Microscopy Stains Technical Data
109 articles
3-Amino-9-Ethylcarbazole Method For Leukocyte Eroxidase
EMS26173 Fixation Formalin-Acetone Fixative Procedure Fix thin blood films in Formalin Acetone Fixative for 15 seconds. Rinse gently in distilled water. Stain in 3-Amino-9-Ethylcarbazole staining solution for 2.5 minutes. Wash in running tap water. Stain in Mayer's Haematoxylin for 8 minutes. Wash in tap water, air dry, and mount in Glycerine Jelly. Stain Results Sites of peroxidase activity Red-Brown granules Nucle Blue
AB/PAS/CG Stain
EMS26920 Components: Alcian Blue 8Gx Solution 1%, pH 2.6-3.0 Sodium Carbonate 0.3% Aqueous Solution Periodic Acid 1% Aqueous Solution Schiff’s Reagent Phophomolybdic Acid-Orange G Solution Fixation: After bisecting the gland and then bisecting each half, fix for 6-10 hours in Formalin Calcium Acetate Solution followed by 4-6 hours in 70% Alcohol before beginning dehydration and embedding. Sections: Paraffin at 3-5µm Procedure staining: Deparaffinise and hydrate to distilled water. Stain in Alcian Blue Solution for 30 minutes. Rinse briefly in water to remove excess Alcian Blue. Place for 30 minutes in Sodium Carbonate 0.3%. Wash in running water for 5 minutes. Oxidise in Periodic Acid, 1% for 10 minutes. Wash for 5 minutes in running water. Stain for 10 minutes in Schiff’s Reagent. Wash for 5 minutes in running water. Stain for 5 seconds in Phosphomolybdic Acid-Orange G Solution. Rinse briefly in water to remove excess Orange G. Dehydrate and then clear in xylene. Examine the sections while in xylene to see if the Orange G staining is too weak or too intense, and then, if necessary, rehydrate and either restain in Orange G Solution or rinse in further water. Mount with synthetic mounting medium. Results: ACTH/MSH cells - purple, red and orange-red TSH cells - light to dark purple-blue Gonadotropin cells - intense red, blue-purple, blue, turquoise GH cells - orange Refernces: McManus, J.F.A. and Mowry, R. W., Staining Methods, Histologic and Histochemical, P. B. Hoeber, NY, 1960 Clark, G., (ed.), Staining Procedures, 3rd edition, Williams & Wilkins, Baltimore, p. 166, c1973
Acridine Orange
EMS10050 Materials PBS pH 7.2 (stock solution 2X) Paraformaldehyde 3.7% (A1) Absolute Methanol (or Ethanol) (A1) Preparation of paraformaldehyde (PFA) solution (100ml): Dissolve 3.7g of paraformaldehyde in 50ml H2O, then add 2 drops of 10N NaOH, heat 30 minutes at 60°C. Add 50ml of PBS 2X. Adjust the pH to 7.2 The PFA solution must be freshly made NOTE: PFA is toxic if inhaled Staining Procedure Wash cells in PBS pH 7.2 Fix cells in 3.7% paraformaldehyde (PFA) in PBS pH 7.2 for 15 minutes at room temperature Wash once with PBS for 5 minutes at room temperature Cover the cells with methanol for 5 minutes at room temperature Wash cells in PBS of pH 7.2 Incubate with staining solution Protocol after DNA Denaturation Acridine Orange (AO) is a metachromatic dye which differentially stains double-stranded (ds) and single-stranded (ss) nucleic acids. When AO intercalates into dsDNA, it releases green fluorescence upon excitation at 480-490 nm. It emits red, however, upon interacting with ssDNA or RNA. Chromatin condensation is seen in the early stage of apoptosis and the condensed chromatin is significantly more sensitive to DNA denaturation than is normal chromatin. If RNA is removed by pre-incubation with RNase A and DNA is denaturated in situ by exposure to HCl immediately before AO staining, apoptotic cells display a strong red fluorescence and a reduced green emission when compared to non-apoptotic interphase cells. Materials Acridine Orange (A1) Citric acid Na2HPO4 Paraformaldehyde PBS DNAse-free RNAse A (A2) Ethanol HCl Equipment Epifluorescence Microscope or Flow Cytometer (A1) Staining solution Acridine Orange 6 µg/ml Citric acid 0.1 M Na2HPO4 0.2 M pH 2.6 Prepare 90 ml of citric acid solution, add acridine orange and 10 ml Na2HPO4 Acridine Orange solution is stable for several weeks when it is stored at 4°C as well as in the dark (A2) RNAse solution Dissolve 1 mg of RNAse A (use DNAse-free RNase) in 1 ml of distilled water Methodology: Wash cells (1x106) in PBS and centrifuge at 200g for 5 minutes Re-suspend the cell pellet in 1 ml PBS Fix cells by transferring the cell suspension in 9ml 1% paraformaldehyde in PBS, on ice. Incubate for 15 minutes on ice Centrifuge at 200g for 5 minutes and re-suspend the cell pellet in 5ml PBS, centrifuge Suspend the cell pellet in 1ml PBS and transfer the suspension in 9ml 70% (vol/vol) ethanol, on ice Incubate for 4 hours (These cells may be stored in ethanol for weeks at a time) Centrifuge at 200g for 5 minutes and re-suspend the cell pellet in 1ml PBS Add 0.2ml of RNAse A solution. Incubate at 37°C for 30 minutes Centrifuge at 200 g for 5 minutes and re-suspend the cell pellet in 0.2ml PBS Add 0.5ml of 0.1M HCl at room temperature After 30-45 seconds add 2ml of AO staining solution Observe the cells under fluorescence microscope with an appropriate filter set You may also analyse cells by cytometry (excitation 488nm; dot plot of green fluorescence at 530 ±20nm versus red fluorescence >600nm)
Alcain Blue (pH 2.5) Method
EMS26323 Components: Alcian Blue 1% Nuclear Fast Red Solution Acetic Acid 3% Aqueous Fixation: 10% Buffered Neutral Formalin Sections: Paraffin at 6µm sections. Staining Procedures: Deparaffinise and hydrate to distilled water. Mordant in Acetic Acid, 3% for three minutes. Stain in Alcian Blue Solution for thirty minutes. Wash in running water for ten minutes and rinse in distilled water. Counterstain in Nuclear Fast Red Solution for five minutes. Wash in running water for one minute. Dehydrate in 95% Alcohol, Absolute Alcohol, and clear in Xylene, two changes each. Mount with Permount or DPX. Stain Results: Weakly Acidic Sulfate Mucosubstances Dark Blue Hyaluronic Acid Dark Blue Sialomucins Dark Blue References: AFIP Manual of Histological Staining Methods, 3rd ed., Ed. L. Luna: New York: McGraw Hill Publications, c. 1968, p. 163. Lev, R. & Spicer, S.S., J. Histochem. Cytochem., 12:309, 1964
Alcoholic Congo Red Solution
EMS26090 Staining Protocol Deparaffinise and hydrate to distilled water. Stain in filtered Congo Red, 1% Aqueous for 1 hour. Note: If using a microwave, stain in Congo Red, 1% Aqueous for 25 seconds. The hot stain jar is removed from the oven and should sit at room temperature for 1 minute. Rinse in distilled water. Quickly differentiate in Alkaline Alcohol. Wash in running tap water for approximately 5 minutes. Counterstain in Mayer's Acid Haemalum for approximately 5 minutes. Note: If using a microwave, oven-stain for 30 seconds. Wash in running tap water for 15 minutes. Dehydrate and clear through 95% ethyl (absolute alcohol changes each), 2 minutes each. Mount using resinous medium. Stain Results Amyloids Pink Red Nuclei Blue Elastic Tissue Light Red
Aldehyde-Fuchsin Method, pH 1.0
EMS26328 Components: Aldehyde-Fuchsin Solution pH 1.0 Metanil Yellow Solution 0.25% Alcohol Solution, 80% pH 1.0 Fixation: 10% Buffered Neutral Formalin or fixative of choice. Sections: Paraffin at 6µm sections Staining Procedures: Deparaffinise and hydrate to distilled water. Rinse in several changes of Alcohol, 80% pH 1.0. Stain in Aldehyde Fuchsin Solution for 30 minutes. Rinse off excess stain with Alcohol, 80% pH 1.0. Counterstain with Metanil Yellow Solution until background is a light yellow (approximately 1 minute). Rinse in distilled water. Dehydrate in 95% alcohol, absolute alcohol, and clear in Xylene, two changes each. Mount with Permount or DPX. Stain Results: All mucins except the highly acidic sulphated mucosubstances are aldehyde fuchsin negative at pH values of 1.0 and below. References: Johnson, W.C., Graham, J.H., and Helwig, E.B., J. Invest. Derm., 42:215-224, 1964. Luna, L.G., (ed), Manual of Histologic Staining Methods of the AFIP, 3rd edition, McGraw-Hill, c 1968, p.166
Alizarin Red for Calcium Deposits
EMS26206 Components: Alizarin S 2% Solution pH 4.2 Reagent Alcohol 50% Acetone Acetone-Xylene Fixation: Alcoholic Formalin Sections: Paraffin, Ester Wax, Celloidin or Gelatin Staining Procedures: Bring sections to Alcohol, 50% and rinse rapidly in distilled water. Cover sections with Alizarin Red S and follow the staining by transmitted light, on the stage of a staining microscope. Calcium sites will be covered with a vivid orange-red calcium-alizarin lake within a period of 30 seconds to 5 minutes. The end point should be chosen when the deposit is heavy but not too diffused. Shake off excess stain and blot carefully with filter paper or alternatively drain very thoroughly. Place immediately into Acetone for 10 – 20 seconds followed by Acetone-Xylene for 10 -20 seconds. Clear in Xylene and mount in Depex or Cedar wood Oil Stain Results: Calcium Sites Covered and surrounded by heavy orange red precipitate which is birefringent between crossed polaroids Background Faint Pink References: McGee-Russell, S.M., J. Histochem., 6:22-42, 1958. Clark, G., (ed.). Staining Procedures, 3rd ed., Williams & Wilkins, Baltimore, p. 140, c. 1973.
Alizarin Red S and Toluidine Blue O
EMS26205 Components: Alizarin Red S 0.02% Aqueous Toluidine Blue O 0.25% Ammonium Alcohol Potassium Hydroxide Sulphuric Acid Alcohol Potassium Hydroxide 4% Aqueous Fixation: At least 1 week in 10% Formalin after evisceration and washing. (Evisceration is not necessary for museum specimen) Staining Procedures: Wash the specimen in 250ml of Ammonium Alcohol. For 24 hours. Stain in Toluidine Blue Solution, 0.25% for 1 week. Harden and de-stain for 72 hours in 4 changes of 95% alcohol. Macerate in several changes of Potassium Hydroxide, 2% for 5 – 7 days, depending on the size of animal. This step is facilitated exposure to sunlight or ultraviolet light. Transfer to Working* Alizarin Red S solution for approximately 24 hours. * Prepare the Working Stain by mixing equal parts of Alizarin Red S, 0.02% and Potassium Hydroxide, 4%. Prepare fresh. If specimen has been insufficiently macerated the soft will appear slightly attained. They may be quickly de-stained in Sulphuric Acid-Alcohol. Dehydrate through 3 changes of cellosolve, 6 hours each. Reduce the time for small embryos. OPTIONAL: One may substitute the alcohol series, 50%, 80% and 90% followed by 3 changes of benzene. Clear by transferring to solution for 25%, 50%, and 75% methyl salicylate in cellosolve for 24 hours each. Then store in methyl salicylate. Modification by using glycerin for clearing: Omit step 6 and transfer directly from Working Alizarin Red S solution into a series of 50%, 70% and 80% glycerin solution for 24 hours each. Store in glycerin. Stain Results: Cartilage Dark Blue Soft Tissue Transparent Osseous Tissue Deep Red Note: The intensity of the stains serves to indicate the relative amounts of ossification and chrondogeneis, which may be taken place. The bone cartilage may be stained separated by omitting step 2 for bone or step 5 for cartilage. References: Dawson, A.B., Stain Tech., 1:123 – 4, 1926. Williams, T.W., Stain Techn., 16:23-5, 1941. Clark, G., (ed)., Staining Procedures, 3rd ed., Williams & Wilkins, Baltimore, p. 139, c. 1973.
Basic Fuchsin Reagent
Preparation of Basic Fuchsin Reagent ("Cold" Method) Perform the preparation of the reagent under a fume hood or in a very well ventilated area (irritating sulphur dioxide fumes are given off during preparation of the reagent). Wear protective gloves and safety glasses and protect your clothes. Ingredients: Basic Fuchsin C.I. 42510 (or New Fuchsin C.I.42520), 1g Potassium or sodium metabisulphite, 2.8g (potassium or sodium metabisulphite can deteriorate on storage (try and obtain a relatively fresh batch) Hydrochloric acid (ex pool shop) Add 1.5ml of the concentrated acid to 100ml of distilled (or deionised water ex Woolworths) Activated charcoal Prepare 100ml of the hydrochloric acid solution, by adding 1.5ml of concentrated hydrochloric acid to 100ml of water (remember "Always do what you oughta, add the acid TO the water"). Dissolve the metabisulphite in the acid solution (if you put the metabisulphite in the solution first, you can see when it is dissolved before then adding the Basic Fuchsin). Add the Basic Fuchsin and mix thoroughly (to facilitate "wetting", damp the powdered dye with a little alcohol). Stopper the container with a lid or parafilm to stop the loss of sulphur dioxide. Shake the solution periodically until it turns a yellow to light brown colour. This takes at least 2 hours or more. The final Schiff's Reagent is a "water white" (clear solution), and to get rid of the yellowish/brown impurity present in most samples of Basic Fuchsin that does not decolourise under the action of the sulphite, the solution is treated with activated charcoal which removes the impurities. You may wish to forego this step. Add half a teaspoon of activated charcoal to the solution, and shark or stir the solution for 1?2 minutes, then filter through filter paper in a plastic funnel. Repeat step 5 until the solution is clear. Transfer the reagent to a clean, stoppered bottle, label, and store in a fridge. You will know if the reagent works, as any spilled solution will turn a bright magenta colour. Notes: Prepare the reagent in a suitable lidded glass jar or a container, or sealed with parafilm; Weigh the ingredients using a good set of digital scales; Weigh the chemicals onto a pre?folded slip of paper or a piece of aluminium foil to facilitate transfer to the container; Measure the concentrated hydrochloric acid with a plastic syringe
Bismuth Stain Kit 1 - Specific En Bloc Staining
EMS11430 Bismuth Stain Kits for Specific En Bloc Staining Kit consists of: 10 x 5ml Solution A (Sodium Tartrate, NaOH and Bismuth Subnitrate) 10 x 10ml Solution B (0.2M Triethanolamine-HCl buffer, pH 7.0) Procedures Staining solution: Mix Solution A and B together (One part of A into two parts of B) then adjust to pH 7.0 with 1N HCl. Aldehyde-fixed tissue blocks are rinsed in the above buffer and stained en-bloc with Bismuth for one hour at room temperature on a gentle rotary shaker. Post fixation is carried out with 1% Osmium Tetroxide in 0.05 M Sodium Cacodylate buffer, pH 7.2. Post-staining is not needed. Reference Locke & Huie, (1977). Bismuth staining for Light and Electron Microscopy. Tissue Cell 9, 347.
Bismuth Stain Kit 2 - Enhancing Contrast in Uranyl Acetate Treated Tissue
EMS11434 Bismuth Stain Kit for Enhancing Contrast in General consists of: 5 x 10ml Solution A (Sodium Tartrate in NaOH) 5 x 200mg Ingredient B (Bismuth Subnitrate) Procedures Drops of solution A are added into the ingredient B (Bismuth Subnitrate). The mixture is cleared after 6-8ml of solution A is added; and after the addition of all of solution A, all the Bismuth is chelated. Conventionally fixed tissue specimens are stained en-bloc with a saturated Uranyl Acetate aqueous solution for 20 minutes. Thin sections on grids are stained under cover for 3 minutes by immersion in the Bismuth Subnitrate chelated above, and then rinsed with distilled water. Reference Riva, A. (1974). A simple and rapid staining method for enhancing the contrast of tissues previously treated with Uranyl Acetate J. Microsc. ( Paris ) 19, 105.
Bismuth Stain Kit 3 - Mucosubstances and Polysaccharides
EMS11436 The Bismuth Stain Kit for Staining of Mucosubstances and Polysaccharides consists of: 10 x 10ml Solution A (Periodic acid, Ethanol, Sodium acetate in distilled water) 10 x 10ml Solution B (Sodium tartrate, NaOH, Bismuth subnitrate) Procedures Thin sections of conventionally fixed tissues are mounted on copper grids and floated on solution A for 10-30 minutes at room temperature. After being thoroughly washed 20-30 times for 5-10 minutes with distilled water to remove periodate, the grids are floated on solution B for 30-60 minutes at room temperature. They are then again thoroughly washed with distilled water. Reference Ainsworth, S.K., Ito S., and Karnovsky,M.J. (1972). Alkaline bismuth reagent for high resolution ultrastructural demonstration of periodic-reactive sites. J. Histochem, Cytochem. 20, 995.
Bodian's Protargol Method
EMS26693 Components Protargol Solution, 1% Aqueous Cooper Shot Gold Chloride, 1% Aqueous Aniline Blue Solution Lissamine Fast Red Solution Oxalic Acid, 2% Aqueous Sodium Thiosulfate, 5% Aqueou Phosphomolybdic Acid, 1% Aqueous Tartrazine Solution, 1.5% Hydraquinone Reducing Solution Fixation Formalin - Alcohol - Acetic Acid Sections Paraffin At 6µm Staining Deparaffinise and hydrate to distilled water. Place slides in Protargol Solution and add 4-6g clean copper shot per 100ml of solution. Let stand at 37°C for 48 hours or 60°C for 4-16 hours. Staining is complete when sections are golden brown. Wash by dipping in distilled water Reduce in hydroquinone solution for 5-10 minutes. Wash thoroughly in distilled water, three changes. Tone in Gold Chloride, 1.0% for 5-10 minutes. Rinse in distilled water, three changes. Develop in oxalic acid 2% until background is gray and nerve fibers appear clearly, 3-5 minutes. Rinse in distilled water, three changes. Place in sodium thiosulfate 5% 2-5 minutes. Rinse in distilled water. Counterstain with aniline blue solution 2 or 3 quick dips. Alternatively Counterstain with lissamine fast red for 5 minutes. Place in phosphomolybdic acid 1% for 5 minutes. Rinse briefly in distilled water. Place in tartrazine solution for 5 minutes. Rinse briefly in 95% alcohol. Dehydrate rapidly in absolute alcohol and clear in xylene, two changes each. Mount. Staining Results Nerve Fibres Black Myelinin, Muscle, Erythrocytes Lissamine Fast Red Counterstain Red Background-Lissamine fast red Counterstain Yellow Background-Aniline Blue Counterstain Blue Copper shot is good as long as it retains a metallic luster. If a dull oxide film forms discard the Copper Shot or clean by immersing in 30% Nitric Acid (30ml of conc. Nitric Acid added slowly and carefully to 70ml of distilled water) until the surface film or oxides is removed. The Copper will be bright in colour and all dark areas of oxide removed. Rinse quickly in D-H20, several changes, followed by one or two changes of alcohol or acetone. Dry in a low temperature (approx. 500°C) oven for 15-30 minutes. Store in a tightly sealed bottle. References Luna, L.G., (ed.), Manual of Histologic Staining Methods of the AFIP 3 rd edition, McGraw Hill, NY, c 1968, p. 195. Clark, G., (ed.) Staining Procedures , 3 rd edition, Williams & Wilkins co., Baltimore c. 1973, 98.
Brown and Brenn Method for Gram + and Gram - Bacteria
EMS26105 Components: Crystal Violet Stain Solution 1% Aqueous Sodium Biocarbonate 5% Aqueous Basic Fuchsin Stock Solution 0.25% Basic Fuchsin Working Solution Gram's Iodine Solution Acetone-Alcohol 1:1 Picric Acid Acetone Solution 0.1% Acetone Acetone:Xylene Fixation: Formalin, 10% Buffered Neutral Section: Paraffin, 6µm *Recommended technique includes a control slide. Staining Procedure: Deparaffinise and hydrate to distilled water. Mix 1.0ml (20dr.) Crystal Violet, 1% Aq. with 5 drops Sodium Bicarbonate, 5%, Aq.; pour onto slides held in a staining rack. Agitate gently to cover section: stain slides for 1 min. Rinse in distilled water. Flood with Gram’s Iodine for 1 min, rinse with water and carefully blot with filter paper to dryness. Decolourise with Acetone-Alcohol, 1:1 by dropping onto the slide until no more colour runs off. Stain in the Basic Fuchsin Working or (dilute one vol. Basic Fuchsin Stock, 0.25%, Aq., with 10 vol. distilled water), 1 minute; wash in water, blot carefully but not to complete dryness as in step #3. Differentiate in Acetone, i.e. one quick dip, then transfer immediately to the Picric Acid – Acetone Solution, 0.1% to complete. Differentiate until sections show yellowish-pink. Rinse quickly in Acetone, then Acetone-Xylene. Clear in 3-4 changes Xylene alone. Mount. Results: Gram+ Bacteria, Nocardia and Actinomyces Filaments Blue Gram- Bacteria, Nuclei Red Additional tissue elements Yellow Note: See also the Taylor modification of the Brown and Brenn+/- technique noted for the varying differentiation available. Over-differentiation in the B&B step #6 is a problem with some sections; run the control slides at varying rates to determine the amount for the specific organism. References: Brown, J.H. and Brenn, L. Bull. Johns Hopkins Hosp., 48:69 (1931). AFIP Manual of Histologic Staining Techniques: 3rd. ed., ed. G. Luna; New York: McGraw-Hill Publications, c. 1968, p. 222.
