Fuchsin staining with naoh clearing for lignified elements of whole plants or plants organs

Three methods that are adapted to the various consistencies of plants are as follows: 1. Samples are placed for 10-14 hr at 60° C in a 1% aqueous solution of basic fuchsin, to which 10 gm of solid NaOH per 100 ml are added. 2. Samples when taken out of 95% alcohol are placed in a 1% solution of basic fuchsin in 95% alcohol for 24 hr; after washing in water, they are placed in a 15% solution of NaOH at 60° C until cleared. 3. Samples are placed in a 15% aqueous solution of NaOH at 60° C until cleared, then for 24 hr at 60° C in 15% NaOH containing basic fuchsin. After being stained and cleared by one of these three methods, the samples are rinsed in water, dehydrated and then passed into a mixture of absolute alcohol and concentrated HC1 (3:1) for 1-15 min, rinsed in absolute alcohol, cleared in xylene and mounted in Canada balsam. The lignified tissues appear red; the others, transparent.     

A Copper Sulfate-Silver Nitrate Method for Nerve Fibers of Planarians

For staining in toto, planarians are fixed in a mixture of 10 ml of commercial formalin, 45 ml of 95% ethanol and 2 ml of glacial acetic acid. After treatment with 70% ethanol 3-10 days, they are washed in distilled water and immersed in 10% CuSO₄. 5H₂O for 3 hr at 50° C, transferred without washing to 1% AgNO₃ for 1.0-1.5 hr at 50° C; and then developed in: 10 ml of 1% pyrogallol, 100 ml of 56% ethanol and 1 ml of 0.2% nitric acid. Gold toning, 5% Na₂S₂O₃ and dehydration follow as usual. For staining sections, material is fixed in the same fixative, embedded in paraffin and sectioned at 10 μ. After bringing sections to water, they are immersed in 20% CuSO₄. 5H₂O for 48 hr at 37° C; then rinsed briefly in distilled water and placed in 7% AgNO₃ for 24 hr at 37° C. They are washed briefly in distilled water and reduced in: hydroquincne, 1 gm; Na₂SO₃, 5 gm and distilled water 100 ml. Gold toning, followed by 5% Na₂S₂O₃ and dehydration completes the process. Any counterstaining may follow.     

Differentiated Staining of Osmium Tetroxide-Fixed, Vestopal-w-Embedded Tissue in thin Sections

Tissues were fixed at 20° C for 1 hr in 1% OsO₄, buffered at pH 7.4 with veronal-acetate (Palade's fixative), soaked 5 min in the same buffer without OsO₄, then dehydrated in buffer-acetone mixtures of 30, 50, 75 and 90% acetone content, and finally in anhydrous acetone. Infiltration was accomplished through Vestopal-W-acetone mixtures of 1:3, 1:1, 3:1 to undiluted Vestopal. After polymerisation at 60° C for 24 hr, 1-2 μ sections were cut, dried on slides without adhesive, and stained by any of the following methods. (1) Mayer's acid hemalum: Flood the slides with the staining solution and allow to stand at 20°C for 2-3 hr while the water of the solution evaporates; wash in distilled water, 2 min; differentiate in 1% HCl; rinse 1-2 sec in 10% NH,OH. (2) Iron-trioxyhematein (of Hansen): Apply the staining solution as in method 1; wash 3-5 min in 5% acetic acid; restain for 1-12 hr by flooding with a mixture consisting of staining solution, 2 parts, and 1 part of a 1:1 mixture of 2% acetic acid and 2% H₂SO₄ (observe under microscope for staining intensity); wash 2 min in distilled water and 1 hr in tap water. (3) Iron-hematoxylin (Heidenhain): Mordant 6 hr in 2.5% iron-alum solution; wash 1 min in distilled water; stain in 1% or 0.5% ripened hematoxylin for 3-12 br; differentiate 8 min in 2.5%, and 15 min in 1% iron-alum solution; wash 1 hr in tap water. (4) Aceto-carmine (Schneider): Stain 12-24 hr; wash 0.5-1.0 min in distilled water. (5) Picrofuchsin: Stain 24-48 hr in 1% acid fuchsin dissolved in saturated aqueous picric acid; differentiate for only 1-2 sec in 96% ethanol. (6) Modified Giemsa: Mix 640 ml of a solution of 9.08 gm KH₂PO₄ in 1000 ml of distilled water and 360 ml of a solution of 11.88 gm Na₂HPO₄-2H₂O in 1000 ml of distilled water. Soak sections in this buffer, 12 hr. Dissolve 1.0 gm of azur I in 125 ml of boiling distilled water; add 0.5 gm of methylene blue; filter and add hot distilled water until a volume of 250 ml is reached (solution “AM”). Dissolve 1.5 gm of eosin, yellowish, in 250 ml of hot distilled water; filter (solution “E”). Mix 1.5 ml of “AM” in 100 ml of buffer with 3 ml of “E” in 100 ml of buffer. Stain 12-24 hr. Differentiate 3 sec in 25 ml methyl benzoate in 75 ml dioxane; 3 sec in 35 ml methyl benzoate in 65 ml acetone; 3 sec in 30 ml acetone in 70 ml methyl benzoate; and 3 sec in 5 ml acetone in 95 ml methyl benzoate. Dehydrated sections may be covered in a neutral synthetic resin (Caedax was used).     

