using nonmetallic forceps, place stidos in preheated working Spirochetes - black methenamine silver solution in the water bath at 56°C to Other bacteria - black 58°C for 15 minutes or until sections turn yellowish brown Background - pale yellow to light brown (paper-bag brown). Remove the control, rinse in distilled water, and chock microscopically for adequate silver impregnation. Fungi should be dark brown at this stage. If DIETERLE METHOD FOR SPIROCHETES AND LEGIONELLA impregnation is not sufficient, return the slide to the methenamine silver and check every 3-5 minutes. ORGANISMS 8.​ Rinse slides in 6 changes of distilled water. 9.​ Tone in 0.1% gold chloride solution for 2-5 minutes. This solution may be used until brown precipitate appears and he solution is cloudy 10.​ Rinse sections in distilled water. 11.​ Remove unreduced silver by placing the slides in 2% sodium thiosulfate solution for 2-5 minutes. 12.​ Wash thoroughly in tap water. 13.​ Counterstain with a working light green solution for 1 1⁄2 minutes. 14.​ Dehydrate with 2 changes each of 95% and absolute alcohols. 15.​ Clear with 2-3 changes of xylene, and mount with a synthetic resin. PURPOSE: Demonstration of spirochetes and Legionella organisms RESULT: PRINCIPLE: Fungi- cell walls should be crisp black and the internal structures Spirochetes are argyrophilic; they will adsorb silver from the silver should be visible. solution. Hydroquinone is used as the reducing agent or “developer” Mucin- taupe to dark gray FIXATIVE: Background- green 10% NBF WARTHIN STARRY TECHNIQUE FOR SPIROCHETES PROCEDURE: Good luck! Page 4 of 7  COLOR CODES: HEADINGS | SUBHEADINGS 1. Preheat the 5% alcoholic uranyl nitrate solution and the 1% silver nitrate solution in a 56 C-58 C oven for at least 30 mins. ENDOGENOUS PIGMENTS Hematogenous or blood 2. Deparaffinize and hydrate sections to distilled water. Use 3 control derived pigments: hemosiderin, slides hemoglobin, bile pigment. 3. Place sections in preheated 5% alcoholic uranyl nitrate in a 55 C-60 Non Hematogenous melanin, C oven for 30 mins-1hr lipofuscin, and chromaffin 4. Dip sections once in distilled water. Endogenous minerals iron, 5. Dip section once in 95% alcohol calcium and copper 6. Place slides in 10% alcoholic gum mastic for 3 mins 7. Dip sections once quickly in 95% alcohol Consists of foreign materials, 9. Place sections in distilled water for 1 min, then allow slides to drain usually minerals introduced to for 15-20 mins until almost dry. Slides may be left overnight. the body thru air, food, 9. Place sections in preheated 1% silver nitrate solution in a 55-56C medications and injections. oven, in the dark for 5 hours. EXOGENOUS PIGMENTS Ex: Tattoos, asbestos, carbon, 10. Quickly dip slides twice in distilled water. silica, iron, silver 11. Place sections in developer dip until sections are tan to gold. Check Carbon- most common a control at 4,8, and 12 mins. Finish all sections when control is ready. exogenous pigment; appears as 12. Quickly dip twice in distilled water. jet black pigments in lung 13. Place in 10% formic acid for 45 seconds secretions and bronchial glands 14. Dip twice in distilled water of chronic smokers. 