Brown-Hopps Method For Gram Positive And Gram Negative Bacteria
EMS26106 Components Crystal Violet Staining Solution, 1% Aqueous Gram's Iodine Solution Basic Fuchsin Staining Solution, 0.5% Aqueous Gallego’S Differentiating Solution Cellosolve Tartrazine, 1.5% Aqueous Fixation 10% Buffered Neutral Formalin Section Paraffin, @ 6µm Staining Procedure Deparaffinise and hydrate to distilled water. Place slides on staining rack and pour on Crystal Violet Solution, 1%, Aqueous for two minutes. Rinse in distilled water. Mordant in Gram's Iodine for five minutes. Rinse in distilled water. Differentiate in Cellosolve, until blue color no longer streams away from the section (approximately 5-10 seconds). (See note) Quickly rinse in distilled water. (See note) Stain in Basic Fuchsin Solution, 0.5%, Aqueous for five minutes. Rinse in distilled water. Gallego's Solution , for five minutes. (Differentiates and "fixes" the Basic Fuchsin). Rinse thoroughly in distilled water and blot lightly to remove excess water (not to dryness). Stain for three seconds in Tartrazine Solution . Immediately blot away excess, but not to dryness). Cellosolve three changes, for six quick dips in each. (See note) Xylene, three changes, for 10 dips each. Slides may remain in Xylene until ready for mounting. Mount with Permount. Results Gram Negative Bacteria red Gram Positive Bacteria blue Background yellow Notes: Preferably, all stains and solutions are applied to the slide while it is in a horizontal position, except for steps 4, 10 and 11, where the slide should be dipped into the solutions contained in coplin jars. Steps 4, 5, 9 and 10 are critical with regard to timing. This procedure does not work well on Bouin's fixed tissues. References AFIP Manual of Histologic Staining Methods; ed. G. Luna, McGraw-Hill Publications, 3 rd . ed.; New York, c. 1968, p.224. Brown, R. C. and Hopps, H.C., Geographic Pathology Division Armed Forces Institute Of Pathology, Washington, D.C. 20305
Canaliculi and Lacunae Stain for Hard Tissues
EMS26201 Components: Cupric Nitrate 1% Aqueous Protargol Solution 1% Aqueous Copper Shot Hydroquinone Reducing Solution Gold Chloride 1% Aqueous Oxalic Acid 2% Aqueous Sodium Thiosulfate 5% Aqueous Fixation: Modified Bouin's Solution Decalcification: Trichloroacetic Acid – Alcohol Embedded in: Paraffin, ester wax, celloidin or gelatin Procedure: Deparaffinise and hydrate to distilled water. Mordant in Cupric Nitrate, 1% overnight. In a coplin jar, add 4-6 gm Copper Shot per 100ml of Protargol Solution and place slides in the solution. Place in a 60°C oven. Staining is complete when the sections have a golden brown color (4-16 hours). Wash very briefly in distilled water (1 dip). Reduce in Hydroquinone Solution for 3-10 minutes. Wash thoroughly in running water to remove all of the reducing solution. Tone 5-10 minutes in Gold Chloride, 1% to which 3 drops of Glacial Acetic Acid per 100 ml have been added. Rinse in water. Develop in Oxalic Acid, 2% until sections have a faint purple or blue color. Rinse in water. Place in Sodium Thiosulfate, 5% for 5-10 minutes. Wash thoroughly in water. Dehydrate, clear and cover. Stain Results: Canaliculated, lacunae, odontoblasts and dentinal tubules Bluish to purplish black References: Powers, M.M., Rasmussen, G.L., and Clark, G., Anat.Rec., 111:171-6, 1951 Clark, G. (ed), Staining Procedures, 3rd edition, Williams & Wilkins, Baltimore, c 1973, p 136 Bodian, D., Anat Rec., 65:89-97, 1936 Bodian, D., Anat. Rec., 69:153-62, 1937
Carstairs' Method for Fibrin & Platelets
EMS26381 Components: Ferric Ammonium Sulphate 5% Mayer's Haematoxylin Picric Acid-Orange G Solution Ponceau Acid Fuchsin Solution Phosphotungstic Acid 1% Aqueous Aniline Blue Solution Fixation: Formalin-Saline for 48 hours or more Sections: Cut paraffin at 5µm sections Staining Procedures: Hydrate slides to distilled water. Mordant in Ferric Ammonium Sulphate, 5% for 5 minutes. Rinse in running tap water. Stain with Mayer's Haematoxylin for 5 minutes. Rinse in running tap water. Stain in the Picric Acid-Orange G Solution for 30 minutes to 1 hour. Rinse once in distilled water. Stain in Ponceau Fuchsin Solution for 1 to 5 minutes. Rinse in distilled water. Differentiate with Phosphotungstic Acid, 1% until the muscle is red and background pale pink. Rinse in distilled water. Stain with Aniline Blue Solution for 30 minutes to 2 hours. Rinse in several changes of distilled water. Dehydrate, clear in Xylene and coverslip, using a synthetic mounting medium. Stain Results: Cell Fixation Time: 48 Hours or more Fixation Time: Less than 48 hours Fibrin Bright Red Orange to Orange Red Platelets Grey Blue or Navy Light Grey Collagen Bright Blue Bright Blue Muscle Red Red Red Blood Cells Clear Yellow Red, Green or Yellow
Cresyl Fast Violet, (FIRM)
EMS12780 Bridging the gap between light and electron microscopy. Fluorophore-Infiltrated Resin Microscopy (FIRM) is a novel staining method providing ultra-high contrast imaging of tissue subcellular structure using a standard widefield fluorescence microscope. FIRM images closely resemble those obtained by transmission electron microscopy (TEM) at low magnifications. The key concept is the infiltration of thin resin sections with a fluorophore. FIRM can be used either in epoxy or acrylate resins. When illuminated by epifluorescence, the brightly fluorescent resin reveals cell and tissue structures in negative relief, according to the light absorbing and scattering properties of such structures. The method provides much greater contrast of fine subcellular structures than obtained by typical semithin section brightfield stains, such as toluidine blue. FIRM images of multiple (>50) serial semithin sections of complex structures, such as kidney glomeruli, are easily rendered in 3-D by open source software programs. General Methods During the testing of a large number of fluorescent dyes as candidate counterstains for thinsection immunofluorescence in LR white sections, one with special properties was encountered. Most dyes, such as sodium fluorescein, non-selectively stained tissue and provided little contrast (not shown). However, one such dye, when applied to either LR White or Epon sections, provided very high contrast images using standard fluorescence microscopy, with the resin appearing brightly fluorescent and tissue structures appearing primarily in negative relief (Figure 1). FIGURE 1: Human anterior pituitary, FIRM, rhodamine channel. Note that the fluorophore illuminates the LR White resin, providing an image of tissue structure primarily in negative relief. Bar, 100µm. All subsequent FIRM images are shown in monochrome. FIGURE 2: Human anterior pituitary, formalin-fixed autopsy specimen, comparison of H&E paraffin section (left hand side) with FIRM (right hand side). 20x dry objective used for both. Scale bar, 20µm FIGURE 3: Human anterior pituitary, FIRM image taken with 150X glycerin objective. Individual secretory granules are easily visualised, ranging from 150- 350nm in diameter. Scale bar, 5µm. FIGURE 4: Correlative FIRM/fluorescent lectin labelling of human pituitary. A single LR White thin section was labeled first with Concanavalin AAlexa-488 (B) imaged, then stained for FIRM imaging (A). Densely granulated cells (arrows in A) are strongly labeled with Con-A (arrows in B). Dark structures resembling large lysosomes or residual bodies (arrowheads in A) are also strongly labeled by Con-A, consistent with reports in the literature. FIGURE 5: Comparison of toluidine blue staining (top) to FIRM (left) on near adjacent sections of human kidney. Note enhanced contrast, especially of glomerular basement membranes in FIRM image. High power FIRM image (C) of glomerulus. Note resolution of podocyte foot processes (arrows). Scale bars in A, B, 20µm, in C, 5µm. DISCLOSURE: James W. Mandell, MD, PhD, Dept. Pathology (Neuropathology), University of Virginia School of Medicine, Charlottesville, VA has filed a provisional patent relating to this invention: U.S. Provisional Patent Application Serial No. 61/298,759 Filed on January 27, 2010 Title: Compositions and Methods for Enhanced Specimen Contrast
Crystal Violet Stain For Amyloid
EMS26301 Components: Crystal Violet Solution Acetic Acid, 1% Aqueous Fixation: Absolute alcohol or 10% Formalin Sections: Paraffin Staining: Stain sections of fixed or unfixed tissue in Crystal Violet Solution for three to five minutes. Wash in Acetic Acid, 10%, followed by a thorough washing in water to remove all traces of acid. Examine in water or glycerine. The stain will keep for some time if sections are mounted in a saturated solution of potassium acetate or in levulose. Mount in Modified Apathy's Gum Syrup or Lieb's Abopon. Stain Results: Amyloid Violet-Red Tissue Blue References: Clark, G.: Staining Procedures, Williams and Wilkins Co., Baltimore 3 rd . ed., c. 1973,p.48. Conn , H.J., Biological Stains 4 th ed. Biotech Publications, Geneva New York , 1940.
Dane's Method for Prekeratin, Keratin, & Mucin
EMS26401 Components: Phloxine Solution 1% Alcian Blue 0.5% Orange G Solution Mayer's Haematoxylin Fixation: 10% Buffered Neutral Formalin Sections: Cut paraffin at 6µm sections Staining Procedures: Deparaffinise and hydrate slides to distilled water. Mayer Haematoxylin for 10 minutes. Blue in running tap water for 10 minutes. Rinse in distilled water. Phloxine B 1% Solution for 3 minutes. Wash in running tap water to remove excess stain. Rinse in distilled water. Alcian Blue, 0.5% for 5 minutes. Wash in tap water for 2 minutes. Rinse in distilled water. Orange G Solutionfor 13 minutes. Transfer to 95% alcohol, two changes, five dips each; absolute alcohol, two changes, fifteen dips each. Clear in Xylene and mount. Stain Results: Acid Mucopolysaccharides Blue Prekeratin and Keratin Orange to Red Nuclei Brown References: Dane, E.T., and Herman, D. L.: Stain Techn., 38:97, 1963 AFIP Manual of Histological Staining Methods, 3rd ed., Ed. L. Luna: NY, McGraw-Hill Publ., c. 1968, p.83.
Differential Quik III Stain Kit (Modified Giemsa)
EMS26096 The Differential Quik III Stain Kit is a high quality, rapid turnaround time staining kit that serves many purposes in the cost-conscious laboratory environment. The Differential Quik III Stain Kit may be used for fine needle aspirations, cytology, blood smears, bone marrow biopsies as well as bone marrow smears, touch preps, various microorgan-isms, as well as detecting the H.Pylori Microorganism. The kit contains a fixative for air-dried cell suspensions and/or touch preparation slides. Applications: Fine needle aspirations Frozen Sections Blood Smears Cytological specimens Cytopreps Bone marrow biopsy Touch preps Microorganisms Kit Contents: Fixative -Teal Solution I- Red/Orange Solution II - Blue Storage and handling: Storage: Room Temperature Safety: Refer to the MSDS for details. Procedure: Fixative: Fix frozen sections, touch preps, bone marrow biopsies and blood smears for 10 seconds inFixative. Solution I-Red: Dip 5 times in Solution I Solution II-Blue: Dip 5 times in Solution II Wash in Distilled water and air dry. Quickly dehydrate in 100% EtOH,then xylene and mount in a synthetic resin. Results: Structure Colour Erthrocytes Pink/yellowish red Platelets Dark blue/violet/purple granules Neutrophils Blue nucleus, pink cytoplasm, purple granules Eosinophils Blue nucleus, blue cytoplasm, red granules Basophils Dark blue purple nucleus, purple granules Monocytes Purple nucleus, light blue cytoplasm Bacteria/microorganisms Deep blue, vary in shape Collagen, cytoplasm & muscle Various shades of pink, blue, orange & yellow
Ehrlich’s Haematoxylin
EMS26753 Components: Ehrlich's Haematoxylin Solution Acid Alcohol 1% Sodium Acetate 1% Aqueous Ammonium Water 0.3% Lithium Carbonate, Saturated Fixation: Formalin 10% Neutral Buffered or fixative of choice Sections: Paraffin @ 5-6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Stain for 2 – 5 minutes in Ehrlich’s Haematoxylin. Wash in distilled water. Differentiate in Acid Alcohol until the sections turn red. Blue section in Sodium Acetate 1% or Ammonium Water or Lithium Carbonate Solution. Wash thoroughly and counterstain. References: P. Ehrlich "Fragekasten" ["Questionbox"] Zeitschr. für Wiss. Mikr. Techn. III,I:150, 1886. Article in German. Mallory, F.B., Pathological Technique. W.B. Saunders Co, Philadelphia, p. 73, 1938. Clark, G., (ed). Staining Procedures, 3rd edition, Williams & Wilkins, Baltimore, pp 33-34, 1973. Luna, L.G. (Ed), Manual of Histologic Staining Methods of the AFIP, 3rd Edition, McGraw-Hill, NY, pp 35 and 38, c. 1968.
Eosin Y Solution
Preparation: Prepare eosin Y stock solution. Add 2.0g of water-soluble eosin Y to 40ml of double-distilled H2O, and mix until dissolved. Then add 160ml of 95% ethanol, and mix. Store at room temperature. Prepare eosin Y working solution. Add 200ml of eosin Y stock solution to 600ml of 80% ethanol and mix well. While working in a fume hood, add 4ml of glacial acetic acid and mix well. Store covered at room temperature.
Fat Stains In Supersaturated Alcohol
EMS26503 Components: Sudan IV-Isopropanol Solution Oil Red O-Isopropanol Solution Sudan III-Isopropanol Solution Mayer's Haematoxylin Erhlich’s Haematoxylin, Diluted Sodium Phosphate 1% Aqueous Fixation: Formalin Sections: Frozen Preparing Working Stain Dilute 6ml of stock solution Sudan IV in Isopropanol or Oil Red O in Isopropanol or Sudan III Isopropanol solution with 4 ml of distilled water. Allow to stand for 5-10 minutes and then filter. The filtrate may be used for several hours Procedure Stain thin frozen sections with the working fat stain for 10 minutes. Wash in tap water. Stain for 5 minutes in either Mayer’s Haematoxylin or Ehrlich’s Haematoxylin diluted. Place in Sodium Phosphate 1 %, or in tap water until blue. Float out in water, take up on a slide and mount on Apathy’s Gum Syrup or Zwemer’s Glychrolgel or any other soluble aqueous medium. Stain Results: Nuclei Blue Fat Orange-Red Erthrocytes Sometimes Green Cytoplasm Lighter Green References: Clark, G, ed.: Staining Procedures, 3rd ed.: Baltimore: Williams & Wilkins Co., 1973, p. 149
Ferritin Cationised for Labelling Negative Charges
EMS15550 Ferritin Cationised Specifications Quantity: 10mg per ml Form: Dissolved in 0.15M NaCl sterile solution Protein Concentration: 1% 10.5 mg/ml based on E 270nm = 79.9 as for native ferritin. Cationised ferritin is prepared according to the publication of D. Danon et. al. Journal of Ultrastructure Research 38, 500-510 (1972). The Nucleophile used for the coupling is DMPA (N,N-dimethyl-1, 3 propanediamine). Storage: Store at 4°C Instructions for use: Labelling of negatively charged cell membranes for visualisation in the electron microscope is carried out by using the cationised ferritin solution (0.1ml), diluted to 0.5ml with Veronal-HCl buffered saline pH 7.2. This working dilution is added to a 10% cell suspension (1ml) before or after a short fixation with glutaraldehyde. Shake the mixture gently and allow to stand at ambient temperature. The optimal incubation for unfixed cells is 5 minutes; longer periods can give rise to toxic effects (the incubation and temperature conditions may be changed according to the requirements of the experiment). Wash the cell suspension with Veronal-HCl buffered saline and prepare for electron microscopy. Note: Laboratory reagent - not to be administered to humans nor used for any drug purposes.
Genta, Robason and Graham Stain for Simultaneous Visualisation of Helicobacter Pylori and Gastric Morphology
EMS26116 Components: Uranium Nitrate Silver Nitrate 1% Aqueous Gum Mastic 2.5% Hydroquinone 2% Silver Nitrate 0.04% Alcian Blue, pH 2.5 Harris Hematoxylin Acid Alcohol 1% Ammonia Water 0.25% Eosin Y 1% Fixation: Formalin fixed Sections: Paraffin @ 4µm sections Staining: Deparaffinise and hydrate sections to distilled water. Sensitise sections by placing them in Uranium Nitrate. 1%, Aqueous at room temperature for 3 minutes and transfer them to distilled water. Rinse slides in distilled water until the possibility of cross contamination is eliminated. Place sections in Silver Nitrate, 1% at room temperature and then heat then in a microwave oven to just below the boiling point. DO NOT BOIL! Rinse slides in 3 changes of distilled water. Rinse in 2 changes of 95% alcohol. Rinse in 2 changes of 100% alcohol. Place slides in 2.5% Gum Mastic for 5 minutes. Air dry sections for 1 minute Rinse in 2 changes of distilled water. Reduce in reducing solution in a 450°C water bath for 10 to 15 minutes or until sections have developed satisfactorily with black or dark brown H-Pylori and a light yellow background. Check periodically under microscope. To prepare reducing solution - Gum Mastic, 2.5 % - 10 ml - Hydroquinone, 2% - 25 ml - Absolute Alcohol - 5 ml - Silver Nitrate, 0.04% - 2.5 ml Mix together - DO NOT FILTER!! Rinse sections in water to stop reduction. Stain in Alcian Blue, pH 2.5 for 10 minutes. Rinse in running tap water. Stain sections in Harris' Haematoxylin for 5 minutes. Rinse in running tap water. Quickly dip in Acid Alcohol. Rinse in running tap water. Rinse in Ammonia Water, 0.25% until blue. Rinse in running tap water. Stain sections in Eosin Y, 1% for 5 minutes. Quickly rinse in tap water. Rinse twice in 95% alcohol. Rinse three times in absolute alcohol. Rinse three times in Xylene and then coverslip. Staining Results: Mucus cells Brownish yellow Lamina propria ground substance Light yellowish or greyish Smooth muscle fibers Light pink Cytoplasm of neutrophils Pale pink Eosinophils Intense red Nucleus Dark brown Intestinal metaplasma Bright blue H-pylori are easily recognised in the mucus, on the cell surface, and within the gastric pits. References: Genta PM., Robason GO, Graham DY- Simultaneous Visualization of Helicobactor Pylon and Gastric Morphology A New Stain Hum Pathol 25: 22 1-226, 1994.
Giemsa for Helicobactor Pylori and Mast Cells
EMS26110 Components Methanol Giemsa Stock Solution Acetic Acid 1% Fixation 10% Buffered Neutral Formalin Sections 3µm Procedure Deparaffinise and hydrate to distilled water. Place promptly in Methanol for 2 minutes. Preheat Working Geimsa solution in microwave on high for 45 seconds. Filter the hot solution into a coplin jar containing the slides. Stain for 2 minutes. To prepare working solution mix: • 7.0ml Geimsa Stock • 38ml Distilled water • 5ml Methanol Prepare fresh- do not reuse!! Rinse quickly in distilled water Dip one slide at a time in Acetic Acid 1% . Rinse quickly in distilled water. Rinse in 3 changes of Methanol. Clear in Xylene and mount. Stain Results Red Blood Cells Pink Nuclei Blue H-Pylori, fungi and other bacteria Blue Mast Cells Purple Rickettsia Purple NOTE: The use of methanol to dehydrate stained sections prior to mounting seems to provide lighter staining of surrounding tissue, offering better contrast with stained organisms. References Helicobacter pylori in peptic ulcer disease, NIH Consensus Statement Online, January 1994, 12 (1): 1:23 . Luna l AFIP 3 rd ed. McGraw Hill, 236. Fail, Rena , AS , HT (ASCP) Medical University of South Carolina , Histologic . Vol. XXXIV, No. 2, Nov. 2001. P 32.
Giemsa Thick Film Stain
EMS26154 Components: Giemsa Thick Film Solution Giemsa Stock solution Phosphate Buffer Fixation: None. Films are prepared by spreading 3 – 5 drops of blood on a scrupulously clean slide to form a circle of about 15mm diameter: the films are air dried by keeping in a horizontal position for 18 – 24 hours at room temperature, protected from dust and insects. Staining: Stain 40-120 minutes in Giemsa Thick Film Staining Solution. To prepare Giemsa Thick Film Staining Solution from Giemsa Stock Solution, mix before use: Giemsa Stock Solution - 1ml Phosphate Buffer - 2ml Distilled Water - 47ml Wash 5-10 minutes in distilled water or Phosphate Buffer Dry in the air and examine. Stain Results: Malarial Parasites Clear Red Chromatin Cytoplasm Clear Blue Red Corpuscles Not seen due to lacking of hemoglobin during drying References: Clark, G., Staining Procedures, Williams and Wilkins Co. Baltimore, 3rd, c. 1973, p. 126 Lillie, R. D. Histologic Technic and Practical Histochemistry, 3rd ed., Mcgraw-Hill, New York, 1965. Barber, M.A., and Komp, H.W., Pub. Health Rep., 44:2330-41, 1929
Giemsa Thin Film Stain
EMS26153 Components: Giemsa Thin Film solution Giemsa Stock solution Phosphate Buffer Fixation: Streak a drop of blood, leukocytes, rickettsiae culture, pus or other exudates over a sterile glass slide by means of another slide or cover glass. Fix film in Methyl Alcohol for 2-3 minutes. Staining: Place in Ethyl or Methyl Alcohol for 5 – 7 minutes. Air dry. Stain in the Giemsa Thin Film Staining solution*. Because different cells accept the stain at varying rates, prepare 3 – 4 sample slides and stain from 15 – 20 minutes. Some membranous structures (Trypanosomes) may require 1 – 2 hour periods. To prepare Giemsa Thin Film Staining solution from Giemsa Stock solution, mix before use: Giemsa Stock Solution - 1ml Phosphate Buffer - 50ml Wash in distilled water and air dry. Stain Results: Nuclei, additional Leukocyte features are similar to Wright Stain results. Reddish Purple Plasmodic cytoplasm Blue Chromatin Red References: Clark, G., Staining Procedures, Williams and Wilkins Co. Baltimore, 3rd, c. 1973, p. 125
Gold Chloride Hydrate
EMS16580 To make 100ml of GOLD SOLS prepare two separate solutions in CLEAN GLASSWARE Components: Sodium Citrate Tannic Acid Potassium Carbonate Solution A: Gold Chloride solution (Use 250ml beaker) 1ml 1% (w/v) Gold Chloride in deionised water 79ml deionised water Solution B: Reducing solution (Use 50ml beaker) 4ml 1% Sodium Citrate, dihydrate (FW. 294.11) Make fresh monthly 1% Tannic Acid make fresh monthly 25mM Potassium carbonate (K2C03) Make fresh on day of use (Volume used is equal to the volume of Tannic Acid) Deionised water to bring to a final volume of 20ml The final diameter of the gold particle is inversely proportional to the volume of Tannic Acid solution added. Some common sizes and the volume of Tannic Acid solution to be added are: Gold Particle Size Volume of Tannic Acid 15nm 0.022ml 10nm 0.100ml 5-6nm 0.900ml Procedure Heat Solution A and B to exactly 60°C in a water bath; cover the solution to eliminate evaporation during heating. Remove Solution A to a hot plate (which is heated sufficiently to boil water); begin mixing vigorously and immediately add the entire contents of Solution B in one quick motion to the beaker of Solution A. Continue to mix the solution until it reaches 100°C. (It may be necessary to reduce the vigour of the stirring in order to achieve 100°C). Try to minimise evaporation during the heating to 100°C. After the solution has been at 100°C for 2 - 5 minutes, remove and allow to cool. Check the volume and adjust to 100ml with deionised water. Store at 4°C in a clear glass bottle, appropriately labeled as to the size of the particle, date and maker.