Staining of fresh, Undecalcified, thin Bone Sections

Fresh, undecalcified bone sections can be reproducibly and reliably stained by any of the following procedures: (A) Basic fuchsin, 1% in 30% alcohol, 48 hr, 22°C. (B) AgNO₃, 0.033 M, 48 hr, 22°C; washing 48 hr in a large volume of distilled water; exposure to light to develop the color. (C) Metallic sulfides (Co⁺⁺, Pb⁺⁺, Hg⁺⁺, Cu⁺⁺): the nitrate of the metal, 0.033 M, 48 hr, 22°C; then Na₂S, 0.033 M, 48 hr, 22° C. (D) Alizarin Red S, 0.1% solution in distilled water, 48 hr, 22°C; differentiated 48 hr at 22°C in weakly alkaline water, pH about 8. (E) KMnO₄: boiling 8-10 min in a 0.1 N, solution. With the exception of D the surface stain must be ground off the section for microscopic examination of its interior. Stain concentration, time and temperature can be altered to suit specific needs.     

Staining Mitochondria With Hematoxylin After Formalin-Sublimate Fixation; A Rapid Method

Mitochondria were stained in liver, kidney, pancreas, adrenal and intestinal mucosa of rat and mouse. Tissues 1 mm thick, were fixed in a mixture of saturated aqueous HgCl₂, 90 ml; formalin (37-38% HCHO), 10 ml, at room temperature (25°C) for 1 hr. Deparaffinized sections 3-4μ thick were treated with Lugol's iodine (U.S.P.) followed by Na₂S₂O₃ (5%), rinsed in water and the ribonucleic acid removed by any of the following procedures: 0.2 M McIlavaine's buffer, pH 7.0, 2 hr, or 0.2 M phosphate buffer, pH 7.0, 2 hr at 37°C; 0.1% aqueous ribonuclease, 2 hr at 37°C; 5% aqueous trichloracetic acid overnight at 37°C; or 1% KOH at room temperature for 1 hr. After washing in water, sections were treated with a saturated solution of ferric ammonium alum at 37°C for 8-12 hr and colored by Regaud's ripened hematoxylin for 18 hr. They were then differentiated in 1% ferric ammonium alum solution while under microscopic observation.     

Permanent Smears of Leaf Tips for the Study of Chromosomes

Young leaf tips are soaked in a saturated aqueous esculin (aesculine) solution at 10-12° C for 15 min to 24 hr and fixed in acetic-alcohol, 1:1. The materials are then stained in a mixture of 2% aceto-orcein and 12V HCl (9:1), 3-4 sec over a flame followed by 30 min or longer at 30° C and then smeared in 1% aceto-orcein. Preparations are made permanent by loosening the cover glass in tertiary butyl alcohol and mounting directly in Canada balsam.     

