Procedure for Staining Fixed Human Brain Slices

The described method provides a new technique for differentiating areas of gray and white matter in fixed human brain slices. The technique is a modification of an existing method permitting use of the nonfading copper phthalocyanine dye alcian blue Stained slices show turquoise gray matter that contrasts sharply with areas of white matter. Procedure Cut human brains from gross anatomy laboratory cadavers into 4 mm slices and wash in running tap water for 14 hr. Oxidize slices in performic acid for 1.5 hr. Wash in running tap water for 14 hr. Stain slices in shallow dishes in 0.05% aqueous alcian blue. Wash in running tap water for 1 hr. Dry for 2-4 hr and embed in plastic.     

Simple and Durable Staining of Thick Sections of the Human Brain for Macroscopic Study

Observations on gross morphology of the human brain is greatly facilitated by enhancing the contrast between gray and white matter. The proposed technique is much more simple than the generally recommended Mulligan method and its variations. Moreover, there is no loss of stain since the fugitive surface impregnation, obtained by the Mulligan method, is replaced by a thoroughgoing block-staining procedure with the nonfading copper phthalocyanine dye astra blue. Staining procedure: wash formalin-fixed brain slices overnight in running tap water. Place slices in performic acid for 1 hour. Wash in running tap water. Place slices individually in staining solution consisting of 0.1 g astra blue (Merck) in 1000 cc distilled water and 1 cc HCl (37%), for 12-24 hours. Wash in running tap water. Embed in gelatin and mount in plastic cuvettes.     

Simple and durable staining of thick sections of the human brain for macroscopic study.

Observations on gross morphology of the brain is greatly facilitated by enhancing the contrast between gray and white matter. The proposed technique is much more simple than the generally recommended Mulligan method and it variations. Moreover, there is no loss of stain since the fugitive surface impregnation, obtained by the Mulligan method, is replaced by a thoroughgoing block-staining procedure with the nonfading copper phathalocyanine dye astra blue. Staining procedure: wash formalin-fixed brain slices overnight in running tap water. Place slices in performic acid for 1 hour. Wash in running tap water. Place slices individually in staining solution consisting of 0.1 g astra blue (Merck) in 1000 cc distilled water and 1 cc HCl (37%), for 12-24 hours. Wash in running tap water. Embed in gelatin and mount in plastic cuvettes.     

Tannic acid-iron alum reactions: stain of choice for macroscopic sections of brain to be embedded in plastic.

As a macroscopic stain for gross brain sections to be embedded in plastic, tannic acid-iron alum is superior to the generally recommended LeMasurier's variation of the Berlin blue technique because of its greater permanency in plastic. However, as originally adopted for use with brain tissue by Mulligan, the intense black staining of gray matter is too dark for plastic embedded specimens. A modification of this method designed to overcome this difficulty is described. Staining procedure: Wash formalin-fixed brain slices overnight in running water. Wash in distilled water, 2 changes, 30 minutes each. Place slices individually in Mulligan's solution at a temperature of 60-65 C for 4 minutes. Rinse in ice water for 10 seconds. Mordant in 0.4% tannic acid in distilled water for 1 minute. Wash in running tap water for 1 minute. Develop in 0.08% ferric ammonium sulfate in distilled water until gray matter is light gray, about 10-15 seconds. Wash in lukewarm running water for 1 hour, then gently hand-rub whitish film from myelinated surfaces. Store briefly in 3% formalin or 25% glycerine if necessary depending on plastic embedding procedure to be followed.     

