Notes on Technic: Mahon's Myelin Stain for Celloidin-Embedded Nervous Tissue

Two methods commonly used to stain myelin sheaths are Kluver and Barrera's luxol fast blue (Kluver and Barrera 1953) and Weil's iron hematoxylin (Weil 1928). Both require differentiation of the stain; in addition, the Kluver-Barrera method specifies 16-24 hour staining. A third method for the selective staining of myelinated axons is that of Mahon (1938), which was introduced for use with paraffin-embedded autopsy tissue. The procedure possesses two distinct advantages since it requires: (1) no differentiation of the stain and (2) only 1 hour staining. Loyez's (1910) myelin stain for celloidin embedded tissue is similar to Mahon's but calls for long staining followed by differentiation. This report describes the application of Mahon's method to celloidin-embedded experimental tissue and emphasizes its utility for staining tissues to be used for reconstructing microelectrode penetrations (fig. 1) and for demonstrating the effect of experimental lesions (fig. 2).     

An Evaluation of Cresyl Echt Violet Acetate as a Nissl Stain

The new cresyl echt violet acetate, of which three different batches have been tested, proves to be a very useful Nissl stain. It is especially valuable for formalin-fixed, frozen-sectioned material. By using a buffered staining bath and controlled timing in dehydration it is possible, on paraffin embedded material, to use these dyes as progressive stains apparently specific for nucleic acids. With a saturated aqueous solution of the dye, especially when a mordant of lithium carbonate is used, it is possible to stain material that has been preserved in formalin for several years and also material from which nucleic acids have been removed. The dye is useful also for staining celloidin embedded material. With the buffered stain proposed, differentiation is much easier than with older methods which included a gross overstaining and a long destaining procedure.     

A Teflon Disk for Processing Loose Serial Celloidin Sections

Celloidin sections are routinely used for Nissl, Golgi, or Golgi-Cox staining (e.g., Glaser and Van der Loos 1981) when sections thicker than 30 μm are required. In spite of the advantages of the celloidin method (see Voogd and Feirabend 1981, Buschke 1979), processing free-floating serial sections of celloidin embedded material, which may often be preferred, is not very convenient.     

Nongraded Dehydration and Low-Pressure Infiltration for Rapid Celloidin Embedding of Brain Tissue

Various ways of shortening single steps in the celloidin process have been combined to form a routine method which may be completed, for tissues of average size, within a week following fixation. Fixed, washed tissue slices 5 mm thick are dehydrated in 1 or 2 changes of absolute ethanol and acetone, 1:1. This requires 24 hr in an incubator at 37 C, or 12-16 hr if a magnetic stirrer is used. After ether-alcohol for 4 hr. the tissues are transferred to 5% celloidin and infiltrated in a vacuum desiccator attached to a filter pump. When the volume of celloidin is reduced to half the original amount (about 2 hr), the tissues are removed from the infiltrating fluid and embedded in 10% celloidin. The blocks are hardened in chloroform and cleared by suspending them in 2 or 3 changes of terpineol agitated by a magnetic stirrer. Sections are cut in terpineol, using any type of microtome. After washing in 95% alcohol, they are mounted on albumenized slides for staining.     

Rapid Nissl Staining for Frozen Sections of Fresh Brain

Working with X-ray film autoradiography of soluble isotopes, we needed a staining technique for the localization of nuclei in frozen sections of fresh brain. We have found no Nissl staining method in the literature concerning autoradiography specially recommended for this purpose, nor have we found in handbooks on staining a Nissl method clearly recommended for unfixed, frozen sections of brain. The methods described are intended for paraffin or celloidin sections, and require fixation of brain before sectioning (which must be avoided when working with soluble isotopes). Because autoradiography is a time-consuming method, any technique which shortens time needed for the overall procedure is welcome. Most Nissl techniques described in the literature require long preprocessing of the tissue. We found two rapid methods, described by Humason (1967) and LaBossiere and Glickstein (1976), but their application to frozen sections did not give good results. After trials with several types of techniques, we succeeded in developing two Nissl modifications with slightly different qualities, one of 12 min and the other of 2-3 h. The longer method includes conventional steps in staining; the shorter method does not include fixation or lipid extraction. These methods were applied to 20-60 μm brain sections cut in the cryostat at -10 to -12 C and dried on gelatinized slides.     