Chromosome Spreading Induced by Vegetable Oils

Seeds soaked in the oil extracted from castor beans (Ricinus communis) for 2 hr were germinated in petri dishes on moist filter papers. Root tips were fixed in acetic alcohol (1:3) at 10-14°C, for 24 hr, washed successively with 70% alcohol (15 min) and water (10 min), hydrolysed in 1 N HCl at 60°C for 15 min and stained in leucobasic fuchsin for 30 min. The stained tip was squashed under a cover glass in a drop of acetocarmine and sealed with paraffin wax. The slides were made permanent by separating the cover glass in a mixture of acetic acid and n-butyl alcohol (1:1), passing through 2 changes of n-butyl alcohol and mounting in balsam. Such a method leads to contraction and spreading of chromosomes, without affecting either the clarity of the constriction regions or the anaphase separation of chromosomes.     

Nile Blue Histochemical Method for Phospholipids

The technic recommended is: Fix 6-12 hr. in 10% formalin containing 1% CaCl₂. Cut frozen sections without embedding or after gelatin or carbowax. Stain 90 min. at 60°C. in saturated aqueous Nile blue sulfate, 500 ml. plus 50 ml. of 0.5% H₂SO₄, boiled 2 hr. before use. Rinse in distilled water, and place in acetone heated to 50°C. Remove the acetone from the source of heat and allow the sections to remain 30 min. Differentiate in 5% acetic acid 30 min., rinse in distilled water, and refine the differentiation in 0.5% HCl for 3 min. Wash in several changes of distilled water and mount in glycerol jelly. Results: phospholipids - blue; everything else - unstained. Counterstaining nuclei with safranin is optional, but if done, it preferably precedes the Nile blue and is then differentiated by the acetic acid. The histochemical principles on which the method is based are as follows: (1) The calcium compounds of phospholipids combine with the oxazine form of Nile blue sulfate and survive subsequent treatment; (2) neutral lipids are dissolved out by acetone; (3) proteins and other interfering substances are destained by the acetic acid and hydrochloric acid baths.     

Staining Tissue of the Central Nervous System with Luxol Fast Blue and Neutral Red

The tissue is fixed in 10% neutral saline formalin for 1 day to 3 wk depending on the size of the block, dehydrated and embedded in paraffin. The sections are stained at 57° C for 2 hr, then at 22° C for 30 min, in a 0.0125% solution of Luxol fast blue in 95% alcohol acidified by 0.1% acetic acid. They are differentiated in a solution consisting of: Li₂CO₃, 5.0 gm; LiOH-H₂O, 0.01 gm; and distilled water, 1 liter at 0-1° C, followed by 70% alcohol, and then treated with 0.2% NaHSO₃. They are soaked 1 min in an acetic acid-sodium acetate buffer 0.1 N, pH 5.6, then stained with 0.03% buffered aqueous neutral red. Sections are washed in distilled water, 1 sec, then treated with the following solution: CuSO₄·5H₂O, 0.5 gm; CrK(SO₄)₂·12H₂O, 0.5 gm; 10% acetic acid, 3 ml; and distilled water, 250 ml. Dehydration, clearing and covering complete the process. Myelin sheaths are stained bright blue; meninges and the adventitia of blood vessels are blue; red blood cells are green. Nissl material is stained brilliant red; axon hillocks, axis cylinders, ependyma, nuclei and some cytoplasm of neuroglia, media and endothelium of blood vessels are pink.     