Tannic Acid-Iron Alum Reaction: Stain of Choice for Macroscopic Sections of Brain to be Embedded in Plastic

As a macroscopic stain for gross brain sections to be embedded in plastic, tannic acid-iron alum is superior to the generally recommended LeMasurier's variation of the Berlin blue technique because of its greater permanency in plastic. However, as originally adopted for use with brain tissue by Mulligan, the intense black staining of gray matter is too dark for plastic embedded specimens. A modification of this method designed to overmme this difficulty is described. Staining procedure: Wash formalin-fixed brain slices overnight in running water. Wash in distilled water, 2 changes, 30 minutes each. Place slices individually in Mulligan's solution at a temperature of 60-65 C for 4 minutes. Rinse in ice water for 10 seconds. Mordant in 0.4% tannic acid in distilled water for 1 minute. Wash in running tap water for 1 minute. Develop in 0.08% ferric ammonium sulfate in distilled water until gray matter is light gray, about 10-15 seconds. Wash in lukewarm running water for 1 hour, then gently hand-rub whitish film from myelinated surfaces. Store briefly in 3% formalin or 25% glycerine if necessary depending on plastic embedding procedure to be followed.     

A Sensitive Myelin-Sheath Stain Particularly Useful for Small Mammals and Neonatal Cats

Staining of myelinated fibers including the delicate myelin sheaths of infantile animals is as follows: perfuse the anesthetized animal with a pH 7.4 posphate-buffered fixative, either 10% formalin, 6% gluteraldehyde or a mixture containing 3% gluteraldehyde and 2% acrolein. Dissect out the brain or spinal cord and continue fixation for at least 24 hr. Cut larger brains to 1 cm in at least one dimension. Wash in running tap water 2-3 hr and soak in 2.5% potassium dichromate in 1% acetic acid (the primary mordant) for 3-5 days in darkness. Wash at least 12 hr in running tap water. Dehydrate and embed in celloidin and store in 80% ethanol. Section at 25-60 μ into 80% ethanol. Wash 1-2 min in distilled water and then immerse in 1-2% ferric alum at 50 C for at least 1 hr (the secondary mordant). Wash in tap water and stain at least 1 hr at 50-60 C in 0.5% unripened hematoxylin in 1% acetic acid. Wash well in tap water and differentiate in a mixture containing 0.5% ferrityanide, 0.5% borax and 0.5% Na₂CO₃; 2 changes. Wash well in distilled water, then in tap water, and dehydrate, clear and mount. Myelin stains black, cell bodies stain tan, and the background is pale yellow. With minor modifications in timing, the method is applicable to frozen and to paraffin sections; the primary mordant being omitted in the freezing technique.     

Selective gray matter staining of human brain slices: optimized use of cadaver materials

We report a novel staining technique for human brain slices that distinguishes clearly gray from white matter. Previously described techniques using either Prussian blue (Berlin blue) or phthalocyanine dyes usually have included a hot phenol pretreatment to prevent white matter staining. The technique we describe here does not require hot phenol pretreatment and allows the use of brains stored for postmortem periods of one to two years prior to staining. Our technique involves staining with copper(II) phthalocyanine-tetrasulfonic acid tetrasodium salt 1% in water for 2 h followed by acetic acid treatment; this produces excellent blue staining of gray matter with little white matter staining. The stained brain slices are excellent for teaching human brain anatomy and/or pathology, or for research purposes.     

Preparation of Brain Slices for Macroscopic Study by the Copper Sulfate-Phenol-Ferrocyanide Technique

The technique of Mulligan (J. Anat., 65: 468-72, 1931) for preparing gross brain slices has been modified to include improvements in fixation, slicing and staining. The iron alum-tannic acid steps have been eliminated and potassium ferrocyanide substituted. Washing times have been reduced and a detailed routine for fixation and slicing developed. After postmortem removal, human brains were perfused with 40% formalin and kept suspended in 10% formalin for 2-4 wk. Serial slices, 4 mm thick, were cut with an electrically driven rotary-blade meat slicer, washed in running water for 12 hr, immersed in CuSO₄-phenol solution (0.5%: 5%) at 60 C for 6 min, washed again in running water for 5 min, and then treated with a 2% solution of potassium ferrocyanide until a red-brown color developed in the gray matter, usually within 30-60 sec. The slices were then washed again for 5 min and stored in 10% formalin. Brains of patients over 60 yr of age frequently stained poorly and were not used.     