A new technique for staining sections and recovering celloidin film in high resolution autoradiography

Ultrathin sections are stained immediately after cutting by placing them in contact with staining solution and then placed on a slide covered by a celloidin film. This method largely avoids precipitates of heavy metals. The recovering of celloidin film is improved using a stainless steel basket. This technique is far more reliable than that involving use of a filter paper.     

Notes on Technic: A Device for Centering Tissue Blocks on the Porter-Blum Microtome

A method of double embedding fixed tissues in 3% low viscosity nitrocellulose and paraffin is described. Five percent phenol in 80% alcohol during dehydration and 5% glycerin in the nitrocellulose solutions enhance cutting qualities. A modified Ruyter's solution is used to flatten sections. After a section is aflixed to a slide, it is passed through chloroform and acetone to remove the paraftin and celloidin. A 1% celloidin dip insures adherence of the seaion to the slide. Slides are stored in 70% alcohol until they are to be stained. Following staining and dehydration in graded alcohols, clearing should be done in a 1: 3 mixture of terpineol and toluene.     

Double Embedding Again

A method of double embedding fixed tissues in 3% low viscosity nitrocellulose and paraffin is described. Five percent phenol in 80% alcohol during dehydration and 5% glycerin in the nitrocellulose solutions enhance cutting qualities. A modified Ruyter's solution is used to flatten sections. After a section is aflixed to a slide, it is passed through chloroform and acetone to remove the paraftin and celloidin. A 1% celloidin dip insures adherence of the seaion to the slide. Slides are stored in 70% alcohol until they are to be stained. Following staining and dehydration in graded alcohols, clearing should be done in a 1: 3 mixture of terpineol and toluene.     

A Rapid Celloidin Method for the Rotary Microtome

A method is described which combines the writer's hot celloidin technic¹ with a form of the clearing-before-cutting procedure. The method requires only 16-17 days and yields a block which may be cut in any microtome, the sections being as thin as those afforded by paraffin with comparable material. The advantages of celloidin over paraffin, listed in the writer's earlier paper, are retained in the present method which, altho consuming more time than the hot process, requires less skill and gives superior results.     

An Improved Fixing Solution for Methylene Blue Preparations

A method is described which combines the writer's hot celloidin technic¹ with a form of the clearing-before-cutting procedure. The method requires only 16–17 days and yields a block which may be cut in any microtome, the sections being as thin as those afforded by paraffin with comparable material. The advantages of celloidin over paraffin, listed in the writer's earlier paper, are retained in the present method which, altho consuming more time than the hot process, requires less skill and gives superior results.     

Demonstration of cathepsin B in the rat kidney using fluorescence histochemistry

The procedure of mounting freeze-dried sections with celloidin was adapted for the fluorescent-histochemical demonstration of cathepsin B in the rat kidney. A good localization of reaction products was shown in freeze-dried, 5-micron sections which had been mounted free floating with 1.5% celloidin solution on albuminized slides. Using this procedure, the reaction products were localized in the lysosomes, particularly those of the convoluted proximal tubule.     

Application of the Periodic Acid-Schiff Technique to Whole Chick Embryos

The practicability of applying histochemical reactions to bulk staining has been explored by subjecting whole chick embryos at early stages to the periodic acid-Schiff (PAS) reaction. A comparison of the microscopic distribution of PAS positive substances revealed by this procedure with that obtained by the standard routine, i.e., staining of deparaffinized sections on slides, has shown similar localizations of PAS positive material and, in addition, finer morphological detail and more intensive reactions by staining the specimens in toto. The following method is recommended for chick embryos between stages 11-17 (Hamburger and Hamilton): Fixation in Gendre's fluid at 4°C; oxidation with alcoholic buffered periodic acid, 15 min; rinsing in distilled water, 10 min; Schift's reagent, 30 min; 3 sulfite rinses, 5 min each; running tap water, 10 min; dehydration, clearing and double-embedding in celloidin and paraffin.     