Staining Procedures for Ultrathin Sections of Tissues Embedded in Polyester Resin

Tissues were fixed for 30 min In cold (0-2° C) 1% OsO₄ (Palade) buffered at pH 7.7, to which 0.1% MgCl₂ was added. Dehydration was in a graded ethanol series (containing 0.5% MgCl₂) at 0-2° C, and terminated with 2 changes of absolute ethanol. Tissues were then transferred by a graded series to anhydrous acetone. Infiltration of the tissue with Vestopal-W (a polyester resin), is gradual with the aid of graded solutions of Vestopal-W in acetone. The infiltrated tissue is encapsulated and initial polymerization is done under ultraviolet light at room temperature for 8-16 hr. This is followed by final hardening at 60° C for 36-48 hr. Sections (0.2-1 μ) were cut, dried on slides, placed in acetone for 1 min and then treated by either of the following staining procedures: (1) Thionin-azure-fuchsin staining: Flood the preparation with 0.2% aqueous thionin and heat to 60-80° C for 3 min; if the preparation begins to dry, add stain. Rinse in distilled water. Flood the slide with 0.2% azure B in phosphate buffer at pH 9. Heat to 60-80° C for 3 min; do not permit the preparation to dry. Rinse in distilled water. Dip the slide in MacCallum's variant of Goodpasture's carbol-fuchsin stain for 1-2 sec. Rinse in distilled water. Check the preparation microscopically for intensity of the fuchsin stain. Repeat dips as may be needed to obtain the desired intensity. Rinse in distilled water. Dehydrate quickly in 95% and absolute alcohol; clear in 2 changes of xylene and cover in Permount or similar synthetic resin. (2) Thionin-azure counterstain for the periodic acid-Schiff reaction: Oxidize the tissue in 0.5% periodic acid for 15 min and transfer to Schiff's leucofuchsin solution for 30 min. Counterstain with 0.5% aqueous thionin for 3 min; wash in distilled water; stain in 0.2% azure B in phosphate buffer at pH 5.5; wash in distilled water; dehydrate; clear and cover as in the first method. For temporary preparations let dry after absolute alcohol and apply a drop of immersion oil directly on the section.     

Staining and Mounting Helminths

The following procedure is recommended: Fix ces-todes and trematodes (while held flat between glass slides) 0.5-2.0 hr. in the following mixture: formalin, 15; acetic acid (gl.), 5; glycerol, 10; 95% ethyl alcohol, 24; distilled H₂O, 46; all proportions by volume. After freeing them from the slides, wash thoroughly in running water and stain immediately thereafter. Stock staining solution: ferric ammonium alum (violet cryst.), 2 g.; distilled H₂O (cold) 100 ml.; after solution, add 2 ml. concentrated H₂SO₄, bring to a boil; add 1 g. coelestin blue B (Nat. Aniline), boil 3-5 min.; cool and add 10 ml. absolute methyl alcohol and 10 ml. glycerol. Dilute 1 vol. with 3 vol. distilled H20 for use. Stain 5-30 min., depending on size of specimens. Wash with 2 changes 0.5 hr. each of distilled H₂O, then 50% isopropyl alcohol 12-16 hr., 50% isopropyl alcohol 2 hr., followed by graded isopropyl alcohol for dehydration. Ether: ethyl alcohol (equal parts), 1 hr., is followed by embedding in celloidin in a sheet just thick enough to cover the specimens. Trim embedded specimens and dehydrate with isopropyl alcohol, 80%, 90% and absolute. Clear in beechwood creosote. Mount in balsam with cover glasses that overlap the edges of the celloidin 1-2 mm. While drying at 37°C, refill edges of mount with fresh balsam as needed. When dry, remove excess balsam and ring the edges with ordinary gloss enamel paint.     

Enhanced Trehalose Accumulation and Desiccation Survival of Entomopathogenic Nematodes Through Cold Preacclimation