A new method for surface staining large slices of fixed brain using a copper phthalocyanine dye

Here we describe a method for gross staining of gray matter in slices of formaldehyde-fixed human brain. After protection of white matter with 4% phenol at 60°C for 5 min followed by a cold water wash, the gray matter was stained for 10-15 min at 20-25°C with 1% aqueous copper(II) phthalocyanine tetrasulfonic acid tetrasodium salt (CPTS). The staining resisted all attempts to be washed from the gray matter. Stained slices can be stored indefinitely in slightly acidified water, or plastinated as permanent dry specimens.     

A new method for surface staining large slices of fixed brain using a copper phthalocyanine dye.

Here we describe a method for gross staining of gray matter in slices of formaldehyde-fixed human brain. After protection of white matter with 4% phenol at 60 degrees C for 5 min followed by a cold water wash, the gray matter was stained for 10-15 min at 20-25 degrees C with 1% aqueous copper(II) phthalocyanine tetrasulfonic acid tetrasodium salt (CPTS). The staining resisted all attempts to be washed from the gray matter. Stained slices can be stored indefinitely in slightly acidified water, or plastinated as permanent dry specimens.     

A new method for surface staining large slices of fixed brain using a copper phthalocyanine dye

Here we describe a method for gross staining of gray matter in slices of formaldehyde-fixed human brain. After protection of white matter with 4% phenol at 60°C for 5 min followed by a cold water wash, the gray matter was stained for 10-15 min at 20-25°C with 1% aqueous copper(II) phthalocyanine tetrasulfonic acid tetrasodium salt (CPTS). The staining resisted all attempts to be washed from the gray matter. Stained slices can be stored indefinitely in slightly acidified water, or plastinated as permanent dry specimens.     

Protective Paraffin Infiltration of Soft Tissues in Insects to Facilitate Softening of Hard Exoskeletons

This technique can produce serial sections as thin as 5 μ from hard chitin-covered materials of insects or other arthropods. Procedures: Fix with alcoholic Bouin's fluid for 3 hr. Henceforth subject material to partial vacuum in each step to ensure a final proper embedding. Wash with 80% ethanol 2 or 3 times for 2 hr or until the picric acid is largely removed. Dehydrate to 90% ethanol and give 2 changes of n-butanol 2 hr each, and one of a 1:1 n-butanol-paraffin mixture in 56-57° oven for 12 hr. Finally, use 2 baths of pure paraffin, 3 hr each, to complete the infiltration. After the last bath, withdraw the specimen from the paraffin, and remove the superficial paraffin, first mechanically and then with a xylene bath for 4 min. Rinse first with n-butanol, and afterwards with absolute ethanol, 2 min each. The compound eyes are protected with a paraffin covering, the specimen is hydrated with a 1% aqueous solution of detergent for 1 hr and then washed with running tap water. The material is treated with a concentrated sulfuric-nitric mixture (H₂SO₄:HNO₃) for 4 hr to eliminate the exoskeleton. After this treatment, the specimen is washed with running tap water for 12 hr, dehydrated with acetone and then bathed in a 2% solution of celloidin in ethyl acetate to form a protective artificial cuticle. This coating is hardened with 2 quick baths of chloroform, the specimen reembedded in paraffin, and the block cast for sectioning.     

Novel Fixation of Plant Tissue, Staining through Paraffin with Alcian Blue and Hematoxylin, and Improved Slide Preparation

Onion (Allium cepa) root tips were fixed in a proprietary solution without aldehyde, toxic metals or acetic acid. Fixed specimens were embedded in paraffin, sectioned on a rotary microtome and mounted on detergent-washed slides without adhesive. Slides with ribbon segments affixed were immersed in 0.2% aqueous alcian blue 8GX in screw-capped Coplin jars in a water bath at 50 C for 1 hr. Excess alcian blue was rinsed off under cold running tap water and the slides were immersed in quick-mixed hematoxylin at room temperature for 15 min. Stained slides were deparaffinized, rinsed with isopropanol, air dried, and coverslips were affixed with resin. Thus, the traditional paraffin microtechnique has been modified at all steps from fixation to finishing slides with coverslips.     