Goldner's Trichrome Method
EMS26386 Components Bouin’s Fluid Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B Ponceau Acid Fuchsin Acetic Acid, 1% Phosphomolybdic Acid Orange G Solution Light Green Stock, 0.2% Fixation 10% Buffered Neutral Formalin or Bouin's Solution Sections Paraffin, frozen or celloidin Microtomy Cut paraffin sections at 6 micrometers. Procedure Decerate slides in Xylene, 2 changes, 2 minutes each. Place in absolute alcohol, 2 changes, 2 minutes each; 95% alcohol, 2 changes, 2 minutes each; then rinse in distilled water. Mordant in Bouin's Fluid solution for 1 hour at 56°C, or overnight at room temperature, if formalin fixed. Cool and wash in running tap water until yellow color disappears. Rinse in distilled water. Place in Weigert's Haematoxylin for 10 minutes. ***To prepare Weigert's Solution, mix equal parts of: Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B *** stir vigorously. Wash in running tap water for 10 minutes. Rinse in distilled water. Stain in Ponceau Acid Fuchsin for 5 minutes. Wash in Acetic Acid, 1%. Place in Phosphomolybdic Acid-Orange G solution until collagen is decolorized. Rinse in Acetic Acid, 1% for 30 seconds. Stain in Light Green Stock Solution for 5 minutes. Rinse in Acetic Acid, 1%, for 5 minutes. Blot sections but do not allow to dry. Dip quickly in absolute alcohol. Dehydrate slides in 95% alcohol and absolute alcohol 3 changes each. Clear in Xylene, 3 changes. Mount coverglass with appropriate medium (refractive index 1.48-1.56). Stain Results Nuclear chromatin Brownish black Cytoplasm Bright red Erythrocytes Orange Muscle Red Collagen Green References Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts Johnson Printers, Downers Grove , IL p 151-152, c 1992.
Gomoris’ Methenamine Silver for Urate Crystals
EMS26614 Components: Silver Nitrate 5% Methenamine 30% Sodium Borate 5% Gold Chloride 0.1% Sodium Thiosulfate 3% Light Green Working Solution Fixation: Absolute Alcohol at 4°C for overnight. Sections: Paraffin @ 4-5µm Staining Procedures: Deparaffinise to Xylene and rinse with 3 changes of absolute alcohol. Place slides in preheated working* Methenamine Silver. Incubate for 30 minutes @60°C. To prepare working solution: Methenamine-Silver Nitrate Stock Solution: Silver Nitrate, 5% - 5ml Methenamine, 3% - 100ml Methenamine-Silver Nitrate Working Solution Methenamine Silver-Nitrate Stock Solution - 25ml Distilled water - 25ml Sodium Borate, 5% - 2ml Prepare fresh prior to use. DO NOT USE IF CLOUDY. Urate Crystals should be black Rinse sections in distilled water. Tone with Gold Chloride, 0.1% for 5 minutes. Give sections 4 or 5 rinses of distilled water. Sodium Thiosulfate, 3% for 5 minutes. Wash in tap water for 5 minutes. Rinse in distilled water. Counterstain in Light Green Working Solution for 2 minutes. Dehydrate, clear and mount with Permount or any synthetic resin Stain Results: Urate Crystals Black Background Green References: Sheehan, Denza C. and Hrapchak, Barbara B. Theory and Practice of Histotechnology, 2nd ed. The C.V. Mosby Company, St. Louis, p.225 – 226, c. 1980.
Gridley's Method for Endamoeba Histoslytica
EMS26109 Components: Harris Haematoxylin Weigert's Iron Haematoxylin Solution A Weigert's Iron Haematoxylin Solution B Analine Eosin Solution Naphthol Green B Stain 1% Acid Alcohol 1% In 75% Alcohol Ammonia Water 0.3% Fixation: 10% Buffered Neutral Formalin Section: Paraffin at 6µm Staining Procedure: Deparaffinise and hydrate to distilled water. Include a control slide with all samples. Stain in Harris Haematoxylin 10 min. or Weigert's Iron Haematoxylin for 3 min. To prepare Weigert's Iron Hematoxylin, mix equal parts just before use: Weigert's Iron Haematoxylin Sol'n A and Weigert's Iron Haematoxylin Sol'n B. Wash in running water for several minutes, differentiate each slide in Acid Alcohol, 1% and then wash again. Blue each slide in Ammonia Water and wash in running water. Stain in Aniline-Eosin for 5 minutes, rinse well in distilled water. Slides should appear deep rose. Counter-stain in Naphthol Green B for 5 minutes. Under a microscope, differentiate the slides in 95% alcohol (two changes) until the erythrocytes remain a deep rose. Dehydrate through two changes absolute alcohol and clear in two changes Xylene. Mount. Results: Erythrocytes (ingested) rose Connective tissue green Amoebae blue-green Nuclei (amoebic) darker blue-green References: Gridley, M.F.: Am. J. Clin., Pathol., 24:243 (1954). Luna, L.G.: Histologic Staining Methods, 3rd ed.: New York: McGraw-Hill Book Co., c. 1968, p. 228.
Gridley's Method for Fungi
EMS26120 Components: Chromic Acid 4% Coleman's Feulgen Aldehyde Fuchsin Metanil Yellow 0.25% Fixation: 10% Buffered Neutral Formalin Sections: Paraffin at 6µm sections Staining: Use a control slide Deparaffinise and hydrate to distilled water. Oxidise in Chromic Acid, 4% for one hour. Wash in running water for five minutes. Coleman’s Feulgen Reagent for 15 minutes. Wash in running tap water for 15 minutes. Rinse in several changes or 70% Alcohol. Aldehyde Fuchsin for 30 minutes. Rinse off excess stain with 95% alcohol Rinse in distilled water. Counterstain lightly with Metanil Yellow. Rinse in distilled water. Dehydrate in 95% alcohol, absolute alcohol, and clear in two changes of Xylene. Mount. Stain Results: Mycelia, Conidia Deep purple Elastic Tissue & Mucin Background Yellow Filaments of Nocardia & Actinomyces are not stained by this method. References: Gridley, M.F. – Amer. J. Clin. Path., 23:303 (1953) AFIP – Manual of Histologic Staining Techniques: 3rd, ED. ED. L. Luna; New York: McGraw-Hill Publications, c 1968, p. 229.
Haematoxylin And Eosin / Methenamine Silver Stain
EMS26123 For The Histological Diagnosis Of Fungi In Tissue Sections Contains Fungi in tissue, includes: Chromic Acid, 10% Sodium Bisulfite, 1% Silver Nitrate, 5% Aqueous Methenamine, 3% Aqueous Sodium Borate, 5% Aqueous Gold Chloride, 0.2% Sodium Thiosulfate, 0.2% Aqueous Harris’ Hematoxylin Acetic Acid 3% Ammonia Water, 0.25% Eosin Y, 1% Alcoholic Fixation 10% Buffered Neutral Formalin (NBF) Sections 3-4µm Staining Procedure Deparaffinise and hydrate to distilled water. Oxidize in Chromic Acid, 10% for 10 minutes. Rinse with 3-4 changes of tap water. Rinse in Sodium Bisulfite, 1% for 1 minute to remove any residual Chromic Acid. Wash in running water, 5-10 minutes. Rinse with 3-4 changes of distilled water. Place in freshly mixed working methenamine Silver Nitrate Solution in a water bath @ 58-60 °C, for 15-25 minutes or until sections turn yellowish brown. Methenamine Silver Nitrate Stock: • Silver Nitrate, 5% Aq - 5ml • Methenamine, 3% Aq - 100ml Use chemically clean glassware. Shake well to dissolve precipitate. Store in refrigerator. Then to prepare Methenamine Silver Nitrate Working solution: • Methenamine Silver Nitrate Stock Solution - 25ml • Water - 25ml • Sodium Borate, 5% Aq - 4ml Rinse in 6 changes of distilled water. Tone in Gold Chloride, 0.2% for 2-5 minutes. Rinse in 3-4 changes of distilled water. Remove unreduced Silver with Sodium Thiosulfate, 2% for 2-5 minutes. Wash thoroughly in tap water for 10 minutes. Place in Harris' Haematoxylin for 10 minutes. Rinse in 3-4 changes of tap water. Place in Acetic Acid, 3%) for 1 minute. Rinse in 2 changes tap water. Place in Ammonia Water, 0.25% 10 minutes. Rinse in 2 changes tap water. Place in 95% alcohol, 1 minute. Place in Eosin Y, 1% Alcoholic for 5 minutes. Dehydrate in 95% alcohol , absolute alcohol and clear in xylene, 2 changes each. Mount with Permount. Stain Results Organisms Blue-Black Background Rose References Lamps, Laura W. et al A Combined Hematoxylin and Eosin/ M ethenamine Silver Stain for the Histological Diagnosis of Fungi in Tissue Sections. , J. of Histotechnology 23:4 p. 341-345
Hall’s Method for Bilirubin
EMS26611 Components: Fouchets Reagent Van Gieson's Solution Fixation: 10% Buffered Neutral Formalin Sections: Paraffin @ 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Place in Fouchet’s Reagent for 5 minutes. Wash in running water, then in distilled water. Stain for 5 minutes in Van Gieson’s Solution. Dehydrate in 95% Alcohol, absolute alcohol, and clear in xylene, two changes each. Mount with Permount. Stain Results: Biliverdin Green Collagen Red Muscle Yellow References: Hall, M.J., Amer. J. Clin. Path., 34:313-316, 1960. Luna, L.G. (ed). Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology, 3rd ed., McGraw-Hill, N.Y., c. 1968, p.174.
Harris' Haematoxylin
EMS26041 Fixation Follow most fixatives. Sections Paraffin Staining Deparaffinise and hydrate to distilled water.. Stain in Harris Haematoxylin containing 2 – 4ml of Glacial Acetic Acid per 100ml for 5-15 minutes. Wash in water. Differentiate in Acid Alcohol until the sections turn red. Blue section in Sodium Acetate, 1% or Ammonia Water, 0.3%, or Lithium Carbonate Solution Wash thoroughly and counterstain. References Harris, H.F., J. Appl. Microscopy, III: 777-781, 1900 Mallory, F. B., Pathological Technique, W. B. Saunders Co., Philadelphia, 1938. Clark, G., (ed.), Staining Procedures, 3rd Edition, William & Wilkins, Baltimore, pp33 & 35, c 1973. Luma, L.G., (ed.) Manual of Histologic Staining Method of the AFIP, 3rd edition, McGraw-Hill, NY, pp 34 & 38, c1968.
Harris' Haematoxylin (1900)
EMS26754 Haematoxylin with or without mercury Components: Harris' Haematoxylin Solution Harris' Haematoxylin Without Mercury Acid Alcohol 1% Sodium Acetate 1% Aqueous Ammonia Water 0.3% Lithium Carbonate Saturated Solution Fixation: Formalin 10% Neutral Buffered or fixative of choice Sections: Paraffin @ 5-6µm Staining Procedures: Deparaffinise and hydrate to distilled water Stain in Harris' Haematoxylin containing 2-4ml Glacial Acetic Acid per 100ml for 5 – 15 minutes. Wash in water. If using Harris; Haematoxylin without Mercury containing 2-4ml Glacial Acetic Acid per 100ml for 5 – 15 minutes. Wash in water. Differentiate in Acid Alcohol until the sections turn red. Blue section in Sodium Acetate, 1% or Ammonium Water or Lithium Carbonate Solution. Wash thoroughly and counterstain. References: Harris, H.F., J. Appl. Microscopy, III: 777-781, 1900. Mallory, F.B., Pathological Technique. W.B. Saunders Co, Philadelphia, 1938. Clark, G., (ed). Staining Procedures, 3rd edition, Williams & Wilkins, Baltimore, pp 33-35, 1973. Luna, L.G. (Ed), Manual of Histologic Staining Methods of the AFIP, 3rd Edition, McGraw-Hill, NY, pp 34 and 38, c. 1968.
Herxheimer Technique for Fat
EMS26500 Components: Sudan IV Staining Solution Harris' Haematoxylin Acetic Acid 1% Aqueous Fixation: Formalin Sections: Frozen Staining Procedures: Immerse in 70% Alcohol for an instant. Stain for 2 – 5 minutes in Sudan IV Staining Solution. Wash quickly in 70% Alcohol. Rinse in distilled water. Counterstain in Harris Haematoxylin. Follow with 2 – 5 minutes in Acetic Acid, 1% for clearer blue staining of nuclei. Wash thoroughly in tap water. Mount (Glycerin or Glycerin Jelly. Stain Results: Nuclei Blue Fat Orange-Red Cholesterol Red References: Clark, G, ed.: Staining Procedures, 3rd ed.: Baltimore: Williams & Wilkins Co., 1973, p. 143 Conn’s, H.J.: Biological Stains; 8th Ed., Baltimore: Williams and Wilks Co., c. 1969, p. 100
Hirano-Zimmerman for Nerve Cells & Fibres
EMS26698 Components: Silver Nitrate 10% Aqueous Ammonia Water 0.1% Formalin Solution 50% Gold Chloride 0.05% Aqueous Sodium Thiosulfate 5% Aqueous Fixation: Formalin 10% Neutral Buffered Sections: Paraffin @ 12 – 15µm Staining Procedures: Deparaffinise and hydrate to distilled water. Silver Nitrate, 10% for two hours or longer. Place sections directly in Ammonia Water for 3 minutes. Place section directly in Formalin Solution 3 minutes. Tissue becomes black-brown. Rinse in distilled water, two changes. Place section again into Silver Nitrate, 10% for 3 – 5 minutes. Repeat step 3 and 4. Tissue becomes darker in color. Tone in Gold Chloride Solution for 20 to 30 minutes. Place sections directly into Sodium Thiosulfate, 5% for 1 minute. Rinse in distilled water. Dehydrate in 95% alcohol, absolute alcohol and clear in Xylene two changes each. Mount with Permount or any synthetic mounting medium Stain Results: Nucleolus and Nuclear Membrane Black Neurofibrils, Dendrites and Axis Cylinders Black Cytoplasm of Astrocytes and the Cytoplasmic membrane of Macrophages Grey Senile Plaques Black Various lipid granules are unstained, but each granule is clearly outlined. References: AFIP, Manual of Histological Staining Methods, 3rd ed., Ed. L. Luna: New York: McGraw-Hill Publications. C. 1968, p. 198. Hirano, A. and Zimmerman, H.M., Arch. Neurol., 6:114-122, 1962.
Jenner's Stain Solution
EMS26033-05 Specimen Collection Fresh blood film or fresh EDTA anticoagulated blood film and bone marrow films. Specimens should be air dried after smear has been prepared and fixed in absolute methanol for 15 seconds. Mode of Action The Jenner Stain Solution is a mixture of several thiazin-dyes in a methanol solvent. Ionic and nonionic forces are involved in the binding of these dyes. The staining solution has anionic and cationic properties. The negatively charged phosphoric acid groups of DNA attract the purple polychromatic cationic dyes to the nuclei. The blue basophilic granules are stained by the polychromatic cationic dyes. Cationic cellular components, such as erythrocytes and eosinophilic granules, are stained by the red and pink anionic dyes. The buffers used in the staining procedure liberate and activate dye ions allowing them to chemically bond with specific cellular components. When staining blood and bone marrow smears, the pH of the staining solution and/or buffer is a critical factor. Technical Procedures Immersion Staining Protocol Thoroughly dry blood or bone marrow smears. Fix smears in absolute methanol for 15 seconds to 5 minutes. Stain smears in Jenner Solution for 2 minutes. Stain in mixture of 50ml of Jenner Stain Solution, 75ml of pH 6.6 Phosphate Buffer solution and 175ml deionised water for 5 minutes. Rinse in standing deionised water for 1.5 minutes, or rinse briefly in running deionised water. Air dry smears. Examine smears under a microscope. Horizontal Staining Protocol Place slide with thoroughly dried film on a horizontal staining rack. Flood smear with absolute methanol for 15-30 seconds and then drain. Flood smear with 1ml Jenner Stain Solution and let stand for 3 minutes. Add 1ml of pH 6.6 Phosphate Buffer solution and 1ml deionised water to smear and let stand for 45 seconds. Rinse briefly with running deionised water. Air dry and examine under a microscope. Results Erythrocytes - Pale Pink Eosinophilic Granules - Reddish Orange Leukocyte Nuclei - Purple Cytoplasm - Bluish Purple Neutrophilic Granules - Light Purple Discussion Jenner Stain Solution should be stored at room temperature. This staining reagent is for "In Vitro" use only. Refer to the Safety Data Sheet for Health and Safety Information. All reagents are stable and should not form precipitants under ordinary storage parameters. It is recommended that the Jenner Stain Solution be discarded after each use. All dyes used in the formulation have been certified by the Biological Stain Commission. Technical Comments Thicker films and bone marrow preparations will require longer staining times. Because there is variation in the pH tap water use of tap water in procedure may hinder staining results. Distilled or deionised water should be used during the staining procedure. The "ripening" of the polychromed dye is a continuous chemical reaction. Therefore, the stock solution should not be used or diluted after the expiration date. The staining procedure may require modification to suit personal preference. Blood films which have not been thoroughly air dried before staining may show sloughing of cells from slide. References Raphael, S.S. Lynch's Medical Laboratory Technology, Fourth Edition. Saunders Company, Philadelphia, PA, 1983. Lillie, R.D. H. J. Conn's Biological Stains, Ninth Edition. Williams and Watkins Company, Baltimore, MD, 1977. Brown, B.A. Hematology: Principles and Procedures, Fourth Edition. Lea and Febiger Company, Philadelphia, PA, 1984.
Johnson's Method For Iron (Microincineration)
EMS26603 Components Potassium Ferrocyanide Solution 5% Hydrochloric Acid 5% Fixation 10% Buffered Neutral Formalin Sections Paraffin at 6µm Staining A. Microincineration Place unstained paraffin section uncovered on a sheet of asbestos and heat cautiously, using the outer cone from the flame of a Fisher or Meker burner. Keep the flame in motion and heat the slide as uniformly as possible. Continue the heating until the specimen, which turns brown then white, has almost completely disappeared. Immediately cover the slide with an inverted pan, (e.g. a pie pan) and allow the slide to cool gradually.to rapid cooling may cause the slide to break. After the slide has cooled, the completely incinerated specimen is examined under strong oblique lighting with dark field illumination, with or without coverslip. Results Iron oxide imparts a bright yellow-orange to dark red color to the ash. Specimen must be completely incinerated as carbon particles can also give a yellowish to red color. B. Staining of Incinerated Specimen Dip incinerated specimen in a thin celloidin solution (approx. 0.015% in alcohol: ether, 1:1). Air dry slides for a few minutes and then quickly hydrate to distilled water. Stain in working hydrochloric acid-potassium ferracyanide solution* for 15 minutes. Rinse in several changes of distilled water. *Prepare working stain by mixing equal parts of 5% hydrochloric acid and 5% potassium ferracyanide just before use. Air dry or dry in a 60°C oven. Mount with Permount. C. Direct observation of Prussian Blue Reaction on the Ash Create a well paraffin around the specimen. Pour working Hydrochloric Acid-Potassium Ferracyanide Solution (for preparation, sheet step B-2) into the paraffin well and observe under the microscope. Staining Results (B & C) Iron Oxide Blue References Fenton, R.H., Johnson, F.B., and Zimmerman, L.E., J. Histo-chem. Cytochem ., 12:153:155, 1964. L una, L.G., (ed.), Manual of Histologic Staining Methods of the AFIP , 3 rd edition, McGraw-Hill, NY., p. 181., c 1968.