Limited storage stability is a major obstacle to further expansion of the use of entomopathogenic nematodes for pest control. Progress has been made that Steinernema carpocapsae can now be stored under partial anhydrobiosis for up to 6 months at 25°C and 10 months at 5°C in a water-dispersible granular (WG) formulation. However, other species have been more difficult to store in the WG formulation due to migration of nematodes out of the granules and sensitivity of some species to desiccation directly at cold temperatures. As acclimation to cold induces trehalose accumulation (a major cryo- and desiccation protectant) in many invertebrates, it was hypothesized that cold preacclimation of entomopathogenic nematodes will enhance their survival in the WG formulation at cold temperatures. This hypothesis was tested using a temperate species Steinernema feltiae , a subtropical species S. carpocapsae , and a tropical species Steinernema riobrave possessing different thermal niche breadths and reproduction temperature optima. Cold acclimation of infective juveniles increased trehalose accumulation in all three species and the amount of trehalose accumulated was both temperature and species dependent. Trehalose content reached at its peak after 6 days at 5°C in S. feltiae (82.28 μg/mg dry weight), after 10 days at 10°C in S. carpocapsae (94.16 μg/mg dry weight) and after 6 days at 15°C in S. riobrave (47.58 μg/mg dry weight). Cold preacclimation at 5°C for 2 days enhanced desiccation survival of S. feltiae in 25% glycerol (osmotic desiccation) at both 5 and 25° and of S. carpocapsae and S. riobrave only at 5°C. Non-cold acclimated S. carpocapsae and S. riobrave were extremely sensitive to desiccation directly at 5°C in 25% glycerol, resulting in over 98% mortality within 6 days, but S. feltiae was more sensitive to desiccation at 25°C than at 5°C. Cold preacclimation increased survival of all the three species in the WG formulation at both 5 and 25°C. The survival of S. riobrave at 5°C in the WG formulation was positively correlated with the length of preacclimation period at 5°C (R 2 = 0.99) and with the amount of trehalose accumulated during cold preacclimation (R 2 = 0.81). These results support the hypothesis that cold preacclimation enhances desiccation survival of entomopathogenic nematodes at cold temperatures and the increased survival correlates well with the increased trehalose accumulation. Results also demonstrate that cold preacclimation can be used as a tool to enhance survival of nematodes in the formulations with reduced water activity.     

Polyoxyethylene Sorbitan Monopalmitate (Tween 40) as a Vehicle for oil Red O Fat Stain

An oil red O fat stain is prepared by dissolving 250 mg of the dye in 100 ml of a 1% Tween 40 solution in 30% alcohol, and incubating the mixture at 60°C for 24 hr. The solution is then filtered at room temperature under vacuum through medium porosity frittedglass. Frozen sections cut from material fixed in CaCl₂-CdCl₂-formalin (1%:1%:10%) are placed in the stain for not less than 4 hr. After washing in the alcoholic-Tween solvent, they are mounted on glass slides from distilled water with Farrants' medium. The resulting preparations appear to be permanent, for in a 2-yr test they have remained free from stain crystalization and the fat particles are still discrete and dark red.     

Freeze-Dehydration for Permanent Mounts after Aceto-Orcein Stain

Fresh smears of mammalian material were stained 5 min (only) by La Cour's (1941) aceto-orcein method at 60°C, briefly rinsed and drained, and then dehydrated in a prechilled equal-parts mixture of absolute alcohol and anhydrous acetone held 2 hr or more at — 20°C or lower by means of dry ice. The preparations were differentiated 30 sec at room temperature in 95% alcohol containing 1% HCl, counterstained (optional) in a 0.01% solution of fast green FCF in absolute alcohol for 10 sec, passed through 2 changes of xylene and mounted in HSR or other synthetic resin.     

Culture And Staining of Pollen Grains of Ricinus communis

Germinating and growing pollen grains (male gametophytes) of Ricinus communis L. in liquid culture is achieved as follows: Pollen is collected over a 10-15 min period from mature anther clusters which have been removed from the male flowers and which have been kept at 25° C and 40-60% relative humidity. Samples weighing between 2.5 and 5.0 mg are brought as quickly as possible into a Desicote treated vial containing 17% sucrose and 30 ppm H₃BO₃ in boiled distilled water. The proportion (w/v) of pollen to culture solution should be 1:100. Shed pollen is kept in a humidity chamber whenever it is not being handled. The air in the culture vial is replaced by O₂ at the pressure of 1 atmosphere plus 5 lb and the sealed vials are shaken gently for 8-10 hr while partially immersed in a waterbath kept at 30° C. The pollen is fixed by the addition to the incubation suspension of an absolute alcohol-lactic acid (4:1) fixing fluid. The proportion used is 36 parts of fixing fluid to 1 part of culture solution. The fixed pollen can be stored in the fixative. Smears are prepared by applying single drops of the constantly agitated suspension of fixed pollen to a microscope slide. After each drop has spread out and dried, an additional drop is added until 10-20 have been applied. The preparations are stained by adding a drop of 1% acetic-orcein and are sealed with fingernail lacquer. The method is well adapted to the following types of studies: pollen germination, physiology of pollen tube growth, morphology of the male gametocyte, and physiology and cytology of the generative cell and nucleus.     