15. Dip twice in 95% alcohol 16. Dip twice in acetone ARTIFACT PIGMENTS Usually lie on top of tissue 17. Clear in 2 changes of xylene, and mount in synthetic resin instead of within the cell. Produced during processing Results: and most commonly result from Spirochetes bacteria- brown to black fixation. Background- pale yellow or tan RESULT: ENDOGENOUS PIGMENTS Spirochetes, bacteria - brown to black 1. HEMOGLOBIN Background- pale yellow or tan ​ Oxygen-containing conjugated protein found in normal RBCs ​ Only hematogenous pigment that is present in normal tissue MICROWAVE STEINER AND STEINER PROCEDURE FOR ​ RBCs are stained black by Heidenhain's iron hematoxylin, SPIROCHETES, HELICOBACTER AND LEGIONELLA ORGANISMS and blue by Mallory's PTAH 2. HEMOSIDERIN ​ Most common hemoglobin derivative; iron-containing pigment of hemoglobin ​ Seen as yellow to brown granule and is normally found inside the cells (macrophages) that have phagocytized and degraded hemoglobin ​ Hemosiderosis may be attributed to transfusion, excess dietary iron consumption or breakdown of RBCs ​ Demonstrated histochemically by Prussian blue reaction ​ Can be removed from tissue sections by 10% sulfuric acid. ​ Oxygen-containing conjugated protein found in normal RBCs ​ Only hematogenous pigment that is present in normal tissue ​ RBCs are stained black by Heidenhain's iron hematoxylin, and blue by Mallory's PTAH PURPOSE: Demonstration of spirochetes, H. pylori and Legionella 3. HEMATOIDIN PRINCIPLE: ​ Iron-free pigments of hemoglobin, found in places where The organisms demonstrated by this method are argyrophilic; they there is poor oxygenation, participating in the formation of will adsorb silver from a silver solution, be chemically reduced to bile pigments visible metallic form using a developer solution which is ​ Derived from RBCs and occurs as yellowish or greenish hydroquinone. granules or masses FIXATIVE: ​ Found in conditions such as infarct and areas of hemorrhage 10% NBF and thrombosis Mercurial end chromate fixative should be avoided 4. HEMATIN RESULT: ​ Hemoglobin minus the globin molecule Spirochetes - dark brown to black ​ Found in old blood clots H. pylori - dark brown to black ​ May be encountered in malaria, pernicious anemia, and toxic L. pneumophila - dark brown to black hemolysis Other non-filamentous bacteria - dark brown to black 5. HEMOZOIN Background-light yellow ​ Malarial pigments; may be seen in the liver, spleen, bone marrow, lymph nodes, and brain capillaries ​ Black granules formed by malarial parasites PIGMENTS AND MINERAL LILLIE'S METHOD FOR FERRIC AND FERROUS IRON Produced within the tissue to Procedure: serve as a physiological function 1.​ Deparaffinize and hydrate to distilled water. or a by-product of normal 2.​ For Ferric Iron, place in Potassium Ferrocyanide Solution for metabolism. 1 hour.​ Good luck! Page 5 of 7  COLOR CODES: HEADINGS | SUBHEADINGS For Ferrous Iron, place in Potassium Ferricyanide Solution for 1 hour. 3.​ Wash well in 1% Acetic Acid. 4.​ Stain for 10 minutes in Basic Fuchsin Solution. 5.​ Rinse in distilled water. 6.​ Dehydrate in 95% alcohol, absolute alcohol, and clear in xylene, two changes each. 7.​ Mount with Permount. Result: ​ Ferric iron – dark Prussian blue ​ Ferrous iron – dark Turnbull’s blue ​ Background – light red Perl’s Prussian Blue Method for Leuco patent blue V stain for Hemosiderin (ferric iron) hemoglobin Procedure: PROCEDURE: 1. Take test and control sections to distilled water. 2. Stain in patent blue solution for 5 minutes at room temperature. 1.​ Sections to water. 3. Rinse in distilled water. 2.​ Treat sections with freshly prepared acid ferrocyanide 4. Lightly counterstain in 0.5% aqueous neutral red or 0.1% aqueous solution for 10 to 30 minutes. nuclear fast red for 1 minute. 3.​ Wash in distilled water. 5. Rinse in distilled water. 4.​ Counterstain the nuclei with eosin, 0.5% aqueous neutral 6. Dehydrate, clear, and mount in synthetic resin. red, or 0.1% nuclear fast red. 5.​ Wash rapidly in distilled water. Result: 6.