Jones' Method for Kidney
EMS26396 Components: Periodic Acid 0.5% Aqueous Methenamine Solution 3% Aqueous Silver Nitrate 5% Aqueous Borate Buffer Working Solution Gold Chloride 0.2% Aqueous Harris Haematoxylin Alcoholic Eosin Y Stain Solution Sodium Thiosulfate 3% Aqueous Potassium Ferricyanide 0.5% Sodium Metabisulfite 3% Aqueous Acid Alcohol 1% Ammonia Water 0.3% Fixation: 10% Buffered Neutral Formalin or Russell’s Zenker. Sections: Cut paraffin at 2µm sections. Staining Procedures: Deparaffinise and hydrate slides to distilled water. Oxidise in Periodic Acid, 0.5% for 11 minutes. Wash well in chloride-free water. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3%, 2.5ml Silver Nitrate, 5% and 12.0ml Borate Buffer, pH 8.2 Place slides in the solution and the entire jar in a water bath at 70°C for approx. 60 – 75 minutes. Check under microscope when slides appear medium brown microscopically. Every 10 minutes, once the medium brown color has been established, rinse slide in 70°C chloride-free water and check under microscope. Rinse again in hot water and return to hot staining solution. As the staining time approaches the end point, check the slides, as above, every 1 – 2 minutes. The entire procedure must be performed quickly to prevent an uneven staining of the tissue. The slides should exhibit a brownish-yellow background, intense black reticulum fibers, and black basement membranes. If the slides become oversaturated, i.e. too black, de-stain in a dilute Potassium Ferricyanide Solution, 0.5% for one or two dips. Rinse well in distilled water. Tone in Gold Chloride, 0.2% for 1 minute. If sections are over-toned, place in Sodium Metabisulfite, 3% for 1 – 2 minutes. Rinse well in distilled water. Place in Sodium Thiosulfate, 3% for 1- 2 minutes. Wash in running tap water, 10 minutes. Rinse well in distilled water. Stain in Harris’ Haematoxylin containing 2 – 4 ml of Glacial Acetic Acid per 100 ml for 5 -15 minutes. Wash in water. Differentiate in Acid Alcohol, 1% until the sections turned red. Blue section in Ammonia Water, 0.3%. Wash thoroughly. Counter-stain in Eosin Y, 1%, Alcoholic Solution Dehydrate in 95% and 100% alcohol, and clear in Xylene, 3 changes each. Mount. Stain Results: Basement membranes, reticulum fibres solution Black Nuclei Blue Cytoplasm, collagen, connective tissue Pink-Orange References: Jones, D.B., Amer. J. Path. 27:99 (1951). AFIP Manual of Histological Staining Methods, 3rd ed., Ed. L. Luna: NY, McGraw-Hill Publ., c. 1968, p.97.
Kinyoun's Method For Acid Fast Bacteria, Microwave
EMS26087 Components Kinyoun's Solution Acid Alcohol, 1% Methylene Blue Working Solution Section Paraffin and frozen sections, air dried smears, touch preps. Procedure Deparaffinise sections in 2 changes of Xylene, and absolute alcohol. Air dry slides. Filter 40ml Kinyoun's Carbol Fuchsin Solution, into a plastic Coplin jar. Place slides in the jar; cover with a loose screw cap. Heat in the microwave for 50 seconds. Remove the Coplin jar from the oven; rinse slides individually in 50% Alcohol. Differentiate in Acid Alcohol, 1%, until sections are faint pink. Dip a few times. Rinse in 50% alcohol, then in distilled water. Counterstain lightly with Methylene Blue Working Solution, for two seconds. To prepare working solution from Methylene Blue Stock Solution, mix: Methylene Blue Stock -10 parts Tap Water - 90 parts Dehydrate clear in Xylene. Mount Note: All the alcohol dilutions used in this method should be prepared with absolute 100% alcohol. Stain Results Acid Fast Bacteria Red Other tissue elements Blue Other types of bacteria Blue References Valle, Sylvia; Journal of Histology , vol. 9, No. 4, December 1986.
Kluver-Barrera Method for Myelin and Nerve Cells
EMS26681 Components: Luxol Fast Blue MBS Solution 0.1% Cresyl Violet Acetate Solution 0.1% Acetic Acid 10% Aqueous Lithium Carbonate Solution 0.05% Fixation: Formalin, 10% Buffered Neutral Sections: Paraffin @ 15 to 20µm Staining Procedure: Deparaffinise and hydrate to 95% alcohol. Leave in Luxol Fast Blue Solution in 56°C oven overnight. Rinse off excess stain with 95% alcohol. Rinse in distilled water. Differentiate the slides singly in Lithium Carbonate Solution for 30 seconds. Continue differentiation in 70% alcohol until the grey matter is clear and white matter sharply defined. Check microscopically. Repeat the differentiation if necessary starting at step 5. When differentiation is complete, place in distilled water. When all slides have been collected in distilled water, add fresh distilled water. Counterstain in (Cresyl Violet Acetate),Working solution for 6 minutes. To prepare working solution add 15 drops of Acetic Acid, 10% to 100ml of Cresyl Violet Acetate 0.1% just before use. Filter. Rinse in 2 changes of 95% alcohol. Continue the dehydration through two changes of absolute ethyl alcohol and xylene, two changes each, for 2 minutes. Mount with resinous medium. Stain Results: Myelin, including phospholipids Blue to Green cells Cells and cell products Pink to violet NOTE: The Luxol Fast Blue procedure works well when combined with other procedures such as the Bodian Method. References: Kluver, H., and Barrera, E. : A method for the combined staining of cells and fibers in the Nervous system. J. Nueropath. Exp. Neurol. 12:400-403, 1953. AFIP, Lab Methods in Histothechnology, ed. Edna B. Prophet, Bob Mills. Jacquelyn B. Arrmyl, Leslie H. Sobin, M.D. Wash, D.C. American Registry of Pathology, c. 1992, p. 94-95.
Lacquers's Method for Alcoholic Hyalin
EMS26306 Components Mayer's Haematoxylin Acid-Fuchsin Aniline Solution Light Green 1% Aqueous Phosphomolybdic Acid 1% Alcoholic Picric Acid Fixation: 10% Buffered Neutral Formalin or Zenker’s Sections: Cut Paraffin, 6µm Staining Procedures: Deparaffinize and hydrate to distilled water. Stain in Mayer Haematoxylin for 15 minutes. Wash in running water for 15 minutes. Flood slides with Acid Fuchsin-Aniline Solution. Shake bottle well before using, solution will separate upon standing, and heat gently until aniline fumes escapes, view against a dark background. Let stand for 15 minutes. Wash in water, 3 dips. Differentiate in Alcoholic Picric Acid until only hyaline and red cells remain red, and collagen is faint gray or unstained. Wash in running water for 3 minutes. Mordant in 1% Phosphomolybdic Acid for 4 to 18 hours. Counterstain in Light Green Solution for 1 hour. Wash in running water for 1 minute. Fibers are seen as distinct fibrils. Dehydrate in 95% alcohol, absolute alcohol, and clear in xylene, two changes each. Mount with Permount. Stain Results: Mallory Bodies Bright Red Erythrocytes Red Cystoplasm Pale Brown Bile Pigment Green Protinaceous material seen occasionally in some liver cells Red Hemosiderin and lipofichsin Unstained References: Lacquer, G.I., Amer. J. Clin. Path., 20:689-690, 1950. Luna, L.G. (ed), Manual of Histologic Staining Methods of the AFIP, 3rd edition, McGraw-Hill, NY, c 1968 Lillies, R.D. Fullmer, H.M., Histopathologic Technic and Pratical Histochemistry 4th ed., McGraw-Hill, N.Y. 1976, pp 670-671. Clark, G., (ed). Staining Procedures, 3rd edition, Williams & Wilkins, Baltimore, p. 52, c.1973.
Leukocyte Alkaline Phosphatase
EMS26913 Components Buffered Acetone Fixative Naphthol AS-BI Phosphate Dimethyl Formamide Fast Red Violet Salt LB Propanediol Buffer Haematoxylin Counter Stain Sample Capillary Blood Or Heparinized Venous Blood Films Fixative Buffered Acetone Fixative Procedure Fix blood films in Buffered Acetone Fixative for 30 seconds. Wash Blood films in running tap water for 30-60 seconds. Air dry or blot. Stain films for exactly 10 minutes in staining Solution. *To prepare staining solution: Dissolve 5mg of Naphthol AS-BI Phosphate in 0.2 to 0.3ml of Dimethyl Formamide. Add 60m of Propanediol Buffer. Add 40mg Fast Red Violet Salt LB or Fast Violet B Salt. Filter and use at once. Wash blood films in running tap water for 30 to 60 seconds. Counterstain in Haematoxylin Solution filtered just prior to use, for 5 - 8 minutes. Rinse in running tap water 1 to 2 minutes. Air dry or blot carefully. Examine microscopically under an oil immersion lens. Stain Results Intensity of precipitated dye in the cytoplasm of neutrophilic leukocytes varies. One hundred are scored from 0 to 4+ based on the intensity of the precipitated dye. Scoring Criteria CELL RATING AMOUNT **% Precipitated azo dye In cytoplasm Size INTENSITY BACKGROUND OF CYTOPLASM 1+ <50 Small Faint to moderate Colourless to very pale pink 2+ 50-80 Small to Med. Moderate to strong Colourless to pale pink 3+ 80-100 Med. To Large Strong Colourless to pink 4+ 100 Med. To Large Brilliant Not visible * When there is no precipitate azo dye in the cytoplasm and the background is not stained, the cells are read as zero. ** Percentage of volume of cytoplasm occupied by azo dye precipitate. Normals Adults - Leukocyte Alkaline Phosphatase Score Of 13 To 130. References Williams, W.J., Beutler, El, Erslev, A.J. and Rundles R.W., Hematolog y, 2 nd ed., McGraw-hill , NY , C. 1977, p1628-1629. Kaplow, L.S., AM.J., Clin. Patbol ., 39:439, 1963. Kaplow, L.S. and Burstone, M.S., Nature , 200:690, 1963.
Lillie Modification of Mason's Trichrome Stain
EMS26364 Components: Phosphomolybdic/Phosphotungstic Acid Solution Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B Biebrich Scarlet Solution 1% Fast Green FCF Solution 2.5% Acetic Acid 1% Aqueous Lugol's Iodine Sodium Thiosulfate 5% Aqueous Gram's Iodine Solution Fixation: Bouin’s or Russell’s Zenker -contain no Mercury Sections: Paraffin at 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. If Zenker’s contain Mercury is used as fixative, remove Mercuric Chloride Crystal by placing in Lugol’s Iodine or Gram’s Iodine for 15 minutes. Rinse in water and clear in Sodium Thiosulfate, 5% for 3 minutes. Wash in water for 10 minutes or longer. If Russell’s Zenker is used, go to step #3. Stain in Weigert’s Iron Hematoxylin Working* solution for 5 minutes. Wash in running water. * To prepare working solution, mix equal parts of Weigert’s Iron Haematoxylin A and Weigert’s Iron Haematoxylin B. Stain in Biebrich Scarlet Solution for 4 minutes. Rinse in water Mordant in Phosphomolybdic Acid-Phosphotungstic Acid for 1 minute. Stain in Fast Green FCF Solution for 4 minutes. Differentiate in Acetic Acid, 1% for 1 minute. Carry through alcohol and dehydrate with acetone or absolute alcohol. Clear and mount in balsam saturated with salicylic acid. Stain Results: Nuclei Black Cytoplasm Brown to Pink Muscle Red Erythrocytes Brilliant Scarlet Collagen Blue to Blue-Green Myelinated Fibres Red Fibrin Red to Pink References: Clark, G.: Staining Procedures. Williams and Wilkins Co., Baltimore, 3rd ed., c. 1973, p. 59 Lillie, R.D., Arch. Path., 29:207, 1940.
Lillie's Method For Ferric and Ferrous Iron
EMS26604 Components: Potassium Ferrocyanide Potassium Ferricyanide Hydrochloric Acid 0.5% Basic Fuchsin 0.5% Acetic Acid 1% Aqueous Fixation: 10% Buffered Neutral Formalin Sections: Paraffin @ 6µm Staining Procedures: (Use control slide. Use chemically clean glassware) a) Preparation of Potassium Ferrocyanide Solution: Prepare fresh, just before use Potassium Ferrocyanide - 0.4g Hydrochloric Acid - 40.0ml b) Preparation of Potassium Ferricyanide Solution: Prepare fresh just before use. Potassium Ferricyanide - 0.4g Hydrochloric Acid - 40.0ml Deparaffinise and hydrate to distilled water. For Ferric Iron, place in Potassium Ferrocyanide Solution for 1 hour. For Ferrous Iron, place in Potassium Ferricyanide Solution for I hour. Wash in Acetic Acid, 1%. Stain for 10 minutes in Basic Fuchsin Solution. Rinse in distilled water, (For critical work, omit this step). Dehydrate in 95% alcohol, absolute alcohol and clear in xylene, two changes each. Mount with Permount. Stain Results: Ferric Iron Dark Prussian’ Blue Ferrous Iron Dark Turnbull’s Blue Background Light Red References: Lillie, R.D., Histopathologic Technic and Practical Histochemistry, 3rd ed. McGraw-Hill, N.Y., p.407, c 1965. Luna, L.G. (ed). Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology, 3rd ed., McGraw-Hill, N.Y., c. 1968, p.182.
Lillie's Method for Nucleic Acids (1965)
EMS26774 Components: Schiff's Reagent Hydrochloric Acid 1N Sodium Bisulphite Solution 0.05M Aqueous Fast Green FCF Solution 0.01% Fixation: 10% Buffered Neutral Formalin Sections: Paraffin @ 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Place slides in preheated 1N Hydrochloric Acid at 60°C for 10 -15 minutes. Stain for 10 minutes in Schiff’s Reagent. Place in Sodium Bisulphite, 0.05M for 3 changes, 2 minutes each. Wash in running water for 5 minutes. Stain for few seconds in Fast Green Solution, 0.01%. Dehydrate in 96% alcohol, absolute alcohol, and clear in xylene, two changes each. Mount with Permount. Stain Results: Nuclear Chromatin Deep Red-Purple Cytoplasm Light Green The Chromatin of plasmodia, sarcosporidia toxoplasma and histoplasma Pale Red NOTE: If the fast Green is too intense, it may be removed promptly in water, it does not wash out in alcohol. References: Lillie, R.D., Histopathologic Technic and Practical Histochemistry, 3rd ed, McGraw-Hill. NY, pp 149-150, 1965. Luna, L.G., (ed). Manual of Histologic Staining Methods of the AFIP 3rd edition. McGraw-Hill, NY, p. 131, c. 1968.
Maldonado's Method For Pancreatic Islet Cells
EMS26458 Components Phloxine 1% Aqueous Solution Phosphotungstic Acid 3% Solution Azure II Solution, 0.05% Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B Fixation Bouin's without Acetic Acid Section Paraffin, @ 6µm Staining Procedure Deparaffinise and hydrate to distilled water. Stain for 10 minutes in Phloxine Solution, 1% Rinse in distilled water. Place for 1 minute in Phosphotungstic Acid, 3%. Rinse in distilled water. Stain for 30 seconds in Azure II Solution, 0.05% Rinse in distilled water. Stain in Weigert's working solution for 1 minute. Rinse in distilled water. To prepare Weigert's Working solution: Mix just before use: 1 part Weigert's A 1 part Weigert's B DO NOT REUSE!!!!! Dehydrate in 95% alcohol, absolute alcohol and clear in xylene, two changes each. Mount with Permount Results Human Tissue Alpha granules: Purple Beta granules: Violet Blue D-cell granules: Light blue with evident granules Exocrine cells: Grayish blue with red granules References Maldonado, R. and San Jose, H. Stain Techn., 42:11-13, 1967 Luna , L.G., (ed.), Manual of Histologic Staining Methods of the AFIP, 3rd edition, McGraw, N.Y. p. 105, c. 1968
Mallory-Heidenhain Azan-Gomori’s Modification for Islet Cells
EMS26450 Components Azocarmine Stain Solution Aniline-Alcohol 1% Ferric Ammonium Sulfate 5% Aniline Blue-Orange Solution Orange G 0.2% In Alcohol Fixation Bouin's or Helly's Section Paraffin, 3-5µm Staining Deparaffinize and hydrate to distilled water as usual. Wash in running water to remove all picric acid. Stain in Azocarmine G 0.1% 45-60 Min. Perform in a covered petri dish at 56°C. Rinse and blot dry carefully. (NOTE: Azocarmine G has been restricted in use as a carcinogen and not recommended, Azocarmine B may possibly be substituted depending upon material to be stained.) Differentiate in the Aniline-Alcohol, 1% according to fixation and species used. Check periodically under a microscope until the beta cells are stained either orangy red with human tissue or red with guinea pig tissue. The beta cells should appear red against the pinkish alpha cells with colorless parenchyma (acinar). Rinse in distilled water. Transfer to Iron Alum Solution for 5 minutes or more- rinse in distilled water. Stain in Aniline Blue-Orange G for 2 to 20 minutes under microscopic control. Rinse briefly and blot carefully. Differentiate in absolute alcohol and dehydrate completely. (NOTE: if the orange colour fades too much in this step, restain the slide in Orange G, 0.2% saturated alcohol, and rewash and redehydrate with clean absolute alcohol.) Clear with xylene, two changes each. Mount. Stain Results: (Bouin Fixative) Human Tissue Guinea Pig Alpha Granules: Red, Bright Orange - Tan Beta Granules: Orange-Brown, dull Red, Fiery D-Cell Granules: Dark Blue Deep Blue References Clark, G, ed.: Staining Procedures 3rd. ed.; Baltimore: Williams and Wilkins Co., c. 1973, p171. Gomori. G: "Studies on the cells of the Pancreatic Islets." Anat Rec. Rec. 74: 439 (1939).
Marking Dyes for Tissue
EMS63020 For Mohs Chemosurgery Technique Marking Dyes for Tissue are designed to perfect orientation of excised surgical specimens. This method of permanently marking the margins of tissue specimens is very beneficial to surgeons and pathologists testing for skin cancer. This method of marking multiple margins of specimens was originally documented by Dr. Frederic Mohs. Another benefit of these dyes is the ability to process more than one specimen in a single cassette, saving the researcher both time and materials. Use this method on specimens where the chance of malignancy is very small. Usual method is to apply the dyes to fresh tissue, but they are successfully used with formalin fixed tissue. Tissues must be dried by blotting or wiping fluids from the tissue surface prior to applying the dye. Small amounts of the dyes are applied with the enclosed wooden sticks or similar applicator. Larger surfaces may require cotton swabs. Dyes will bind permanently to the tissue surface within 1 to 2 minutes when allowed to air dry. Freezing or formalin fixation will not interfere with the dye performance. Re-cap bottles after use to prevent dehydration.
Masson's Trichrome for Connective Tissue
EMS26367 Components: Bouin's Fixative Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B Biebrich Scarlet Solution 1% Phosphomolybdic/Phosphotungstic Acid Solution Aniline Blue Solution Acetic Acid 1% Aqueous Fixation: Bouin’s or 10% Buffered Formalin Sections: Paraffin at 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Mordant in Bouin’s Fixative for 1 hour at 56°C or overnight at room temperature. Cool and wash in running water until yellow colour disappears. Rinse in distilled water. Stain in Weigert’s Iron Hematoxylin Working* solution for 5 minutes. Wash in running water for 10 minutes. To prepare working solution, mix equal parts of Weigert’s Iron Haematoxylin A and Weigert’s Iron Haematoxylin B. Rinse in distilled water. Place in Biebrich Scarlet-Acid Fuchsin for 15 minutes. Rinse in distilled water. Place in Phosphomolybdic Acid-Phosphotungstic Acid for 15 minutes. Stain in Aniline Blue Solution for 10 to 20 minutes. Rinse in distilled water. Differentiate in Acetic Acid, 1% for 3 to 5 minutes. Discarded solution. Dehydrate in 95% alcohol and absolute alcohol, and clear in xylene, 2 changes each. Mount. Shorten staining times when using 2-4µm sections of renal biopsy specimens: Biebrich Scarlet – Acid Fuchsin: 5 minutes Phosphomolybdic Acid-Phosphotungstic Acid: 10-12 minutes Aniline Blue Solution: 3 minutes Stain Results: Nuclei Black Cytoplasm, Keratin, Muscle Fibres Red Collagen, Mucin Blue NOTE: To ensure proper mordanting Bouin’s Fixative should be preheated to 58 – 60°C References: Sheehan and Hrapchak. Theory and Practice of Histotechnology. St. Lous. The Mosby Comp. 1980, p. 190
Maximow's Method for Bone Marrow
EMS26252 Components: Mayer's Haematoxylin Eosin Y 0.1% Aqueous Azure II 0.1% Aqueous Phosphate Buffer 0.1M, pH 6.8-7.0 Fixation: Zenker’s / Formalin Sections: Paraffin at 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Mayer’s Haematoxylin for 15 minutes. Wash in tepid tap water for 15 minutes. Stain in working solution overnight. Prepare working solution fresh each time as follows: Eosin Y, 0.1%, Aqueous - 10ml Triple distilled water or Phosphate Buffer - 100ml Azur II, 0.1%, Aqueous - 10ml Glacial Acetic Acid - 2 drops Differentiate in 95% alcohol until blue ceases to come out into alcohol and erythrocytes and collagen are pink. Dehydrate in absolute alcohol, clear in xylene, two changes each. Mount with Permount. Stain Results: Nuclei Blue Basophile leucocyte, mast cell Purple to Violet Cartilage Purple Erythrocytes, eosinophile granules Pink Cytoplasm Blue to Pink References: AFIP, Manual of Histologic Staining Methods, 3rd edition, Ed. L. Luna: New York: McGraw Hill Publications, c. 1968, p. 120 Mallory, F.B., Pathological Technique, New York, Hafner Publishing Co., 1961, p. 196.
May-Grunwald Giemsa For Helicobactor Pylori
EMS26114 Components Jenner’s Stain Stock Solution Giemsa Stock Solution Acetic Acid 0.1% Solution Fixation 10% Buffered Neutral Formalin Sections 3µm Procedure Deparaffinise and hydrate to distilled water. Place promptly in methanol (methyl alcohol), two changes, three times each. Stain in Jenner's Working solution for 6 minutes. To prepare working solution mix: • 1 part Jenner's Stain • 1 part Distilled water Prepare fresh! Stain in Working Giemsa solution for 10 minutes. To prepare working solution mix: • 2.4ml Giemsa Stock • 50ml Distilled water Prepare fresh-do not reuse! Differentiate in Acetic Acid, 0.1% (A-114-3) one slide at a time, gently agitate, checking often under microscope until defined. DO NOT OVER DIFFERENTIATE! Dehydrate quickly in tert-Butyl Alcohol to Xylene Mount. Stain Results H-pylori Blue Background Light purple References Carson, F. Histotechnology: A Self-Instructional Text . 1980-109.