A HYDROXYL-DEPENDENT SILVER METHOD FOR NERVE AXOPLASM

Axoplasm is selectively impregnated by the following steps: (1) fixation in 10% formalin or in 10% formalin with added sucrose, 15%, and concentrated NH₄OH, 1%, for 1-7 days; (2) frozen sections; (3) extraction of the sections in 95% ethyl alcohol, absolute alcohol, xylene, and 95% ethyl alcohol and absolute alcohol, 1 hr each; (4) distilled water, 3 changes of 10 min each; (5) 20% AgNO₃ (aq.) at 25°C, 30 min; (6) distilled water, 3 changes of 1-2 sec each; (7) 6.9% K₂CO₃, 1 hr; (8) water, 3 changes of about 1 min each; (9) 0.2%AuCl₃, 2 min; (10) distilled water; (11) 5% Na₂S₂O₃, 2 min; (12) washing, clearing and mounting. This procedure is proposed as a simplified stain for axoplasm, with other tissue components remaining unstained. The few reagents necessary suit this method for histochemical investigation of the mechanism of silver staining.     

Polyester Resin Embedding for Sectioning of Hard and Soft Tissues

Soft and calcareous tissues embedded in polyester resin may be cut on a sledge microtome to produce thin sections of 3-4 β thickness. Fixed tissues, dehydrated in ethyl alcohol, cleared in methyl benzoate and chloroform, are taken into a wide-necked bottle containing equal parts of polyester resin and chloroform with 0.75% catalyst. The bottle kept in water bath at 37°C is connected to a vacuum pump. With the evaporation of the chloroform under reduced pressure (approximately 10 mm Hg) infiltration is complete. Tissues transferred into a blocking form containing pure polyester resin with 1.5% catalyst are polymerized at 37° C until blocks are firm (48 hr or more). Blocks are prepared with at least 5 mm margin of plastic surrounding the tissue. The edge of the block adjacent to the knife is then filed at an angle of 45° to the cutting movement. Sections are cut with a wide-backed biplanar knife having a cutting edge of 40-44° positioned at an angle of 30° to the plastic block. As the resin is permeable to most stains, staining is carried out through the plastic Sections carried through staining procedures in wire baskets are floated onto slides and mounted in polystyrene; the cover-glass is compressed with a spring-clamp. Microscopic examination shows no staining of plastic, minimal shrinkage and good cellular detail.     

Histological Localization of Microelectrode Placement in Brain by Ferrocyanide and Silver Staining

After recordings had been taken from a microelectrode used for mapping nerve impulses, a current of 100 μa from the positive pole of a direct current generator was run through the electrode for 5 sec while it was still in place. On terminating the experiment, in which the use of several electrodes was possible, 50-75 ml of a 1:1 mixture of 4% potassium ferrocyanide and 4% acetic acid was injected into each common carotid artery, and the brain left in situ for 0.5 hr. It was then removed and the electrode-bearing part fixed 5-6 hr in a 1:1 mixture of 40% formalin and 95% ethyl alcohol at 55 °C. This specimen was washed in running water 5-10 min, the electrodes removed and frozen sections of 40-80 μ cut and placed in 95% alcohol. Sections were stained 5-10 min at 25-30°C in 10% silver nitrate solution in 75-80% alcohol acidified by 3-4 drops of glacial acetic acid per 50 ml, washed 4-5 sec in each of 2 baths of 95% alcohol, and reduced while being agitated constantly in a 2% solution of pyrogallol and 6-7% formalin in 75-80% alcohol. Washing in 95% alcohol, clearing in clove oil or methyl salicylate followed by xylene and mounting in synthetic resin or balsam completed the process. Sites of electrolysis at the tips of electrodes (under magnification) were blue before silver staining and black after staining. Axons stained brown to black on a yellow background.     