​ Dehydrate, clear, and mount in a neutral mounting medium. ​ Hemoglobin peroxidase (RBCs and neutrophil) - dark blue ​ Nuclei - red Result: ​ Hemosiderin and ferric salts: Deep blue ​ ​ Other pigments retain their natural color. Tissue and nuclei stain red (according to counterstain). Modified Fouchet's Technique for Liver Bile Pigments PROCEDURE: 1. Take test and control sections to distilled water. Gomori’s Prussian Blue Stain for Iron 2. Treat with the freshly prepared Fouchet's solution for 10 minutes. 3. Wash well in running tap water for 1 minute. Procedure: 4. Rinse in distilled water. 5. Counterstain with van Gieson's solution for 2 minutes. 1.​ Immerse slides in equal parts of 20% hydrochloric acid and 6. Dehydrate, clear, and mount in synthetic resin 10% potassium ferrocyanide mixed immediately before use in chemically cleaned glassware for 20 minutes. Results: 2.​ Wash thoroughly in distilled water. Do not use tap water. Bile pigments- emerald to blue 3.​ Counterstain in nuclear fast red for 2 minutes. Muscle-green-yellow 4.​ Rinse in distilled water. Collagen-red 5.​ Dehydrate, clear, and mount. Result: Gmelin Technique for Bile and Hematoidin ​ Iron pigments: Bright blue ​ Nuclei: Red Sections: Paraffin, frozen or cryostat ​ Cytoplasm: Pink to rose PROCEDURE: 1. Sections to distilled water and mount in distilled water. 2. Place mounted section under the microscope using an objective with reasonable working distance. Turnbull's Blue for Ferrous Iron 3. Place 2-3 drops of concentrated nitric acid to one side of the coverglass and draw under the cover glass by means of a piece of -​ Rarely used in routine histology blotting paper on the opposite side. -​ Reaction depends upon the union of ferrous iron and 4. Remove excess solution and observe pigment for color changes. potassium ferricyanide to form a blue preccipitate of complex ferrous ferricyandie. Result: Bile pigments will gradually produce the following spectrum of Results: color change: yellow-green-blue-purple-red ​ Ferrous iron - blue Good luck! Page 6 of 7  COLOR CODES: HEADINGS | SUBHEADINGS Method orange Lindquist's Copper Orange- red Modified Rhodanine Technique Gomori’s Urate crystals Black Methenamine Silver Stain Fontana Masson Method For Melanin (Melanin has the ability to reduce solutions of ammoniacal silver nitrate to metallic silver) Procedure: 1. Deparaffinize and hydrate to distilled water 2. Treat with silver solution (45 mins at 56C to demonstrate melanin; at RT overnight in the dark to demonstrate argentaffin) 3. Wash well in several changes of distilled water 4. Treat with 5% sodium thiosulfate for 2 mins 5. Wash in tap water 6. Counterstain in 1% neutral red for 3 mins 7. Wash in tap water 8. Dehydrate through graded alcohol 9. Mount and label Result: Melanin-Black Argentaffin granules - Black Nuclei-red Modified Von Kossa Method For Calcium -Von Kossa method involves the replacement of the anionic part of calcium salts with silver. Silver is made visible following reduction by exposure to bright light. PROCEDURE: 1, Deparaffinize and hydrate to distilled water. Rinse well in distilled water. 2. Place in Silver nitrate solution and expose to strong light for 10 to 20 minutes. Check the slides periodically and stop the reaction when the calcium salts are brown-black 3. Rinse sections in distilled water, 4. Place slides in 5% sodium thiosulfate for 2 to 3 minutes. 5. Wash slides in distilled water. 6. Counterstain in nuclear fast red for 5 minutes, 7. Wash sections well in water. 8. Dehydrate and clear in 2 changes each of 95% alcohol, absolute alcohol, and xylene. 9. Mount with synthetic resin. Results: Mineralized bone-black Osteoid - red Nuclei-blue OTHER SPECIAL STAINS FOR MINERALS Alizarin Red S Calcium Salts Intense reddish Good luck! Page 7 of 7