May-Grunwald/Geimsa Combined Stain For Bone Marrow
EMS26064 and 15940 Components: Lugol’s or Gram’s Iodine solution Sodium Thiosulphate 5% Methyl Alcohol May-Grunwald Solution or Jenner's Solution Giemsa Stock Solution Acetic acid 1% Fixation: Smears: Draw a curetted fragment lightly across a sterile slide in a serpentine path. Don’t allow the streak to dry completely. Start with step #4 Sections: Zenker’s or other well-fixed tissue. Start with step 1 Sections: Paraffin, 6µm Staining: Deparaffinise and hydrate to distilled water Remove mercuric chloride crystals with Lugol’s or Gram’s Iodine solution with Sodium Thiosulfate, 5% for three minutes. Wash in tap water, 10 minutes and rinse in distilled water for two changes. Place promptly in Methyl Alcohol, two changes, three times each. Stain with May-Grunwald Solution or Jenner's Solution 3 minutes. Add an equal volume of distilled water and allow standing one minute. Drain, no rinsing. Alternatively stain in working Jenner’s (dilute 1:1 with distilled water, 6 minutes. a. Sections: Stain in working Giemsa (freshly stain in working Giemsa, by diluted 2.5ml Giemsa Stock Solution (EMS15940) in 50ml distilled water, do not re-use), 45 minute b. Smears: Cover for 12 minutes with working Giemsa (15 drops of Giemsa Stock (EMS15940) to 10 drops of distilled water) a. Sections: Differentiate each slide individually in Acetic acid, 1% with gentle agitation, checking often under the microscope until nuclei are well-defined. Rinse quickly in distilled water. b. Smears: Differentiate in distilled water, agitating for approximately 5 seconds and checking under the microscope. (Section only) Dehydrated quickly in 95% alcohol and absolute alcohol and clear in two changes of xylene. a. Sections: Mount with Permount or other synthetic mount and b. Smears: Blot with fine grain filter paper and mount. Results: Blue Nuclei, Bacteria Pink – Rose Cytoplasm Tones are redder than when stained with Giemsa alone. References: Beck, R.C., Laboratory Manual of Hematological Technique, W.B. Saunders & Co., Philadelphia, c1938, pp233 - 236 Clark, G., (ed.), Staining Procedures, 3rd Edition, William & Wilkins, Baltimore, pp131 & 132, c 1973. Strumia, M.M., J. Lab. Clin. Med., 21:930-934, 1935 – 1936. Luma, L.G., (ed.) Manual of Histologic Staining Method of the AFIP, 3rd edition, McGraw-Hill, NY, pp 121 & 122, c1968.
Mayer's Mucicarmine Method
EMS26320 Components Mucicarmine Stock Solution Metanil Yellow 0.25% Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B Fixation 10% Buffered Neutral Formalin Section Paraffin @ 6µm Staining Procedure Deparaffinise and hydrate to distilled water. Stain in working Weigert's Haematoxylin [mix 1 volume of Weigert's A, plus 1 volume of Weigert's B,] for approximately seven minutes. Wash in running water for 10 minutes. ***If using a microwave oven - place slides in working Weigert's Haematoxylin solution and heat in the microwave oven for 30 seconds and immediately transfer to tap water. Stain in the working Mucicarmine (mix 1 volume of Mucicarmine Stock, plus 4 volumes of tap water) for 60 minutes of longer. Prepare working solution fresh, use once and discard. ***If using a microwave oven - place slides in Mucicarmine Working Solution and heat in the microwave oven for 60 seconds. The hot stain jar is removed from the oven and allowed to remain at room temperature (22°C) for 5 minutes. Rinse quickly in distilled water for 2 or 3 changes. Counterstain in Metanil Yellow, 0.25%, for 15 seconds to 1 minute or longer. (Older solutions require longer times) Rinse quickly in distilled water. (2-4 seconds) Dehydrate in 95% and 100% alcohol's, two changes; clear through two changes of Xylene. Mount with Permount or other synthetic medium. Stain Results Mucin, Cryptococci Capsulae deep rose to red Other Tissue Elements yellow Nuclei black Note: The rose color due to carmin staining will be obscured if sections are overstained with Weigert's Hematoxylin and/or Metanil Yellow solution. References Mallory, F.B.: Pathological Technique, Hafner Publishing Co., N.Y., c. 1961, p. 130 Luna, L.G., (ed), AFIP Manual of Histologic Staining Methods, 3 rd . ed., McGraw-Hill Co. , NY, c. 1968, pp 161-162. Sheehan , D.C. & Hrapenak, B.B., Theory and Practice of Histotechnology, 2 nd ed., C.V. Mosby Co., St. Louis, c. 1980, pp 168-169.
McManus Pas Method for Glycogen
EMS26312 Components: Coleman's Feulgen Light Green Stock Solution 0.2% Periodic Acid 0.5% Aqueous Acid-Alcohol 1% Ammonia Water 0.3% Schiff'S Reagent Light Green Working Solution Harris Haematoxylin Fixation: 10% Buffered Neutral Formalin or Zenker’s Sections: Cut Paraffin, 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Oxidise in Periodic Acid, 0.5%, 5 minutes and rinse in distilled water. Stain in Coleman’s Feulgen or Schiff’s Reagent, 15 minutes and wash in running water to develop the pink colour, 10 minutes. Counterstain in Harris Hematoxylin 6 minutes, or Light Green Working Solution 5 – 10 seconds. Light Green is better used when delineation of fungi is required. Prepare the working solution by diluting Light Green, 0.2% Solution 1:5 ratio with distilled water; proceed to step #7, dehydration. Tap water and ammonia decolourise Light Green. Wash in running water and transfer to Acid Alcohol, 1%, for 3 -10 quick dips. Wash again in distilled water. Dip in dilute Ammonia Water, 0.3%, to blue the sections and again wash in running water for ten minutes. Dehydrate in 95% alcohol, absolute alcohol, and clear in xylene, two changes each. Mount with Permount or DPX. Stain Results: Nuclei Blue Fungi Red Background, when Light Green is used as the counterstain Green, pale References: McManus, JFA: Stain Tech.; 23:99 (1948). AFIP, Manual of Histologic Staining Techniques; L.G. Luna, 3rd edition, New York: McGraw-Hill Publication, c. 1968, p. 160. Mowry, R.W.: Annals of the New York Academy of Science; 106: 402 (1963).
Methenamin Silver Method for Argentaffin Cells
EMS26481 Components: Weigert's Iodine Solution Sodium Thiosulfate 5% Aqueous Silver Nitrate 5% Aqueous Methenamine 3% Aqueous Borate Buffer pH 7.8 Gold Chloride 0.1% Aqueous Safranin O Solution 0.1% Fixation: Formalin, Bouin’s or other Formalin containing fixatives. Dichromates and mercury salts impair contrast between granules and background. Sections: Cut paraffin at 3-5µm sections. Staining Procedures: Deparaffinise and hydrate slides to distilled water. Treat for 10 minutes in Weigert’s Iodine followed by bleaching for 10 minutes in Sodium Thiosulfate, 5%. Wash for 10 minutes in running water. Rinse in two changes in distilled water. Place section in chemically clean coplin jars containing the buffered methenamine-silver nitrate solution at room temperature and place in 60°C paraffin oven for 3 – 3.5 hours. (preheating the solution to 60°C reduces impregnation time by ½ to 1 hour). Rinse in distilled water. * prepare methenamine-silver nitrate by mixing: Silver Nitrate, 5% - 5ml Methenamine, 3% - 100ml For working buffered stain add 8 ml of Borate Buffer, pH 7.8 to 30ml of methenamine-silver nitrate. Use chemically clean glassware throughout. Tone for 10 minutes in Gold Chloride, 0.1%. Rinse in distilled water. Fix in Sodium Thiosulfate, 5% for 2 minutes. Wash in running water for 5 minutes Counterstain in Acetic Safranin O, 0.1% for 5 minutes. Dehydrate with Acetone, clear in Xylene and mount. Stain Results: Argentaffin Cells Black Coarse connective tissue of the submucosa Variable amount of blackening by 3-3.5 hour of impregnation Granules of eosinophil leukocytes, nuclei, smooth muscle & surface epithelium Show additional blackening after incubation beyond 3-3.5 hour. Granules of mast cells Remain red after nuclei & reticulum are blackened References: Clark, G, ed.: Staining Procedures, 3rd ed.: Baltimore: Williams & Wilkins Co., 1973, p. 161 Gomori, G.: Arch Path., 45:48, 1948 Burtner, H.J. and Lillie, R.D., Stain Tech., 24:225-7, 1949.
Microwave PAS-Alcian Blue Method For Mucosubstance
Components: Alcian Blue Solution Periodic Acid, 1% Aqueous Schiff's Reagent Gill's Haematoxylin Ammonia Water, 1% Fixation: 10% buffered Neutral Formalin Section: Paraffin, @ 3-5µm Staining Procedure: Deparafinised and hydrate in the usual manner. Place slides in a loosely covered plastic Coplin jar containing 60ml Alcian Blue Solution Microwave on medium level 60 seconds. Tightly cover, agitate, and let stand 5 minutes. Wash in distilled water. Place slides in a loosely covered plastic Coplin jar containing 60ml Periodic Acid, 1% Aqueous. Microwave on medium level 60 seconds, tighten cover, agitate, and let stand one minute. Wash in distilled water. Place slides in a loosely covered plastic Coplin jar containing 60ml Schiff's Reagent Microwave on medium level 60 seconds, tighten cover, agitate, let stand 30 seconds. Wash in warm running water for three minutes. Place slides in a loosely covered Coplin jar containing 60ml Gill's Haematoxylin #2 Microwave on medium level for 30 seconds. Immediately rinse slides in distilled water. Dip once in Ammonia Water, 1% to blue haematoxylin Wash slides in distilled water, dehydrate in 70%, 95% and two changes of absolute alcohol. Clear in two changes Xylene and mount with Permount Result: Acid Mucins turquoise Neutral Mucins magenta Combined acid and neutral mucins purple Nuclei blue References: Mathews Karen and James K. Kelly. Dept. of Histology, Foothills Hospital. Calgary , Alberta. Canada . T2N 2T9. Journal of Histotechnology Vol. 12. No 4 Dec. 1989.
Modification of Mayer's Mucihematein
EMS26321 Fixation: Absolute Alcohol 4 – 8 hours. Sections: Paraffin or Celloidin Staining Procedures: Deparaffinise and hydrate to distilled water. Place on staining rack and rinse several times with distilled water. Place 2 ml of Mayer Mucihematein, Modified on each slide. Allow to remain for five minutes. Drain off stain and wash well with distilled water three times for 5 minutes each. Dehydrate in two changes of 95% alcohol, two changes of absolute alcohol, Absolute Alcohol-Xylene (50:50), and two changes of Xylene. Mount with Permount or DPX. Stain Results: Mucin Deep Violet Cell Nuclei Pale Grey-Blue Connective Tissue Pale Grey to Colourless References: Laskey, Alice: Stain Techn., 25:33 (1950). Clark G.: Staining Procedures, William & Wilkins Company, Baltimore, 3rd Ed., c. 1973, p. 47
Modified Alizarin Red S for Foetal Specimens
EMS26204 Components: Alizarin S 0.01% Aqueous Solution Potassium Hydroxide 1% Aqueous Solution Potassium Hydroxide 10% Aqueous Solution Glycerol Potassium Hydroxide Fixation: Two or more weeks in 95% alcohol or Formalin after evisceration. (evisceration omitted for small embryos). Staining Procedure: Rinse in water and place in Potassium Hydroxide, 1% for a few hours to a week or more (Formalin fixed specimens require approximately 1 month in Potassium Hydroxide, 10% until bones are clearly viable through the soft tissue. Note: If the tissue becomes too soft, harden for 12-24 hours in solution containing equal parts of 95% alcohol, water and glycerin and then return to the Potassium Hydroxide 1%. The Potassium Hydroxide, 1% may be diluted to 0.5% with distilled water during the last few days of clearing. Wash in running water for 12 hours. Stain in Working Alizarin Red S Solution* for 30-60 minutes. Wash in running water for 30 minutes. (Prepare Working Stain by adding 6-10 drops of Potassium Hydroxide, 1% to 100ml of Alizarin Red S, 0.01%. Prepare fresh.) Decolourise in Glycerol-Potassium Hydroxide Solution (C-204-3) for 1-2 weeks. Mount on a glass frame and dehydrate by passing slowly through the following alcohol-glycerin-water mixtures, in the order given: 1:2:7, 2:2:6, 3:3:4, 4:4:2 and finally 5:5 without water. Seal in the final glycerin-alcohol mixture. Stain Results: Bones - Red Soft Tissue - Transparent and unstained References: Dawson, A.B., Stain Techn., 1:123-4, 1926 Richmond, G.W. and Bennett, L., Stain Techn., 13:77-79, 1938. Clark, G., (ed.), Staining Procedures, 3rd edition, Williams & Wilkins, Baltimore, p.137, c. 1973.
Modified Movat's Stain
EMS26385 Components Alcian Blue, 1% Alkaline Alcohol Orcein, 0.2% Haematoxylin Alcoholic, 5% Ferric Chloride, 10% Lugol’s Iodine Woodstain Scarlet-Acid Fuchsin Working Solution Acetic Acid, 0.5% Phosphotungstic Acid, 5% Alcoholic Saffron Fixation 10% Buffered Neutral Formalin or Bouin's Solution Note: If formalin fixative is used, decerated tissue sections must be mordanted in Bouin's Fluid for 1 hour in a 50°C oven. Wash well in running water to remove picric acid deposits. Sections 4-6µm Staining Procedure Decerate section and hydrate to distilled water. Stain in Alcian Blue, 1% for 20 minutes. Dip five times in distilled water. Place in Alkaline Alcohol in a 56°C oven for 10 minutes. Wash in running tap water for 2 minutes. Stain in Orcein-Verhoeff working solution for 2 hours. To prepare Orcein-Verhoeff working solution: Immediately before use, in the following order, combine the solutions below: Orcein 0.2% 25.0ml Alcoholic Haematoxylin 5% 8.0ml Ferric Chloride 10% 5.0ml Lugol's Iodine 5.0ml Wash in running tap water for 3 minutes. Stain in Woodstain Scarlet- Acid Fuchsin working solution for 2 1/2 minutes. Place in Acetic Acid, 0.5% for 30 seconds. Differentiate in Phosphotungstic Acid, 5% for 5-10 minutes, well differentiated sections demonstrate colourless collagen and blue-green mucopolysaccharides. Rinse in Acetic Acid, 0.5% for 30 seconds. Three changes of 100% ethyl alcohol, 1 minute each, Stain in Alcoholic Saffron for 8 minutes. Dehydrate in 100% ethyl alcohol, two changes. Clear in xylene, three changes. Coverslip. Stain Results Nuclei Black Cytoplasm Red Elastic Fibres Purple to Black Collagen and Bone Yellow Mucopolysaccharides Blue-Green Muscle Red References Movat, H.Z.,: Demonstration of all Connective Tissue Elements in a Single Section. Arch Path., 60:289-295, 1955. Roman N., et al: Orcein-Hematoxylin in Iodized Ferric Chloride as a Stain for Elastic Tissue. Stain Techn., 42:199-202, 1967. Silverman Jefferey., A Modified Movat Pentachrome Stain. Histologic , p. 33. 1972
Modified Steiner Using Chapman's Modification
EMS26119 Components Zinc Formalin Silver Nitrate, 1% Aqueous Gum Mastic, 2.5% Hydroquinone, 2% Silver Nitrate, 0.04%, Aqueous Fixation 10% Buffered Neutral Formalin Section 4-6µm Staining Procedure Deparaffinise and re-hydrate tissues to distilled water. Sensitize sections in Zinc Formalin at 60°C in a water bath for 15 minutes (cover jar loosely). Preheat this solution. Rinse in distilled water, 3 times for 2 minutes each. Place in Silver Nitrate, 1%, in a Coplin Jar. Place in microwave (800W); start and run for 80 seconds. Remove from microwave and allow to stand in hot silver solution for 20 minutes (cover jar loosely). Rinse in distilled water 3 times for 2 minutes each. Dehydrate in 2 changes of 95% ethanol followed by 2 changes of 100% ethanol. Treat with Gum Mastic, 2.5%, for 5 minutes. Allow to air dry for 1 minute. Rinse in distilled water (2 changes). Slides may stand in distilled water while reducing solution is being made, or longer if necessary. To prepare reducing solution, mix just before use: a. Gum Mastic, 2.5% 10.0ml b. Hydroquinone, 2% 25.0ml c. Ethanol, 100% 5.0ml Filter and then add: d. Silver Nitrate, 0.04% 2.5ml DO NOT filter this silver! Reducing solution at 45°C in water bath for 5-30 minutes or until sections have developed satisfactorily with the spirochetes at desired intensity and the background light yellow. Avoid an intensely stained background by checking microscopically. DO NOT PREHEAT THIS SOLUTION. Rinse in distilled water to stop reaction. Dehydrate through graded alcohol's clear in, Xylene (not Xylene substitute) and mount with Permount. Results Spirochetes, Donovan Bodies, General Bacteria Legionnaries' Disease Bacteria dark brown or black Background bright yellow to light brown Note: Acid wash all glassware used with silver solutions! References Margeson, L.S. Chapman CM: Use of Zinc Formalin as a sensitizer in Silver Stains for Spirochetes. Journal of Histotechnology , 19:2 p. 135-138, 6-96.
Modified Verhoeff Elastic-Van Gieson Stain
EMS26369 Components Alcoholic Haematoxylin Ferric Chloride 2% aqueous Lugol’s Iodine Ferric Chloride 0.4% aqueous Van Geisons Solution Consistent and shorter staining time! Fixation 10% Buffered Neutral Formalin Sections 4-5µm Staining Deparaffinise and hydrate sections. Stain in modified Verhoeff at room temperature for 7 minutes or in a 60°C oven for 3.5 minutes. To prepare modified Verhoeff Working Elastic solution, mix just before use: • 3 parts Alcoholic Haematoxylin, 3% • 2 parts Ferric Chloride 2%, aqueous • 1 part Lugol's Iodine Wash in running warm tap water for 1 minute. Differentiate in Ferric Chloride, 0.4% aq. for 75 seconds. Wash in running warm tap water for 5 minutes. Counterstain in Van Geisons solution for 60 seconds. Dehydrate, clear and mount with Permount. Results Nuclei Greyish-black Elastic Fibres Bluish-black Collagen fibres Red Muscle Greenish yellow References Garvey, Winsome et al "A modified Verhoff Elastic Van Geison Stain" The Journal of Histotechnology, Vol. 14, No. 2 pp113-114 June 1991.
Neat Stain Gram Stain Kit
EMS27100-01 Summary For staining bacteria from cultures or patient specimens by the differential Gram stain method. This procedure is based upon the Hucker modification of the original Gram stain method. Gram-positive and Gram-negative bacteria are both stained by the crystal violet stain. Addition of iodine leads to the formation of a crystal violet-iodine complex within the cell wall. The decolouriser extracts lipids from the cell wall of Gram-negative bacteria, thereby increasing the porosity of the cell wall and allowing the crystal violet-iodine complex to diffuse from the cell. At the same time, Gram-positive bacteria are dehydrated, causing a decrease in cell wall porosity and trapping the crystal violet-iodine complex within the cell. Due to the increase in porosity, the safranin counterstain is able to permeate the cell wall of the Gram-negative bacteria. Storage Conditions Store at controlled room temperature (15-30°C) with bottles tightly sealed. Neat Stain Gram stain reagent packs may be resealed with adhesive backed foil strips between uses. Do not use beyond the expiration date on the reagent pack. Indications of Deterioration If precipitate forms in the Crystal Violet Solution, warm gently to 37°C and shake well to re-dissolve. Specimen Collection & Preparation Liquid Media: Spread the bacteria suspension on a clean glass slide using a sterilised wire loop and allow to air-dry. Heat-fix the smears by passing the slide through a flame three times and allow them to cool. Solid Media: Place a loopful of deionized water on a clean glass slide. Touch a sterilised wire loop to a colony and suspend the cells in the water droplet; spread the suspension on the slide and allow to air dry. Heat-fix the smear by passing the slide through a flame three times and allow it to cool. Patient: Obtain the patient specimen using a sterile wire loop, pipette or swab. Smear the specimen on a clean glass slide and allow to air dry. Heat-fix the smear by passing the slide through a flame three times and allow it to cool. Technique Immerse the prepared slide into chamber 1 (Crystal Violet Stain) for one minute. Rinse the slide gently with DI water. Immerse the slide into chamber 2 (Stabilized Iodine Solution) for approximately two minutes. Rinse the slide gently with DI water. Repeatedly dip the slide into chamber 3 (Decolorizer) inspecting between each dip. Decolorizing is complete as soon as the solution runs clear from the smear on the slide. Rinse the slide gently with DI water. Immerse the slide into chamber 4 (Safranin) for one minute. Rinse the slide gently with DI water and blot the slide dry with absorbent paper. Examine the slide accordingly. Results Gram-positive bacteria are stained purple and Gram-negative are stained pink to red. Limitations of Procedure Application of excessive heat during fixation of smear may rupture bacterial cell walls and affect the Gram staining results. Treatment with antibacterial drugs may cause Gram-positive bacteria to appear Gram-negative. Controls: Performance of the Set may be tested using the following procedure: Stain a prepared slide of Staphylococcus aureus from an 18 to 24 hour culture with Crystal Violet Stain allowing to stand for one minute and rinsing with DI water. Gram positive purple cocci should be observed; if not, discard this solution. Stain a prepared slide of Escherichia coli from an 18 to 24 hour culture with Safranin Stain allowing to stand for one minute and rising with DI water. Gram- negative pink rods should be observed; if not, discard this solution. Following the normal Gram staining procedure, stain a prepared slide of S. aureus from an 18 to 24 hour culture. Gram-positive purple cocci should be observed; if not, discard the Gram Iodine Solution. Precautions Crystal Violet: Causes eye irritation. Wash thoroughly after handling. In case of contact, flush eyes with water for 15 minutes. Contact a physician. Iodine: Causes eye irritation. Wash thoroughly after handling. In case of contact, flush eyes with water for 15 minutes. Contact a physician. Decolorizer: Is extremely flammable. Use with adequate ventilation. Avoid excessive heat, sparks and open flame. Decolorizer causes eye irritation. Wash thoroughly after handling. In case of contact, flush your eyes with water for 15 minutes. Contact a physician. Safranin: Is a combustible liquid. Avoid excessive heat, sparks and open flame. Causes eye irritation. Wash thoroughly after handling. In case of contact, flush eyes with water for 15 minutes. Contact a physician. Limited Warranty This product is warranted to perform as described in its labeling and product literature. Electron Microscopy Sciences, Inc. disclaims all other warranties, expressed, implied and statutory with respect to this product, including, but not limited to the warranty of fitness for a particular purpose and the warranty of merchantability. In no event shall Electron Microscopy Sciences, Inc. or any of its affiliates be liable for incidental or consequential damages to the buyer or any third party, howsoever arising, even if advised of such damages.