A Celloidin Infiltration-Frozen Section Sequence for Enhanced Preservation of Phosphatases in Bone

The effect of the following embedding procedures on the acid and alkaline phosphatase content of decalcified mouse tibiae has been studied: embedding in 23% gelatine for 18 hr at 37° C, embedding in paraffin wax in vacuo for 1 hr at 58° C, and impregnation with 4% celloidin in diethyl ether and ethanol at 4° C for 2-3 days. Unsupported tissues were also used to demonstrate these enzymes for comparison with the above procedures. Tibiae were first fixed in 10% neutral formalin at 4° C for 15 hr, decalcified in equal volumes of 2% formic acid and 20% sodium citrate at pH 4.9 for not more than 5 days and then washed in distilled water before carrying out the embedding schedules. The celloidin-impregnated tibiae were placed in 70% ethanol to harden the celloidin and then washed in distilled water for 1-2 hr. These tibiae and those embedded in gelatine were cast in a gelatine block which was then hardened in 10% neutral formalin at 4° C for 2 hr. Sections of these and unsupported tibiae were cut at 15 μ on a freezing microtome. Decalcified tibiae embedded and blocked in paraffin wax were sectioned at 15 μ on a base sledge microtome. The enzymes were demonstrated using the coupling azo dye method given by Pearse (Histochemistry, 1st Ed. 1954). The stable diazotates of 4 benzoyl amino 2-5 diethoxyanilene, 3 nitro toluidine and o-dianisidine were used. Of the embedding procedures paraffin wax embedding produced the greatest loss of both enzymes. Gelatine embedding and infiltration with celloidin were equally good for the demonstration of acid phosphatase but for alkaline phosphatase the celloidin method was superior. The gelatine embedded material did not produce consistently good results. Celloidin-impregnated tibiae could be stored without marked deterioration of the enzyme content for longer than gelatine-embedded tibiae.     

Basic Dye Counterstaining of Sections Impregnated by the Golgi-Cox Method

Celloidin blocks of Golgi-Cox impregnated material are cut at 50 μ, the sections collected in 70% alcohol, transferred to a 3:1 mixture of absolute alcohol and chloroform for 2 min, and then stored in xylene or toluene for at least 3 min, or up to 2 wk until processed further. Mounting is done on glass slides which have been coated with fresh egg albumen diluted in 0.2% ammonia water (or a 0.5% solution of dry powdered egg albumen) and then dried at 60°C overnight. For attachment to these coated slides, sections are first soaked for 2-3 min in a freshly prepared mixture of methyl benzoate, 50 ml; benzyl alcohol, 200 ml; chloroform, 150 ml; and then transferred quickly to the slides by means of a brush. After 2-3 min the chloroform evaporates and the celloidin softens. The slides are then immersed in toluene which hardens the celloidin and anchors the sections to the slides. Alcohols of descending concentrations to 40% are followed by alkalinizations, first in: absolute alcohol, 40 ml; strong ammonia water 60 ml, for 2 min, then in: absolute alcohol, 70 ml; strong ammonia water, 30 ml, for 1 hr. Excess alkali is then removed by 70% and 40% alcohol, 2 min each, and a 10 min wash in running tap water. Bleaching in 1% Na₂S₂O₃, for 10 min and washing again in tap water for 10 min completes the process preliminary to staining. The preparations are then stained for 90 min in an aqueous solution of either 0.5% cresylecht violet, neutral red, or Darrow red, buffered at pH 3.6. Dehydration and differentiation in ascending grades of alcohol, clearing with toluene or xylene, and applying a cover glass with a mounting medium having a refractive index of about 1.61 completes the process.