Selective staining methods for cartilage of rat fetal specimens previously treated with alizarin red S.

A convenient method for staining cartilage with several basic stains after alizarin red S staining of bone was investigated in rat fetuses. It was found that bromophenol blue was useful and effective for staining of the margin and center areas of cartilage, even in specimens stored in glycerin for over 10 years. The specimens were washed in running tap water for 1 hr, and subsequently were immersed in water or in 70% ethanol at pH 4 for 1 hr or longer. The specimens were then stained with 0.005% bromophenol blue in 40% ethanol adjusted to pH 4 for 2 hr, or with 0.001% bromophenol blue in 40% ethanol adjusted to pH 4 for 24 hr. Furthermore, the bromophenol blue stain color actually faded when the specimens were immersed in water or in 70% ethanol at pH 8. Descending order of the stain-effective action on fetal rat cartilage for the basic stains tested was bromophenol blue, aniline blue, Evans blue, methyl violet, trypan blue, and water blue.     

Tannic acid, Iron hematoxylin, Alcian blue, and Basic fuchsin for staining islets and reticular fibers of the pancreas

In this technique alpha cells are stained by basic fuchsin, beta cells by iron-hematoxylin, reticular fibers by ferric tannate, and much by alcian blue. Among 6 commonly used fixatives tested, Bouin's fluid fixation (8-12 hr) gave the best staining results. Procedure: paraffin sections to water; 0.5% Li₂CO₃ to remove picric acid; 20% tannic acid, 15 min; wash well; 2-4 sec in 0.5% basic fuchsin containing 10% alcohol; rinse, then differentiate in 1% aniline in 90% alcohol until alpha cells are red and beta cells pink; 1% phosphomolybdic acid, 1 min; 5% hematoxylin in 2% iron alum, 0.5 min; wash well; 1% filtered alcian blue SGX, 15 sec; rinse, dehydrate, clear, and mount in synthtic resin. Results: reticular fibers, black; acinar cells, orange to gray; alpha cells, red; collagenous fibers, red; beta cells, gray granules; ducts, bluish-green. The method was tested on rat, rabbit, dog, hamster, cow and man.     

Silver Staining, Featuring Rapid Reduction, For Whole Mounts of Retina and Optic Pathways in Chick Embryos

Developing and established nerve fibers in the retina and in superficial tracts of the brain can be stained and viewed en bloc. The method was developed on chick embryos of 2 days of incubation to several months post-hatching but could be used on other material provided that the objects of interest were within 35 μ of the surface. Procedure: (1) Place the entire eye or head in 50% pyridine for at least 16 hr. (2) Wash well for 5-7 hr with hourly changes of distilled water or with running tap water for 4-6 hr followed by several changes of distilled water. (3) Transfer to 95% ethanol for 16-48 hr. (4) Impregnate with 1.5% AgNO₃ for 2 days at 37 C. (5) Submerge in water and, when staining the retina, remove the vitreous body and apply an aqueous solution of 0.25% pyrogallic acid in 1.25% formalin by directing a narrow stream of this reducer against the retina for 2-5 sec. Wash the eye with distilled water 30-60 sec after applying the reducer. When staining the brain, remove the meninges under water, direct the stream of reducer against the brain for 20-30 sec, and rinse the brain immediately after the nerves have stained. (6) Dissect the specimen and make temporary mounts in glycerol; or, dehydrate and clear for resin mounting. The technique stains both mature and growing axons with their growth cones and sometimes their cell bodies. The fiber patterns show best on the surfaces of the retina and brain. The stain works consistently and is suited to the study of both normal and abnormal development.     