Neat Stain Haematology Stain Kit
EMS27104-01 This kit is designed for the differentiation of morphological cell types in peripheral blood smears. The staining characteristics are similar to the traditional Wright’s and Wright-Giemsa stains. The simple three-step procedure is useful for laboratories performing manual staining of microorganisms and spermatozoa. The kit is also well suited for stat procedures. Each kit contains the following components: 4 oz. bottle of QUICK III Fixative Solution 4 oz. bottle of QUICK III Solution I 4 oz. bottle of QUICK III Solution II The Neat Stain Haematology Stain Kit contains exceptional stain quality with each reagent in a convenient, wide-mouth reagent bottle with a screw cap lid. Storage Conditions Store at controlled room temperature (15-30°C). Keep cuvettes tightly sealed when not in use. Do not use after specified expiration date. Indications of Deterioration Mould growth or excessive precipitation in reagents are indications of contamination and or degradation. Specimen Collection & Preparation Smears prepared from fresh uncoagulated venous or arterial blood should be air-dried. If the smears are not prepared immediately, 2mg of EDTA/ml of blood sample should be added to prevent coagulation. Technique Dip the prepared slide into the QUICK III Fixative Solution for 1 second and then remove, repeat for a total of 5 dips total. Ensure that the sample area on the slide is submerged in solution while dipping. Drain or blot the slide carefully to remove excess reagent. (Slides can be dipped directly into the reagent bottle for convenience.) Repeat the procedure in Step 1, substituting the QUICK III Solution I. Repeat the procedure in Step 1, substituting the QUICK III Solution II. Rinse slides with deionised water and allow to dry. Note: Staining intensity may be varied by increasing or decreasing the number of dips in Solution I or Solution II. For a more eosinophilic result (red intensity), increase the number of dips into Solution I; for a more basophilic result (blue intensity), increase the number of dips into Solution II. Ideally, five one second dips into each Solution will produce the desired results. Any variation of the procedure other than described above may adversely affect the results. Stained Specimen Results Specimen Nucleus Granules Cytoplasum Erythrocytes n/a n/a Pink Leukocytes Granular Polymorphonuclear Neutrophils Purple Red-lilac Light pink Eosinophils Dark blue Red/orange Medium blue Basophils Dark blue Dark purple Light blue Leucocytes Nongranular Monocytes Violet n/a Light blue Lymphocytes Violet n/a Medium blue Platelets n/a Purple Lilac Microorganisms Blue/violet n/a n/a Spermatozoa Dark blue Danger Poison may be fatal or cause blindness if swallowed. Vapour is harmful and will cause eye irritation. Do not breathe vapour. Avoid contact with eyes, skin or clothing. Keep container closed. Use with adequate ventilation. Keep from heat, sparks and open flame. Wash thoroughly after handling. CANNOT BE MADE NONPOISONOUS. First Aid: If swallowed, call a physician immediately. In case of contact with eyes, immediately flush with water for 15 minutes and notify a physician. Wash exposed areas thoroughly after handling.
New Methylene Blue Solution
EMS26077 Introduction New Methylene Blue N Stain, Aqueous, provides a staining technique for reticulum of immature erythrocytes. Immature erythrocytes (reticulocytes) contain RNA which is lost as cells age. When New Methylene Blue N Stain, Aqueous, is used to supravitally stain viable erythrocytes, the RNA in young cells is precipitated and stained deep blue. The stained RNA appear as blue granules which may be connected into a reticulo-filamentous pattern. Staining Procedure Filter New Methylene Blue N Stain, Aqueous, prior to use. Collect appropriate blood sample for reticulocyte count, per laboratory protocol. Proceed with staining procedure as soon as possible after blood draw. Pre-label microscopic slide(s) with appropriate patient identifiers. Mix five drops of New Methylene Blue N Stain, Aqueous with five drops of whole blood; mix gently with a pipette. See Procedure Notes #1, #2, and #3. Incubate mixture at room temperature for 10-15 minutes. See Procedure Note #4. Thoroughly remix stain/blood suspension after incubation. See Procedure Note #5. Prepare wedge smear(s) on pre-labeled microscopic slide(s) with the remixed stain/blood suspension. Allow slide(s) to thoroughly air-dry. Evaluate reticulocyte count under oil immersion. Results Reticulocytes Pale blue with dark blue granular/reticular material Red cells Pale blue or blue-green Procedure Notes A small test tube, vial, or centrifuge tube can be used for mixing purposes. Smaller or larger amounts of stain and blood can be mixed as long as volumes are of equal proportions. Separate pipettes should be used for each solution and step to avoid any possibility of sample contamination. Incubating longer than 15 minutes may increase the possibility that mature erythrocytes will also be darkly stained. Reticulocytes have lower density than mature erythrocytes and will be near the top during incubation. Remixing prior to preparing smears allows for equal cell distribution. References Bauer, John D. Clinical Laboratory Methods. 9th ed. St. Louis: Mosby, 1982. 195-198. Lillie, R.D., and Harold Fullmer. Histopathologic Technic and Practical Histochemistry. 4th ed. New York: McGraw-Hill, 1976. 752-753. McPherson, Richard and Matthew Pincus. Henry's Clinical Diagnosis and Management by Laboratory Methods. 22nd ed. Philadelphia: Elsevier Saunders, 2011. 514. 544.
Nile Blue A Stain
EMS26511 Components Nile Blue A Staining Solution Sulphuric Acid 1% Aqueous Fixation Formalin fixatives Application Show differentiation of melanins and lipofuscins Staining Deparaffiniz=se and hydrate to distilled water. Stain 20 minutes in Nile Blue A Staining solution. Rinse quickly with sulphuric acid, 1%. Dehydrate immediately in four changes of acetone, 15 seconds each. Rinse in Xylene-acetone. Clear in two changes of Xylene. Mount. Stain Results Lipofuscins Unstained - natural yellow Melanins Green References Staining Procedures, G. Clark: Williams and Wilkins Co., Baltimore, 3 rd ed., c. 1973, p. 122. Lillie, R.D.: Stain Techn ., 31:151.
Oil Red in Propylene Glycol Method
EMS26504 Components: Oil Red O Solution Mayer's Haematoxylin Harris' Haematoxylin Propylene Glycol 85% Aqueous Solution Hydrochloric Acid 5% Ammonia Water 0.3% Fixation: 10% Buffered Neutral Formalin Section: Frozen Staining Procedure: Cut frozen section and collect in distilled water. Propylene Glycol, Absolute, for 2 minutes Oil Red O Solution, for one hour. If sections are mounted on glass slides before staining, allow staining process to continue overnight. Differentiate in Propylene Glycol, 85%, for one minute. Rinse in two changes of distilled water. Stain for a few seconds in Mayer’s Haematoxylin, or Harris’s Haematoxylin. Rinse in two changes of distilled water. If over-stained differentiate in Hydrochloric Acid, 5%, wash in water; neutralise in Ammonia Water, 0.3%. Wash in two changes of distilled water. Mount with Glycerin Jelly. Results: Fat = Red Nuclei = Blue References: AFIP Manual of Histologic Staining Methods, Ed. L. Luna, 3rd ed., New York, McGraw-hill Publications, c. 1968, p. 140.
Oil Red O Method For Lipofuchsin
EMS26609 Components Oil Red O Solution 0.5% Propylene Glycol 85% Aqueous Mayer’s Haematoxylin Fixation 10% Buffered Neutral Formalin Section Paraffin at 6µm Staining Procedure Deparaffinise and hydrate to distilled water. Stain in Oil Red O Solution for 72 hours. Agitate the slide in Propylene Glycol, 85% for 1 minute. Two changes of distilled water, 2 minutes each Stain in Mayer's Haematoxylin for 5 minutes. Wash in running water for 10 minutes. Mount with Glycerin Jelly Staining Results Lipofuchsin Red Nuclei Blue References Luna, L.G. (ed.), Manual of Histologic Staining Methods of the AFIP , 3 rd edition, The AFIP, 3 rd edition, McGraw-Hill, NY, P. 186, c 1968.
Papanicolaou Stain for Membrane Filters
EMS26801 Components: Gill’S Haematoxylin No 1 Hydrochloric Acid 0.05% Solution Scott’s Water Solution Modified Orange G Solution Modified EA Staining Solution Fixation: 95% Ethyl Alcohol Staining Procedure: Fixation in 95% ethyl alcohol for 15 minutes. Two changes of distilled water, 20 dips each. Stain in Gills haematoxylin No. 1 for 2-3 minutes. Wash in two changes of distilled water until the excess stain is removed from the filter. Place in Hydrochloric Acid 0.05% for 45 seconds or until the filter turns yellow. Wash in two changes of distilled water, 20 dips each. Scott’s Tap Water Substitute for 1 minute. Wash in two changes of distilled water, 20 dips each. Two changes of 95% ethyl alcohol, 10 dips each. Counterstain in Modified OG-6 Staining Solution for 2 to 3 minutes. Two changes 95% ethyl alcohol, 1 minute each Counterstain further in Modified EA Staining Solution for 2 to 3 minutes. Dehydrate in 2 changes each of 95% ethyl alcohol and absolute ethyl alcohol. Clear in 2 changes of Xylene. Soak filter in Permount for 15 minutes and mount in Permount. Results: Nuclei Deep Blue to Purple References: Gill. G.W., Acta Cytol., 18:300-311, 1974 Diagnostic Cytology by Membrane Filter, Application Bulletin 100, Gelman Instrument Co., Ann Arbor, MI, 1978.
PAS-Alcian Blue Method For Mucosubstances
Components: Alcian Blue Solution Periodic Acid 1% Schiff's Reagent Sodium Metabisulphite 0.5% aqueous Fixation: 10% Buffered Neutral Formalin Section: Paraffin, @ 6µm Staining Procedure: Deparaffinise and hydrate to distilled water. Stain in Alcian Blue Solution, pH 1.0 or Alcian Blue, pH 2.5 for 30 minutes. If Alcian Blue solution, pH 1.0 is used, blot section dry with filter paper. If Alcian Blue solution pH 2.5 is used wash, in water for 5 minutes. Oxidize in Periodic Acid 1% for 30 minutes. Wash in running water for 5 minutes. Schiff's Reagent, for 10 minutes. Rinse in Sodium Metabisulfite, 0.5% aqueous, three changes, two minutes each. Wash in running water for ten minutes. Dehydrate in 95% Alcohol, Absolute Alcohol and clear in Xylene, two changes each. Mount with Permount. Results: PAS-Alcian Blue pH 2.5 - All polysaccharides and mucosubstances containing hexoses or deoxyhexoses with vicimal glycol groups stain magenta to red. Mucosubstances staining red include neutral mucosubstances. Hyaluronic acid, sialomucins and all but the most strongly acidic sulphated mucosubstances stain blue. PAS-Alcian Blue pH 1.0 - PAS positive same as above. Alcinophilic substances at this pH include only the sulphated mucocaccharides. References: AFIP Manual of Histological Staining Methods, 3 rd ed. Ed. L. Luna: New York : McGraw Hill Publications c. 1968, p. 168. Lev. R. & Spicer S.S. J. Histochem. Cytochem., 12:309. 1964.
Pearse's Method for Phospholipids
EMS26516 Components: Luxol Fast Blue MBS Solution 0.1% Lithium Carbonate 0.05% Aqueous Neutral Red Solution 1% Aqueous Fixation: Bakers Calcium Cadmium Formal. Sections: Paraffin @ 6µm Staining Procedures: Deparaffinise and place in absolute alcohol. Stain in Luxol Fast Blue Solution overnight at 58°C. Rinse in 70% Alcohol and then rinse in distilled water. Differentiate for 30 minutes to 2 hours in Lithium Carbonate 0.05%. Rinse in distilled water. Counterstain for 30 minutes in Neutral Red, 1%. Rinse in distilled water and blot dry. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene, two changes, two minutes each change. Mount with Permount Stain Results: Phospholipids Blue Nuclei and Nucleoli Red Early Lipofuchin, Eosinophil, Granules, Keratin, Keratohyalin, Human Elastic Tissues Blur to Purple References: Pearse, A.G.E., J. Path. Bact., 70:554-557, 1995. Luna, L.G. (ed). Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology, 3rd ed., McGraw-Hill, N.Y., c. 1968, p.148
Periodic Acid Leucofushin Method (PAS)
EMS26853 Components: Schiff’s Reagent Periodic Acid 1%, Aqueous Sodium Bisulphite 0.05M Aqueous Mayer’s Acid Hemalum Solution Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B Picric Acid, Saturated Aqueous Orange G Solution, 1% Fixation: Zenker Fluid or 10% Formalin Sections: Paraffin @ 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Rinse in tap water. Oxidize in Periodic Acid 1% Aqueous for ten minutes. Wash five minutes in running water. Stain ten minutes with Schiff’s Reagent Pass directly to three successive baths of two minutes each in Sodium Bisulfite 0.05M. Wash in running tap water for ten minutes. Stain in Weigert’s Hematoxylin or Mayer’s Acid Hemalum for two to five minutes, wash in tap water. To prepare Weigert’s Hematoxylin: Mix equal parts of Weigert’s A and Weigert’s B just before use. Counterstain for one minute woth Picric Acid, Saturated (Aq) or Orange G, 1%. (A gram-Weigert staining procedure may be substituted for steps 6-8). Dehydrate in two changes each of 95% and absolute alcohol. Clear through Alcohol-Xylene and two changes of Xylene Mount in Clarita. Stain Results: Nuclei Black on Blue Collagen Pink (Orange if Picric Acid is used as counterstain) Reticulm Purplish Red (Orange-red if Picric Acid is used) Glycogen- Dark Purplish Red Epithelial Mucin Red-Purple to Violet Filbrin Pink (Pink to Violet if the Weigert Fibrin Variant was used as a counterstain) Cytoplasm Gray, yellow or Orange (depending on the counterstain) References: Clark, G: Staining Procedures, Williams & Wilkins. Co., Baltimore, 3rd Ed. C.1973, P. 156.
Picrosirius Red Staining Of Cardiac Muscle Following Phosphomolybdic Acid Treatment
EMS26357 This method eliminates cytoplasmic staining, revealing thin septa and collegen fibres clearly. Components Phosphomolybdic Acid, 0.2% Aqueous Sirius Red, 0.1% In Saturated Picric Acid Hydrochloric Acid, 0.01N Fixation Bouins' Fixative Sections Parrafin @ 7 microns. Staining Deparaffinise and hydrate to distilled water. Wash in running tap water, 10 minutes. Rinse in distilled water. Treat in Phosphomolybdic Acid 0.2% Aqueous (1-5 minutes) Stain in Sirius Red, 0.1% in Sat'd Picric Acid , for 90 minutes. Wash for 2 minutes in 0.01 N Hydrochloric Acid. Rinse in 70% Alcohol for 45 seconds. Dehydrate, clear and mount in Permount Results Septa Red References Dolber, Paul C. and Madison S. Spach. Picrosirus Red Staining of Cardiac Muscle Following Phosphomolybdic Acid Treatment , Stain Technology , Vol.62 No.1 (1987) 23-26.
Pinkerton's Method for Rickettsia
EMS26131 Components: Methylene Blue Solution 1% Basic Fuchsin 0.25% Aqueous Citric Acid 0.5% Aqueous Solution Gram's Iodine Solution Lugol's Iodine Sodium Thiosulfate 5% Aqueous Fixation: Zenker’s, Buffered Neutral Formalin, 10% or Moller’s. Sections: Paraffin at 6µm sections Staining: Deparaffinise and hydrate to distilled water. If Zenker fixed, remove “Zenker’s crystals” by placing in Gram’s Iodine or Lugol’s Iodine for 15 minutes. Rinse in water and clear in Sodium Thiosulfate, 5% for three minutes. Wash in water for 10 minutes or longer. Stain overnight in Methylene Blue Solution. Rinse in 95% alcohol for 5 seconds to prevent loss of blue colour. Rinse quickly in distilled water for 2-3 seconds. Stain for 30 minutes in Basic Fuchsin, 0.25%. Decolourise rapidly in Citric Acid, 0.5% for 1-2 seconds, never more than 3 seconds. Differentiate further in absolute alcohol until nuclei stand out blue and rickettsia clumps red. Clear with 2 changes of Xylene. Mount with Permount or SHUR/Mount. Stain Results: Rickettsia Bright Red Nuclei Blue References: Simons, J.S. and Gentzkow, C.J., - Laboratory Methods of the United State Army, 5th ed,. Lee & Febiger, Philadelphia, p. 572, 1944. Luna, L.G., (ed) Manual of Histologic Staining Methods of the AFIP, 3rd edition, McGraw-Hill, New York, p. 237, c. 1968.
Pizzolato's Method For Calcium Oxalate
EMS26212 Fixation Buffered Neutral Formalin, 10% Sections Paraffin @ 6µm Staining Deparaffinise and hydrate to distilled water. Pour enough working silver nitrate-hydrogen peroxide solution* onto slides to cover sections. Place a 60W light bulb 6 inches from sections for 30 minutes. Rinse thoroughly with distilled water. Prepare working solution by mixing, just before use, equal quantities of • Silver Nitrate, 5% • Hydrogen Peroxide, 30%. Counterstain for 5 minutes in Nuclear Fast Red Solution. Rinse in distilled water. Dehydrate in 95% alcohol, absolute alcohol and clear in Xylene, two changes each. Mount with Permount. Stain Results Calcium oxalate Black All other tissue elements Red Note: If a large amount of egg albumin section adhesive is present, numerous gas bubbles will develop during the light bulb staining treatment (step 2). This tends to inhibit the reaction. Should this occur, pour off the mixture and add fresh solution. References Pizzolato, P., J. Histochem. Cytochem., 12:333-336, 1964. Luna, L.G., (ed), Manual of Staining Methods of the AFIP, 3 rd edition, McGraw-Hill, NY, c 1968, p 177.
Platinum Blue, EM Stain
EMS22407 Description IBI Blue Platinum Stain Pt4N8H6O24C20 Preparation Adjust pH of distilled water with concentrated acetic acid to 3.5 Dilute PtBlue stock solution as supplied 1:100 in pH-adjusted water Use Incubate grid/Epon section on a drop of PtBlue 1:100 for 5 min at RT Wash on a water drop for about 2 min, repeat 5x Dry grid/section for approx. 10 min in an incubator Stain with lead citrate for 2 min at RT Jet-wash grid 5x (water stream from a plastic pastette) Suck up remaining water with a filter paper and dry thoroughly Cartilage ECM, constrasted with PtBlue + LC Cartilage ECM, constrasted with UA + LC HepG2 cells, constrasted with PtBlue + LC Depending on the sample, PtBlue can be nearly as good as uranyl acetate in its ability to add contrast. It does not work well with chondrocytes or T. brucei in our hands. Credits Schmidt, Katy. "Technical Note." IBI Labs. Web. 15 May 2015.
Poly-L-Lysine Solution
EMS19320 Intended Use: Poly-L-Lysine solution is intended for use as an adhesive subbing solution for immunoperoxidase and routine Histologic staining preparations. Backgroung & Principle: The loss of paraffin and frozen sections from slides has long been a problem during routine Histologic staining procedures. Various adhesives including albumin, gelatin and chrome alum have been applied to slides to minimise this loss. 1-3 different solutions of Poly-L-Lysine have been shown to be most effective in promoting adhesive of sections. 4-5 The polycationic nature of this molecule allows interaction with the anionic sites of tissue sections resulting in strong adhesive properties. 5. Poly-L-Lysine has been demonstrated as an effective tissue adhesive for use in various microwave procedures. Reagent: Poly-L-Lysine Solution. Poly-L-Lysine, 0.1% (w/v), in deionised water. Preservative added. Precautions: Poly-L-Lysine solution is for "In Vitro Diagnostic Use". Normal precautions exercised in handling laboratory reagents should be followed. Observe all local, state and Federal laws when disposing of waste. Refer to Material Safety Data Sheet for any updated risk, hazard or safety information. When diluted Poly-L-Lysine solution is prepared according to instructions, the maximum number of slides that can be coated is 900 slides/L of diluted solution. Exceeding 900 slides/L will affect the performance of the product. Preparation: Dilute Poly-L-Lysine solution 1:10 with deionised water prior to coating slides. Use plastic containers and graduated cylinders when mixing or storing solution and coating slides. Do not add fresh solution to used diluted solution. Storage & Stability: Dilute Poly-L-Lysine solution at room temperature (18-26°C). Reagent is stable until expiration date shown on label. Store diluted Poly-L-Lysine solution in refrigerator (2-8°C). The diluted solution is stable for at least three months. Filter diluted solution after use. Deterioration: Discard solutions if turbidity or bacterial growth develops. Procedure: Material required but not provided: Microscope slides, slide rack, plastic containers & graduated cylinder, drying oven (optional) Notes: Total slides coated should not exceed 900/L diluted solution. Do not add fresh solution to sued diluted solution. Slides must be clean before attempting this procedure. Clean with acid alcohol (i.e. 1% HCI in 70% ethanol) if necessary. Coating Procedure: Allow diluted Poly-L-Lysine solution to come to room temperature (18-26°C) before use. Place clean slides, a rack at a time, in diluted Poly-L-Lysine solution for 5 minutes. Increasing incubation time does not improve performance. Drain slides and dry in 60°C oven for one hour or at room temperature (18-26°C) overnight. References: Culling CFA, Allison RT, Bair WT: Cellular Pathology Technique, 4 th ed. Butterworth & Co., Ltd Boston , 1985, p98 Theory and Practice of Histological Techniques, JD Bancroft, A Stevens, Editors, Churchill Livingston, New York , 1982, pp 75-76 Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology, 3 rd ed., LG Luna, Editor, McGraw-Hill, New York, 1968, p 28 Mazia D. Schatten G, Sale W: Adhesion of cells to surfaces coated with Poly-L-Lysine. J Cell Biol 66:198, 1975 Huang Wm., Gibson SJ, Facer P, Gu J, Polak JM: Improved section adhesion for Immunocytochemistry using high molecular weight polymers of L-Lysine as a slide coating. Histochem 77:275, 1983
Price's Giemsa Stain Method
EMS26156 Components Giemsa Stock Solution Phosphate Buffer, pH 7.0 Acetic Acid, 0.2%, Aqueous Alcoholic Iodine, 2% Fixation 10% Buffered Neutral Formalin or Zenker's. Use no fixation technique that will destroy erythrocytes. Sections Paraffin @ 4µm Staining Deparaffinise with two changes at 2 minutes each in xylene. Rinse in absolute alcohol for two changes, 2 minutes each. Rinse in 95% Alcohol for two changes after a one minute immersion. If necessary to further remove mercuric chloride crystals, use an Alcoholic Iodine Solution which will be further removed in successive alcohols. Rinse through 80%, 70%, 50%, alcohols for one change, 1 minute each. Rinse in distilled water for 15 seconds. Buffer in the Phosphate Buffer Solution, for 30 minutes. Stain in the Giemsa Working overnight. To prepare Giemsa Working: • Geimsa Stock - 3 parts • Phosphate Buffer pH 7.0 - 97 parts Rinse in the buffer solution. Rinse in Acetic Acid, 0.2%, for one minute, absolute alcohol for two changes, 15 seconds each. This step should take only 90 seconds!!! Dip in xylene for two changes, two minutes each. Mount in Permount. Stains Blue Malarial parasites Blue Tissue Nuclei Dark Blue Bacteria Black Malarial Pigment Blue Schistosomic egg shells Pale Pink Collagen, etc. Pink-Rose Erthrocytes References AFIP Manual of Histological Staining Methods , 3 rd ed., L.Luna:
Prussian Blue Method for Haemosiderin
EMS26170, EMS26613 Components: Potassium Ferrocyanide 2% Aqueous Solution Hydrochloric Acid 2% Safranin O 0.2% Acetic Acid 1% Aqueous Fixation: 10% Buffered Neutral Formalin Sections: Paraffin Staining: Deparaffinise and hydrate to Distilled Water. Place sections in working staining solution for 30 minutes) at 60°C or for one hour at room temperature. Rinse in distilled water. Prepare working stain by mixing equal volumes of Potassium Ferrocyanide, 2% and Hydrochloric Acid, 2%. Prepare fresh just before use. Counterstain in Safranin O, 0.2% (#26170-03) for 2 minutes. Wash in Acetic Acid 1%. Dehydrate in 95% Alcohol, Absolute Alcohol, clear in Xylene , and mount in synthetic resin, such as DPX. Stain Results: Hemosiderin Blue or Green Nuclei Red Background Pink References: Perls, M., Virchow’s Arch. F. Path. Anat., 39:42-48, 1867. Highman, B., Arch. Path., 33:937-938, 1942. Lillie, R.D., Histopathologic Technic and Pratical Histochemistry, 3rd. ed., McGraw Hill, N.Y., 1965 Clark, G., (ed.), Staining Procedures, 3rd ed., Williams & Wilkins, Baltimore, p. 157, c. 1973
Prussian Blue Stain for Non-Haemoglobin Iron
EMS26161 Components: Potassium Ferrocyanide 2% Aqueous Solution Hydrochloric Acid 1% Wright Giemsa Stain Phosphate Buffer pH 6.5 Fixation: None Staining: Rapidly air dry smear or imprint. Flood slide with Wright Staining Solution. Dilute the stain (on slide) with Phosphate Buffer. Leave for a length of time suitable to stain strength and cellularity of film. Immerse slide in Prussian Blue Staining Solution for 10 minutes. Prepare Prussian Blue Staining Solution by mixing two parts of Potassium Ferrocyanide 2% and three parts of Hydrochloric Acid, 1%. Prepare fresh, just before use. Wash briefly in distilled water until pink colour appears, and drain dry. If desired dehydrate, clear and mount with synthetic resin. Stain Results: Particulate Iron and non- granular cytoplasm of many Macrophages Blue to Blue-green Other Comparable to original films stained with Wright Staining Solution References: Clark, G. Staining Procedures, Williams and Wilkins Co. Baltimore, 3rd ed., c. 1973, p.130. Sundberg, R., and Broman, H., Blood, 10160-6, 1955.
PTAH Method for Central Nervous System Tissue
EMS26658 Components: Potassium Permanganate 1% Phosphotungstic Acid-Haematoxylin Oxalic Acid 5% Aqueous Lugol's Iodine Solution Russels Modified Zenkers Fluid Fixation: Formalin, 10% Neutral Buffered. Sections: Paraffin @ 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Incubate sections in Russell Modified Zenker’s Solution (with Acetic Acid) at 60°C, 1 hour. Wash sections in running water, 15 minutes. Tone in Lugol’s Iodine15 minutes, then decolourise in 96% alcohol, 1 hour. Do not use Sodium Thiosulphate in this procedure because it will impair the further staining technique. Wash in distilled water for 5 minutes, and then oxidise with Potassium Permanganate Solution for approx. 5 minutes. Again decolourise the section using Oxalic Acid, 5%, 5 minutes, then wash in distilled water followed by running water for 5 – 10 minutes. Stain in the PTAH (Phosphotungstic Acid – Haematoxylin) reagent, 24 hours. Dip quickly into 95% alcohol, then further in absolute alcohol, two fast changes. Clear the slides in two changes of Xylene. Mount Stain Results: Nuclei, Fibrin, Fibroglia, Microglia Blue Elastic Fibrils, Coarse Purplish References: AFIP Manual of Histologic Staining Technique; ed. G. Luna., 3rd ed. McGraw-Hill Publications, c. 1968, p. 199.
Puchtler-Sweat Method for Basement Membranes
EMS26356 Components: Resorcin-Fuchsin Solution Nuclear Fast Red Solution Periodic Acid 0.5% Aqueous Sodium Bisulphite Solution Fixation: Carnoy’s Solution Sections: Paraffin at 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Periodic Acid, 0.5% for 5 minutes. Rinse in two changes of distilled water. Sodium Bisulphite Solution for 15 hours. Rinse in five or six changes of distilled water. Resorcin-Fuchsin Solution for 4 hours> Rinse in three changes of distilled water. Counterstain in Nuclear Fast Red for 4 hours. Rinse in 3 changes of distilled water. Dehydrate in 95% Alcohol, Absolute Alcohol, and clear in Xylene two changes each. Mount with Permount or DPX. Stain Results: Basement Membranes Black in cross section/Grey in tangential sections Nuclei Pink to Red References: Puchtler, H. and Sweat, F.: Stain Techn., 39:163, 1964. AFIP Manual of Histological Staining Methods, 3rd ed., ED. L. Luna: New York: McGraw Hill Publications, c. 1968, p. 73
Relative Acidophilia Stain
EMS26901 Components Phloxine B Staining Solution Tartrazine Saturated Solution In Cellosolve N-Butyl Alcohol Fixation Formalin Solutions Sections Paraffin Staining (Use 5 Slides) Deparaffinise and hydrate to distilled water. Stain for all 5 slides in Phioxine B, 0.5% Solution for 30 minutes. Remove one slide, rinse in water and 95% alcohol and hold in n-Butyl Alcohol. Rinse the remaining 4 slides quickly in water. Stain the remaining 4 slides in Tartrazine Solution and remove at 2,5, 10 and 20 minutes. Each time, rinse in 95% alcohol and hold in n-Butyl Alcohol following removal. When all 5 slides are in n-Butyl Alcohol, change to fresh n-Butyl Alcohol for 4 minutes, then clear and mount. Staining Results The least acidophilic elements stained in the phloxine control slide lose the red colour after 2 minutes in the tartrazine solution. The most acidophilic retain the phloxine after 20 minutes in the tartrazine. Reference Lendrum, A.C., J. Path. Bact ., 59:399-404, 1947. Clark ,G., (ed.), Staining Procedures , 3 rd edition, Williams & Wilkins, Baltimore , p. 163, c 1973.
Rhodanine Method for Copper
EMS26602 Components: Rhodamine Saturated Solution Mayer’s Haematoxylin Sodium Borate 5% Aqueous Solution Fixation: 10% Buffered Neutral Formalin Embedding: Cut paraffin sections at 6 to 10µm. (Thicker sections may stain better) Staining Procedure: Hydrate slides to distilled water. Incubate in Rhodanine working solution at 37°C for 18 hours. *To prepare Rhodanine Working Solution mix: Rhodanine Saturated Solution 6ml Distilled water 94ml. Note: Shake stock solution before measuring and mixing solutions and shake working solution when pouring it on slides. Wash well in several changes of distilled water. Stain in diluted Mayer’s Haematoxylin for 10 minutes. (To prepare diluted Mayer’s Haematoxlin mix - Mayers Haematoxylin 50ml Distilled water 50ml) Rinse with distilled water. Quickly rinse in Sodium Borate, 5%. Rinse well with distilled water. Dehydrate through 95% alcohol to absolute alcohol. Clear in Xylene, 2 changes each, and coverslip using a synthetic mounting medium. Note: The use of chemically clean glassware is necessary. Shake stock solution before measuring and mixing solutions and shake the working solution before pouring it onto the slides. Results: Copper – orange/red Nuclei – light blue Note: With low copper concentrations in tissue, slight fading may occur after coverslipping and the golden precipitate may be difficult to distinguish from lipofuscin. References: Sheehan and Hrapchack, Theory and Practice of Histotechnology. St. Louis, The Mosby Company 1980 p. 230.
Rhodinile Blue Stain
EMS26162 Fixation None Procedure To a few drops of heparinised or EDTA-treated blood, add an equal amount of Rhodinile Blue Stain. Swirl gently and let stand 2 minutes. Prepare thin smears and air dry. Smears can be examined as usual or can be mounted with a synthetic resin. Stain Results Deep Purple Heinz Bodies Yellow-Orange to Blue-Green Erythrocytes Blue Reticulocytes References Simpson, C.F., Carlisle, J.W., and Mallard, L.: Stain Techn., 45:221(1970) Clark, G.: Staining Procedures, Williams and Wilkins Company, Baltimore , 3 rd Ed., C. 1973, p. 131.
Sweat-Puchter Method For Amyloids
EMS26303 Components: Sirius Red F3Ba Solution 1% Mayer's Haematoxylin Borate Buffer, 0.1M, pH 9.0 Alkaline Alcohol Fixation: 10% Buffered Neutral Formalin Sections Paraffin at 6µm. 12µm sections may be used to demonstrate minute amyloid deposits. Staining is brighter on 12µm sections. Staining: Deparaffinise and hydrate to distilled water. Wash in running water 5 minutes. Mordant in Buffered Neutral Formalin overnight. Wash in water for 15 minutes. Alkaline Alcohol Solution for 1 hour. Rinse in distilled water for 10 seconds. Stain in preheated Sirius Red Solution for 90 minutes @ 60 °C. Rinse briefly in two changes of Borate Buffer. Wash in running water for 5 minutes. Stain in Haematoxylin Solution for five minutes. Wash in running water for at least 20 minutes to prevent fading of stain. Hydrate rapidly in Absolute Alcohol. Clear in three changes of Xylene. Mount. Stain Results: Amyloid Pink to Red Nuclei Blue Elasticia Pink to Red Background Unstained References: Sweat, F. and Puchtler, H.: Arch. Path. 80:613, 1965. AFIP Manual of Histologic Staining Methods, 3 rd edition, Ed L. Luna: New York : McGraw Hill Publications, c. 1968, p. 154.
Taft's Method for Nucleic Acid (1951)
EMS26777 Components: Methyl Green Pyronin Solution Differentiating Solution Fixation: Carnoy’s or absolute alcohol. Sections: Paraffin @ 4µm Staining Procedures: Deparaffinise and hydrate to distilled water. Immerse in the Methylene Green-Pyronin Stain for 10 minutes. The stain may be filtered before use. Rinse in distilled water, twice briefly. Carefully blot dry with several thicknesses of smooth filter paper. Place in the Differentiating Solution for 1 – 2 minutes, then hydrate again in a fresh change of the solution. If the slides are left up to 4 – 5 minutes in the second t-butyl alcohol solution, additional differentiation should not take place. Clear in Xylene, 2 changes, 10 minutes each. Mount in Permount Stain Results: DNA, Deoxyribonucleic Acid Blue Green RNA, Ribonucleic Acid Bright Red Microwave Procedures: Deparrafinise and hydrate to distilled water. Place slide in 40 ml filtered Methyl Green-Pyronin. Solution in a glass coplin jar; cover with a loose plastic cap. Heat in microwave oven for 15 seconds. Rinse in distilled water and blot with filter paper. Differentiate the damp section in Differentiating Solution, 2 changes, 2-3 dips. Clear in Xylene, 2 changes and mount. Staining Results: Chromotin DNA Blue Green RMA Red Rose Other Elements Pale Pink References: Taft, E.B. Stain Tech., 26:205-212, 1951.
Taylor's Method for Bacteria
EMS26108 Components: Harris Haematoxylin Hucker’s Crystal Violet Basic Fuchsin Stock Sol. 0.1% Gram’s Iodine Solution Lithium Carbonate Solution Acid Alcohol, 1% Acetone-Alcohol, 1:1 Acetone Picric Acid-Acetone Sol., 0.1% Acetone-Xylene I Acetone-Xylene II Fixation: 10% Buffered Neutral Formalin Section: Paraffin @ 6µm Staining Procedure: Use Control Slide Deparaffinise and hydrate to distilled water. Harris Haematoxylin for 5-10 minutes. Wash in running distilled water 1 minute. Differentiate in Acid Alcohol. Wash in running water 3 minutes. Wash in Lithium Carbonate Solution to intensify blue ***From this point carry only one slide at a time. Wash in running water, 5 minutes. Hucker's Crystal Violet two minutes. Wash quickly in water. Mordant in Gram's Iodine1 minute. Wash in water. Blot with filter paper moistened with water, but do not allow to dry. Decolorize in Acetone:Alcohol until no more blue comes off. Blot with filter paper moistened with Acetone-Alcohol, but do not allow to dry. Basic Fuchsin Working (or 5.0ml Basic Fuchsin Stock and 60ml Distilled water) for 3 minutes. Wash in water. Blot with filter paper moistened with water, but do not allow to dry. Dip in Acetone until section begins to decolourise. Differentiate in Picric Acid-Acetone, (A-108-9) until section becomes reddish-brown-yellow (15 seconds). Pass quickly through Acetone-Xylene I, then Acetone-Xylene II Clear in Xylene, two changes. Mount. Results: Gram-Positive Organisms blue to blue-black Gram-Negative Organisms bright red Nuclei brownish red Erythrocytes red to yellow-green Necrotic Tissue yellow-green Cytoplasm yellow Connective Tissue red References: AFIP Manual of Histological Staining Methods, 3rd ed., Ed. L. Luna: New York: McGraw Hill Publications, c. 1968, p. 226. Clark, G.: Staining Procedures, Williams and Wilkins Company, Baltimore, 3rd Ed., c. 1973, p. 320 Taylor, R.D., Amer. J. Clin. Path., 46:472 (1966).
Thomas' Method for Malarial Parasites
EMS26155 Components: Phloxine B Solution 0.5% Methylene Blue Azure B Solution Acetic Acid 0.2% Aqueous Fixation: Any well fixed tissue. Section: Paraffin at 6µm Staining: De-paraffinised and hydrate to distilled water. Stain for two minutes in Phloxine B Solution, 0.5%. Rinse well in distilled water. Stain for 1 minutes in Methylene Blue-Azure B Solution. Differentiate in Acetic Acid 0.2%. Complete the differentiation in three changes of 95% alcohol. Dehydrate in absolute alcohol and clear in xylene, two changes each. Mount with Permount. Stain Results: Nuclei Blue Plasma Cell Cytoplasm Blue Malarial Parasites Blue Erythrocytes Pink Other Tissue Elements Shades of rose to red References: Thomas, J.T. Stain Techn., 28:311-312, 1953 Luna, L.G. (ed.) Manual of Histologic Staining Methods of the AFIP, 3rd edition, McGraw-Hill, NY., p. 129, c. 1968
Toren's Method For Mast Cells
EMS26472 Components Giemsa Stain Solution Acetic Acid, O.5%, Aqueous Pollacks’s Trichrome Solution Fixation 10% Buffered Neutral Formalin Section Paraffin, @ 6µm Staining Procedure Deparaffinise and hydrate to distilled water. Stain in *Working Giemsa Solution for 75 minutes. Rinse in distilled water. ***Prepare Working Giemsa Solution by adding 40 drops of Giemsa Stock Solution to 40 ml of distilled water and mixing thoroughly. Prepare fresh, immediately before use. Place in Acetic Acid 0.5% for 5 to 10 seconds. Rinse in distilled water. Stain in Pollack's Trichrome Solution for 10 to 15 seconds. Rinse in Acetic Acid 0.5% until no more stain washes off, 3 to 5 seconds. Dehydrate in 95% Alcohol, Absolute Alcohol and clear in Xylene. Mount with Permount. Results Mast Cell Granules deep blue Collagen & Cartilage blue green Muscle & Elastic Fibres red Fibrin purple Bone green Colloid, Keratin & Erythrocytes orange red to orange References Toren, D.A.: Stain Techn ., 38:249-250, 1963. Luna, L., (ed.), Manual of Histologic Staining Methods of the AFIP , 3 rd . ed., McGraw-Hill , New York , c. 1968, pp. 116-117.
Truant's Fluorescent Method For Acid Fast Organisms
EMS26102 Components Weigert’S Iron Haematoxylin Solution A Weigert’S Iron Haematoxylin Solution B Auramine-Rhodamine Staining Solution Acid Alcohol 1% Fixation 10% Buffered Neutral Formalin Section Paraffin, @ 6µm Staining Procedure Use control slide Deparaffinise and hydrate to distilled water. Weigert's Iron Hematoxylin solution for 10 minutes. Prepare working Weigert's Haematoxylin by mixing 1 vol. of Weigert's Haematoxylin A, with 1 vol. of Weigert's Haematoxylin B. Wash in running water for 10 minutes. Auramine-Rhodamine Solution, @ 60 °C for 10 minutes, or 37 °C for 15 minutes, or 25 °C for 20 minutes. Wash in tap water. Remove excess stain with Acid Alcohol, 1%, for 2-3 minutes. Wash in running water. Rinse in distilled water. Dehydrate in 95% alcohol, absolute alcohol, and clear in Xylene, two changes each. Mount with Permount. Results Acid Fast fluorescent reddish yellow Artifacts fluorescent yellow References Traunt, J.P. Henry Ford Hospital Med. Bull., 10:287, 1962. AFIP Manual of Histological Staining Methods, 3 rd ed. , Ed. L. Luna: New York : McGraw Hill Publications, c. 1968, p. 219. Clark, G.: Staining Procedures, Williams and Wilkins Co., Baltimore , 3 rd Ed., c. 1973, p. 225.
UranyLess EM Stain – Protocols of Use and FAQs
EMS22409 CLASSIC CONTRAST This protocol is used for double staining with UranyLess / Lead citrate on ultrafine sections. This protocol is adapted to biologic samples that have been fixed with glutaraldehyde, osmium, or ruthenium and embedded in an epoxy type resin (Epon, Araldite, Spurr) or acrylic type (LRWhite, HM20). Staining protocol: Place a drop of UranyLess on parafilm or any other hydrophobic slide. Place the grid on the UranyLess drop for 1 to 2 minutes. Blot the grid on a filter paper and then wash in room temperature distilled water. Let it dry. After drying, go to the lead citrate staining according to Reynolds method (1963). Place the grid on the lead citrate drop according to the Reynolds method, for 1 minute. Blot the grid on a filter paper before rinsing with room temperature distilled water. Let it dry. Technical Tip: UranyLess is not air or light sensitive, unlike Uranyl Acetate. After lead citrate, drain immediately in a freshly prepared room temperature distilled water bath or wash with 0.01N of NaOH solution. Cold water is charged with CO2 and causes precipitate on the grid. If there is a precipitate in the solution, filter it prior to use. If solution was refrigerated, allow solution to return to room temperature prior to use. Do not keep lead citrate refrigerated. NEGATIVE STAINING Negative staining is a very useful technique in electron microscopy. It allows characterization of isolated particles of morphology as bacteria, virus, protein, nanoparticles, liposomes, exosomes, etc. Staining Protocol #1: On a piece of parafilm or any other hydrophobic carrier, place a drop of your solution (~ 10µl) and a UranyLess drop. Using our fine tweezers, place your sample drop on a formvar-carbon coated grid. for about 1 minute. Blot your grid using filter paper. Place your grid on the UranyLess solution for 1 minute. Blot, let it dry for 5 minutes and observe under the microscope. Technical Tip: If the staining is too intense, wash with room temperature water for 1 minute. Staining Protocol #2 (Debra M. Townley, Baylor College of Medicine): One drop of suspension of 300 mesh formvar coated grids – 30 minutes Wick away excess suspension with paper arrow (Watman No.1) Place one drop of Uranyless on the grid before it is completely dry Stain for 3 minutes Wick away excess stain with paper arrow (Watman No.1) Allow the grids(s) to dry completely before examination on the TEM Times can be altered (suspension time, stain time) to suit the "look" you desire FREQUENTLY ASKED QUESTION What is UranyLess made from? UranyLess is a solution ready for use, a mix of lanthanides (rare-earths). How is UranyLess sold? In an aqueous solution (water). What is it's shelf life? One year. What are the storage conditions for UranyLess? Store it at room temperature away from direct sunlight. Does it need to be diluted? No, it is sold ready for use. What is its pH? UranyLess pH is 6.8-7. How to stain with UranyLess? Simply drop UranyLess on your grid, and wait a minute. Dry, then contrast with lead citrate according to Reynolds method. How do I avoid precipitates (dirt) on the grids? Precipitates form when lead citrate is washed with cold water which is loaded with CO2. Use water that has never been at low temperature and never store lead citrate at low temperature. Is it the same protocol for every kind of tissue (animal, plant, marine)? Yes it is - a double stain of UranyLess plus Lead citrate. Does it adjust to every kind of resin? Yes, it operates with every kind of resin (Epon, Araldite, Spurr). Can it be used on negative staining? Yes, it can be used on negative staining. Can it be used for bloc contrast? Some tests are in progress. Is it adapted to a cryo use? No, because it is prepared in water. However, we are currently developing many formulations of UranyLess, including ethyl UranyLess and acetone UranyLess, the latter being the best-adapted to cryogenic use. Is it efficient on marine material? Yes. How is UranyLess packaged? We sell UranyLess in an airless 30ml bottle and also in a brown 200ml bottle. How does an airless bottle operate? Its use is very simple; simply push on the head of the bottle to get a drop. When you release, the bottle back pump actuator lifts up. It prevents any air inlet in the bottle. What is the advantage of an airless bottle? It is a bottle in which air never enters. Some products, such as lead citrate, are atmospheric CO2 sensitive. Thanks to this system, those products have a longer shelf life. It also allows the product to be deposited drop by drop, quickly, cleanly and in any position. Can it be used with automated staining equipment? Yes, the 200ml bottle is available for use with automated staining equipment. When using UranyLess for automated staining, do not wash longer than 10 minutes or you run the risk of losing all contrast.