The Demonstration of Beta Cells in Pancreatic Islets and Their Tumors

The stain is applied routinely to tissues fixed in 10% buffered formalin (pH near 7.0) or in Bouin's fluid. Bring paraffin section to water as usual and mordant 72 hr in 5% CrCl₃ dissolved in 5% acetic acid. Wash in water and in 70% alcohol and stain 6 hr. Formula of staining solution: new fuchsin, 1% in 70% alcohol, 100 ml; HCl, conc., 2 ml and paraldehyde, 2 ml, mixed together and added to the dye solution; let stand 24 hr before use. After staining, wash in running tap water 5-10 min, rinse in distilled water and counterstain if desired. Dehydration in alcohol, clearing and covering completes the process. When the paraldehyde is obtained from a freshly opened bottle, standardized staining times can be used and thus eliminate the necessity of differentiating individual slides. The granules of beta cells stained deep blue to purple and were demonstrated in the pancreatic islet of man, dog, mouse, frog, guinea pig and rabbit.     

A batch staining method for brain slices allowing volume measurements of grey and white matter using an image analyzing computer (Quantimet 720)

Normal formalin-fixed gelatin-embedded cerebral hemispheres were serially sliced and the 25 to 30 slices form each hemisphere were batch stained by a modification of Mulligan's method. following washing in water the slices were immersed in Mulligan's acid/copper sulfate/phenol solution for 20 minutes at room temperature, treated with a xylene/Polyclens mixture for 20 seconds and immediately transferred to a 2% sodium hydroxide solution for 10 seconds. Final staining was by immersion in 2% potassium ferrocyanide which was followed by washing in tap water. The grey matter was stained a brick red color while the whiteness of the white matter was accentuated. Following staining the slices were stored between sheets of black paper in 2% aqueous formalin prior to measurement of the respective areas of grey and white matter using a Quantimet 720 image analyzing computer. The method is rapid and color stable, and reduces the risk of exposure to toxic fumes by eliminating the need for hot phenol solutions. This technique is also suitable for the macroscopic demonstration and quantitation of demyelinating conditions in the brain.     

A Batch Staining Method for Brain Slices Allowing Volume Measurements of Grey and White Matter Using an Image Analyzing Computer (Quanttmet 720)

Normal formalin-fixed gelatin-embedded cerebral hemispheres were serially sliced and the 25 to 30 slices from each hemisphere were batch stained by a modification of Mulligan's method. Following washing in water the slices were immersed in Mulligan's acid/copper sulfate/ phenol solution for 20 minutes at room temperature, treated with a xylene/Polyclens mixture for 20 seconds and immediately transferred to a 2% sodium hydroxide solution for 10 seconds. Final staining was by immersion in 2% potassium ferrocyanide which was followed by washing in tap water. The grey matter was stained a brick red color while the whiteness of the white matter was accentuated. Following staining the slices were stored between sheets of black paper in 2% aqueous formalin prior to measurement of the respective areas of grey and white matter using a Quantimet 720 image analyzing computer. The method is rapid and color stable, and reduces the risk of exposure to toxic fumes by eliminating the need for hot phenol solutions. This technique is also suitable for the macroscopic demonstration and quantitation of demyelinating conditions in the brain.     

Comparison of methylprednisolone with dexamethasone in treatment of acute spinal injury in rats

Effect of methylprednisolone sodium succinate (MPSS) and its comparison with dexamethasone in experimentally induced acute spinal cord compression in adult rats was studied. The rats were divided into group A (control) and group B, which was subdivided into B1, B2, B3 where MPSS was given after 1, 8 and 24 hr and B4 where dexamethasone was given after 1 hr of cord injury respectively. Proper neurological evaluation was done with mobility, running and climbing score. Recovery index was evaluated for 7 days. After sacrificing the rats, spinal cord was observed histopathologically. Mean recovery index and microscopic findings based on hemorrhage in gray and white matter, neuronal degeneration, hematomyelia and edema in white matter were recorded. The results suggested that MPSS was effective in promoting post-traumatic clinical and histological recovery and to a greater extent, when given 1 hr after trauma. MPSS is more effective than dexamethasone in reducing edema when both are given after interval of 1 hr.