Van Gieson's Method for Collagen Fibres
EMS26046 Components: Van Gieson's Solution Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B Fixation: Any Sections: Paraffin @ 6µm or Celloidin Staining: Deparaffinise and hydrate to distilled water. Weigert's Working Hematoxylin* for 10 minutes. Wash in distilled water. Stain 1-3 minutes in Van Gieson's Solution. Dehydrate in 95% alcohol, absolute alcohol, two changes each. Clear in two changes xylene. Mount with Permount or equivalents. *Weigert's Working Solution is prepared by mixing equal parts of Weigert's Iron Haematoxylin A and Weigert's Iron Haematoxylin B Staining Results: Collagen: Red Smooth and striated muscle: Yellowish to Brownish Cornified epthelium: Yellow Hyalin: Yellow Nuclei: Blue to Black References: Clark, G., (ed.) Staining Procedures 3 rd Ed., Williams & Wilkins Co. Baltimore, c. 1973, p.50 Luna, L., (ed.) AFIP Manual of Histological Staining Methods, 3 rd Ed., New York . McGraw Hill Publication, c. 1968, p. 76. Van Gieson, I. , New York Med. J., 50:57-60, 1889. Mallory, F.B., Pathological Technique, W.B. Saunders Co., Philadelphia 1938.
Van Gieson's Solution
EMS26046 VAN GIESON’S METHOD FOR COLLAGEN FIBRES Fixation: Any Sections: Paraffin @ 6µm or Celloidin Staining: Deparaffinise and hydrate to distilled water Weigert's Working Haematoxylin* for 10 minutes. Wash in distilled water. Stain 1-3 minutes in Van Gieson's Solution (EMS26046-05) Dehydrate in 95% alcohol, absolute alcohol, two changes each. Clear in two changes xylene. Mount with Permount (EMS17986-01) or equivalents. *Weigert's Working Solution is prepared by mixing equal parts of Weigert's Iron Hematoxylin A (EMS26044-05) and Weigert's Iron Hematoxylin B (EMS26044-15) Staining Results: Collagen: Red Smooth and striated muscle: Yellowish to Brownish Cornified epthelium: Yellow Hyalin: Yellow Nuclei: Blue to Black References: Clark, G., (ed.) Staining Procedures 3 rd Ed., Williams & Wilkins Co. Baltimore, c. 1973, p.50 Luna, L., (ed.) AFIP Manual of Histological Staining Methods, 3 rd Ed., New York . McGraw Hill Publication, c. 1968, p. 76. Van Gieson, I. , New York Med. J., 50:57-60, 1889. Mallory, F.B., Pathological Technique, W.B. Saunders Co., Philadelphia 1938. VERHOEFF’S ELASTIC TISSUE STAIN Fixation: 10% Formalin or Zenker's Stock Solution (EMS64123) Section: Paraffin @ 6µm Staining Procedure: Deparaffinise and hydrate to distilled water. Place in staining solution for 15 minutes. Staining solution should be prepared fresh. Combine: 25ml Iron solution. 10ml Potassium Iodide, 3%, aqueous. 3.5ml Alcoholic Haematoxylin, 10%. Use once and discard. Wash in tap water. Differentiate in Iron Solution, diluted 1-4 with distilled water Counter stain with Van Gieson's Solution, 3 - 5 minutes. Dehydrate, clear and mount. Results: Elastic Fibres: Intensely blue-black Nuclei: Blue to black Collagen: Red Smooth and Striated Muscle: Brownish References: Clark, G., Ed: Staining Procedures. 3 rd ed., Baltimore : William and Wilkens Co. c. 1972, p. 61 AFIP Manual of Histologic Staining Techniques. 3 rd ed., Ed. G.. Luna: New York : McGraw-Hill Publications, C. 1968, p. 76
Verhoeff's Elastic Tissue Stain
Components: Iron solution Potassium Iodide, 3%, aqueous Alcoholic Haematoxylin, 10% Van Gieson's Solution Fixation: 10% Formalin or Zenker's Stock Solution Section: Paraffin @ 6µm Staining Procedure: Deparaffinize and hydrate to distilled water. Place in staining solution for 15 minutes. Staining solution should be prepared fresh. Combine: 25ml Iron solution, 10ml Potassium Iodide, 3%, aqueous., 3.5ml Alcoholic Haematoxylin, 10%. Use once and discard. Wash in tap water. Differentiate in Iron Solution, diluted 1-4 with distilled water Counter stain with Van Gieson's Solution, 3 - 5 minutes. Dehydrate, clear and mount. Results: Elastic Fibres: Intensely blue-black Nuclei: Blue to black Collagen: Red Smooth and Striated Muscle: Brownish References: Clark, G., Ed: Staining Procedures. 3 rd ed., Baltimore : William and Wilkens Co. c. 1972, p. 61 AFIP Manual of Histologic Staining Techniques. 3 rd ed., Ed. G.. Luna: New York: McGraw-Hill Publications, C. 1968, p. 76
Verhoff's Van Geison Stain
EMS26374 Components: Alcoholic Haematoxylin 5% Ferric Chloride 10% Weigert's Iodine Ferric Chloride 2% Sodium Thiosulfate 5% Aqueous Van Gieson's Solution Fixation: Any common well-fixed tissue – 10% Buffered Neutral Formalin or Russell’s Zenker is preferred. Sections: 5µm sections. Staining Procedures: Hydrate slides to distilled water. Stain in Verhoff’s Working Solution* for 1 hours. Tissue should be completely black. Depth of stain required for the elastic fibres. (Nuclei may be stained in Weigert's Iron) * To prepare Verhoff’s Working Solution, add in order the following reagents – prepare fresh! Alcoholic Haematoxylin, 5% - 20ml Ferric Chloride, 10% - 8ml Weigert’s Iodine Solution - 8ml Mix well, solution should be jet black. Used immediately. At the conclusion of staining time, pour off the Verhoff mixture into a container and save it until after the differentiation process has been properly completed. If it should prove necessary, the saved solution may be used. Rinse in tap water with 2 or 3 changes. Differentiate in Ferric Chloride, 2%. Agitate Slide gently during the process. Stop differentiation with several changes of tap water and check microscopically for black elastic fiber staining and gray background. Repeat Ferric Chloride, 2% treatment and tap water rinses as necessary for adequate demonstration. Kidney and myometrium are good controls. If the elastic fiber staining is too pale, re-stain in the saved Verhoffs solution for 30 minutes and then proceed with the subsequent Van Gieson counterstain can extract the elastic stain somewhat. Wash slides in tap water. Treat with Sodium Thiosulfate, 5% for 1 minute. Discard solution. Wash in running tap water for 5 minutes Counterstain in Van Gieson Solution for 3 to 5 minutes. Dehydrate, clear in Xylene and coverslip, using a synthetic mounting medium. Stain Results: Elastic Fibres Blue-black to Black. Fine elastic fibrils may not be stained with this methods Nuclei Blue to Black Collagen Red Other Tissue Elements Yellow References: Sheehan, Dezna C., Hrapchack, Barbara B. Theory and Practice of Histotechnology, 2nd ed. The C.V. Mosby Company, St Louis, p. 196 – 197. c. 1980.
Vogt's Method for Nerve Cell Products
EMS26671 Components Cresyl Violet Acetate 2% Aqueous Acetate Buffer Cresyl Violet Acetate Working Solution Fixation Formalin, 10% Neutral Buffered Sections Paraffin @ 6µm Staining Deparaffinise and carry through absolute alcohol, two changes. Let stand in absolute alcohol for two hours. Stain in Cresyl Violet Acetate Working Solution for 40 – 60 minutes. Differentiate rapidly in 95% alcohol, then dehydrate in absolute alcohol, and clear in xylene, two changes each. Mount with Permount. Staining Results Nissl Substances Intense Purple Nuclei Purple Background Clear References AFIP Manual of Histologic Staining Technique; ed. L. Luna., 3rd ed. McGraw-Hill Publications, c. 1968, p. 212.
Weigert's Iron Haematoxylin With Metachromatic Dyes Fixation
EMS26758 Components Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B Fast Green FCF Solution, 0.02% Bismark Brown Y Solution, 0.1% Safranin O, 0.1% Acetic Acid, 1% Aqueous Fixation Formalin, Bouin's, Carnoy, Orth's or Zenker's Sections Paraffin Staining Deparaffinise and hydrate to distilled water. Stain in working Weigert's Solution* for 6 minutes. Wash in water. *Prepare working solution by mixing equal volumes of Weigert's Iron Haematoxylin A and Weigert's Iron Haematoxylin B. Stain in 0.02% Fast Green FCF for 3 minutes. Wash in 1% Acetic Acid. Stain for 4-6 minutes in 0.1% Bismarck Brown or in 0.1% Safranin O. Dehydrate with 95% alcohol and 2 changes of absolute alcohol. Clear with Alcohol: Xylene, 1:1, 1 change and two changes of Xylene. Mount with a synthetic resin. Staining Results Nuclei Black Cytoplasm Grey-green Mucus, cartilage and mast cell granules Yellowish brown, deep red or orange -red depending on dye used. References Weigert, K., Z. Wissen. Mikr., 21:1-5, 1904. Lillie, R.D., Histopathologic Technic and Practical Histochemistry , 3 rd , edition, McGraw-Hill , NY , 1965. Clark , G., (ed.), Staining Procedures , 3 rd edition, Williams & Wilkins, Baltimore , p. 40, c 1973.
Weigert's Resorcin Fuchsin
EMS26370 Components: Resorcin-Fuchsin Solution Van Gieson's Solution Weigert's Iron Haematoxylin A Weigert's Iron Haematoxylin B Acid Alcohol 1% Fixation: Any common reagent may be used. Sections: Paraffin at 6µm. Staining Procedures: Deparaffinise and hydrate to distilled water. Stain in Resorcin-Fuchsin Solution for 1 – 12 hours depending upon the depth of stain required for the elastic fibres. (Nuclei may be stained in Weigert Iron Hematoxylin Working Solution* prior to the Resorcin Fuchsin rather than alter as is described here). * To prepare working solution, (prepare fresh just before use) mix equal parts of Weigert’s Iron Haematoxylin A and Weigert’s Iron Haematoxylin B. Clear the stain in 95% Alcohol. Acid Alcohol may be used to differentiate any slide that has stained diffusely wash in tap water. Stain slides in Waeigert’s Iron Haematoxylin Working Solution (see above) and then counterstain in Van Gieson’s Solution Rinse in distilled water to remove excess Van Gieson’s Stain from slides. Differentiate and hydrate the slides in two changes each of 95% Alcohol and absolute alcohol. Clear the slides in two changes of Xylene. Mount. Stain Results: Elastic Tissues Dark Blue Nuclei Dark Blue Collagen Red Pink Miscellaneous Tissue Elements Yellow References: Mallory, F.B.: Pathological Technique; New York, Hafner Publishing Co., c. 1961, p. 168 Clark, G.: Staining Procedures: Baltimore: Williams and Wikins Co., 3rd ed., c. 1973, p. 60 Luna, L.G.: AFIP Histologic Staining Methods: New York: McGraw-Hill Book Co., 3rd ed., c. 1968, p.80.
Wilder Modification of the Bielschowsky Stain for Reticulum
EMS26392 Components: Silver Nitrate 10.2% Aqueous Solution Ammonium Hydroxide, concentrated Sodium Hydroxide 3.1% Formalin 40% Harris' Haematoxylin Nuclear Fast Red Phosphomolybdic Acid 10% Uranium Nitrate 1% Gold Chloride Sodium Thiosulfate 5% Aqueous Solution Van Gieson's Solution Fixation: 10% Buffered Neutral Formalin or Russell’s Zenker Sections: Cut paraffin at 6µm sections. Staining Procedures: Deparaffinise and hydrate slides to distilled water. Oxidise in Phosphomolybdic Acid, 10% for 1 minute. Rinse well in distilled water or sunning water to prevent the cells from holding yellow colours. Sensitise in Uranyl Nitrate, 1% for 1 minute or less. Rinse 10 – 20 seconds in distilled water. Stain in fresh Ammonical Silver for one minute (changing solution frequently). The solution may be prepared as follows: Add concentrate Ammonium Hydroxide to 5.0ml of Silver Nitrate, 10.2% drop by drop, until the forming precipitate is almost dissolved. Add 5.0ml of Sodium Hydroxide, 3.1% and just dissolve the resultant precipitate by slowly adding concentrated Ammonium Hydroxide dropwise. Bring volume up to 50ml with distilled water. Use at once or filter and store in refrigerator in amber bottle. Dip quickly in 95% alcohol and transfer to freshly prepared reducing solution. Prepare the reducing solution as follows: Mix 50.0ml distilled water, 1.5ml Uranyl Nitrate, 1% and 0.5ml of Formalin, 40% neutralised. Prepare just before use and change frequently during the process. Rinse in distilled water. Tone in Gold Chloride, 0.2% approx. 1 minute, until the yellow brown colour leaves the sections and they turn lavender. Check the slides at this point individually with a microscope to prevent over toning; this could lead to red sections. Rinse in distilled water and transfer to Sodium Thiosulphate, 5% – approx. 1 minute – to fix the colour. Wash in tap water. Counter-stain in Nuclear Fast Red if desired, rinsing well in distilled water. Alternatively Harris' Haematoxylin with either Van Gieson's or phloxine can be used as a counterstain. Dehydrate in 95% and 100% alcohol, 2 changes and clear in xylene, 2 changes. Mount. If celloidin is used, clear in oil of origanium followed by 25% alcohol, usually mounted in balsam with Permount. Stain Results: Reticulum fibre (If beading is observed and/or dark background, replace the ammonium Hydroxide Silver Nitrate solution and the reducing solution Black Collagen Rose Other tissue elements Red References: Wilder, H.C., Amer. J. Clin. Path., 11:817, 1935. Mallory, F.B. Pathological Technique, Phil., W.B. Saunders & Co, c. 1938 AFIP Manual of Histological Staining Techniques, NY, McGraw-Hill Publ., c. 1968/3rd ed. P.92
Woelck’s Method for Myelin Sheath
EMS26684 Components: Ferric Ammonium Sulphate 2.5% Aqueous Alcoholic Haematoxylin Stock 10% Lithium Carbonate Saturated Solution Fixation: Formalin 10% Neutral Buffered Sections: Paraffin @ 15µm Staining Procedures: Deparaffinise and hydrate to distilled water. Mordant in Ferric Ammonium Sulphate, 2.5% overnight. Rinse in two changes of distilled water. Working Haematoxylin solution for 2 hours. To prepare working solution mix the following ingredients in order listed just before use: 45.0ml Distilled water 10.0ml Haematoxylin Stock – filtered 7.0ml Lithium Carbonate, Saturated – filtered 45.0ml Distilled water Rinse in two changes of distilled water. Differentiate in 80% Alcohol until background is clear and myelin stands out sharply. Dehydrate with 95% Alcohol, absolute alcohol, and clear in Xylene two changes each. Mount. Stain Results: Glial Cells, Nucleoli of Nuerons Black Myelin Sheath Blue Background Clear References: AFIP, Manual of Histological Staining Methods, 3rd ed., Ed. L. Luna: New York: McGraw-Hill Publications. C. 1968, p. 206.
Wolbach’s Giemsa Method
EMS26132, EMS26773 Components: Giemsa Stock Solution Giemsa Working Solution Rosin Alcoholic Stock Solution Rosin Alcoholic Working Solution Lugol's Iodine Gram's Iodine Solution Sodium Thiosulfate 5% Aqueous Fixation: Zenker’s or other well fixed tissue fixatives. Sections: Paraffin @ 6µm Staining Procedures: Deparaffinise and hydrate to distilled water. Remove “Zenker Crystals” by placing in Lugol’s Iodine or Gram’s Iodine for 15 minutes. Rinse in water, place in Sodium Thiosulfate, 5% for 3 minutes and wash in running water for 15 minutes. Rinse in distilled water and stain in Working Giemsa Solution* overnight. Working Giemsa Solution may be made from the stock solution by mixing: Giemsa Stock - 1.25ml Methanol - 1.50ml Distilled water - 50.0ml NOTE: Giemsa stain colours more effectively in tissue at an acid pH. If this has not occurred in the preparation or decalcifications steps, wash in an acid alcohol, and begin stain. Differentiate in Rosin Alcohol Working until the sections are purplish-pink color. Check under microscope. Dehydrate in two changes of absolute alcohol and clear in two changes of Xylene. Mount with Permount. Stain Results: Nuclei, Bacteria Blue Rickettsia Purple Collagen, other Tissue Elements Pink to Rose References: Wolbach, S.B., Todd, J.L., and Paltrey, F.W., The Etiology of Pathology Typhus. Harvard University, University Press, Cambridge, MA, p. 13-14, c. 1922.. Luna, L.G., (ed). Manual of Histologic Staining Methods of the AFIP 3rd edition. McGraw-Hill, NY, p. 119, c. 1968.
Wright Stain Method
EMS26060 Components: Wright Stain Solution Phosphate buffer pH 6.5 Fixation: Streak thin (approximately one cell thick) smears across a sterile slide by means of a second slide or cover glass. Air dry quickly. Staining: Place 1.0ml of the Wright Stain Solution upon the smear 1 – 3 minutes. Add 2.0ml distilled water or Phosphate buffer pH 6.5 and let stand twice as long as in step 1. Rinse stained smear with water or the Phosphate buffer pH 6.5 until the edges show faintly pinkish-red. For a Giemsa appearance stain 10 minutes in one volume Wright Stain and 4 volumes Phosphate buffer, pH 6.5. Blot dry very carefully. Stain may be adjusted by further dilution or in the timing of either before or after dilution in the above procedure. Stain Results: Erthrocytes yellowish-red Polymorphonuclears: Nucleus dark purple Polymorphonuclears: Granules reddish-lilac Polymorphonuclears: Cytoplasm pale-pink Eosinophiles: Nuclei blue Eosinophiles: Granules red to orange-red Eosinophiles: Cytoplasm blue Basophiles: Nucleus purple to dark blue Basophiles: Granules very dark purple Lymphocytes: Nuclei dark purple Lymphocytes: Cytoplasm sky blue Platelets violet to purple granules References: Conn, J I Biological Stains, 8th Ed.,Williams and Wilkins Co. Baltimore, 1969. Clark, G (ed.) Staining Procedures, Williams and Wilkens Co., Baltimore, 3rd Ed., c 1972, p. 122
Wright Stain Method
EMS26060 Fixation: Streak thin (approximately one cell thick) smears across a sterile slide by means of a second slide or cover glass. Air dry quickly. Staining: Place 1.0ml of the Wright Stain Solution upon the smear 1 – 3 minutes. Add 2.0ml distilled water or Phosphate buffer pH 6.5 and let stand twice as long as in step 1. Rinse stained smear with water or the Phosphate buffer pH 6.5 until the edges show faintly pinkish-red. For a Giemsa appearance stain 10 minutes in one volume Wright Stain and 4 volumes Phosphate buffer, pH 6.5. Blot dry very carefully. Stain may be adjusted by further dilution or in the timing of either before or after dilution in the above procedure. Results: Erthrocytes yellowish-red Polymorphonuclears: Nucleus dark purple Polymorphonuclears: Granules reddish-lilac Polymorphonuclears: Cytoplasm pale-pink Eosinophiles: Nuclei blue Eosinophiles: Granules red to orange-red Eosinophiles: Cytoplasm blue Basophiles: Nucleus purple to dark blue Basophiles: Granules very dark purple Lymphocytes: Nuclei dark purple Lymphocytes: Cytoplasm sky blue Platelets violet to purple granules References: Conn, J I Biological Stains, 8th Ed.,Williams and Wilkins Co. Baltimore, 1969. Clark, G (ed.) Staining Procedures, Williams and Wilkens Co., Baltimore, 3rd Ed., c 1972, p. 122
Wright-Giemsa Stain Method
EMS26149 Components: Wright Giemsa Stain Phosphate Buffer, pH 6.5 Fixation: Streak thin (approx. one cell thick) smears across a sterile slide by means of a second slide or cover glass. Air-dry quickly. Staining Procedure: Place 1.0ml of the Wright-Giemsa Stain upon the smear, in sufficient quantity to cover the entire surface, for 3-4 minutes. Add 2.0ml distilled water or Phosphate Buffer, pH 6.5 and let stand twice as long as in step 1. Rinse stained smear with water or the Phosphate Buffer, pH 6.5 until the edges show faintly pinkish red. The film is allowed to dry in the air. The preparation may be blotted gently to hasten drying. Permanent mounts can be made with mounting medium. Stain Results: Erythrocytes Pink-Tan Eosinophiles Red Lymphocytes: Granules Red-Purple Lymphocytes: Cystoplasm Blue Neutrophils: Granules Purple-Pink References: Lillie, R.D.: Factors influencing the staining of blood films and the role of methylene violet. J. Lab. Clin. Med. 29: 1181, 1944 Lillie, R.D., H.J. Conn’s Biological Stains, 8th edition, William’s & Wilkins, Baltimore, p. 438, 1972.