Quick Staining Method For Frozen Sections

Since the advent and general acceptance of frozen sections in histological and pathological laboratories it has been necessary to devise methods for staining these sections. The usual method is fixing the tissue to a slide by the use of celloidin. This paper is an attempt to describe a permanent, quick method of staining frozen sections without distortion or mechanical tearing of the tissues.     

Factors which Influence the Latent Image in Autoradiography

Sections treated with N-ethyl-maleim'ide, a sulfhydryl blocking reagent, exhibit similar grain counts over nonradioactive and S-labeled tissues. This reveals that no chemo-graphic effects were produced by sulfhydryl groups in tissues fixed and prepared in the manner described. It was noted, however, that background count over tissue sections is consistently tower than over adjacent him, indicating perhaps the existence of substances in tissues which interfere with the production of the latent image. These effects have been eliminated by coating the tissues with celloidin. Celloidin coating can be used for most emitters including S but not for tritium. A special method for tritium, not involving celloidin coating is given.     

Factors which Influence the Latent Image in Autoradiography

Sections treated with N-ethyl-maleim'ide, a sulfhydryl blocking reagent, exhibit similar grain counts over nonradioactive and S-labeled tissues. This reveals that no chemo-graphic effects were produced by sulfhydryl groups in tissues fixed and prepared in the manner described. It was noted, however, that background count over tissue sections is consistently tower than over adjacent him, indicating perhaps the existence of substances in tissues which interfere with the production of the latent image. These effects have been eliminated by coating the tissues with celloidin. Celloidin coating can be used for most emitters including S but not for tritium. A special method for tritium, not involving celloidin coating is given.     

Streptococcal Desoxyribonuclease for the Removal of Feulgen-Stainable Material

Removal of Feulgen-stainable material from the cell nucleus was accomplished by treatment of sections with streptococcal desoxyribonuclease. The procedure recommended is (1) Deparaffinize with xylene, followed by descending grades of alcohol. (2) Wash in tap water. (3) Treat slides for 1 hour at 37°C. with streptococcal desoxyribonuclease (1000 units/ml.) in 0.025M veronal buffer of pH 7.5 containing 0.003M MgSO₄. Treat control slides for an equal length of time at the same temperature. Renew the enzyme approximately every 15 minutes. (4) Wash slides briefly in tap or distilled water. (5) Dehydrate, then coat the sections by dipping in a 1% solution of celloidin in alcohol-ether. (6) Subject the preparations to the Feulgen reaction. Control slides showed characteristic nuclear staining; enzyme treated slides did not stain.     

A Universal Stain for the Sex Chromatin Body

For the demonstration of the sex chromatin body in human tissues, fixation in 95% alcohol or modified Davidson's solution (95% alcohol, 30; formalin, 20; glacial acetic acid, 10; distilled water, 30) was best. The staining procedure chosen for most materials is the following: Mounted preparations are coated with celloidin, hydrated, hydrolyzed 20 min in 52V HCl at 20-25°C, rinsed thoroughly in several changes of distilled water and transferred to a buffered thionin solution. This consists of 3 parts: (1) A saturated solution of thionin in 50% alcohol (filtered); (2) Michaelis buffer: sodium acetate (3 H₂O), 9.714 gm; sodium barbiturate, 14.714 gm; CO₂-free distilled water, 500 ml; and (3) 0.1N HCl. To make the staining solution, mix 28.0 ml of the buffer solution with 32.0 ml of 0.1N HCl and bring the total volume to 100.0 ml with the thionin solution. Its pH should be 5.7 × 0.2, and care should be exercised that no acid is carried over from the hydrolyzing solution, since this would progressively lower the pH. The staining time varies from 15 to 60 min, depending on the specimen, but the shortest time consistent with adequate staining gives the clearest preparations. Slides are rinsed in distilled water and 50% alcohol and allowed to remain in 70% alcohol until the heavy clouds of stain cease to appear. Differentiation is completed in 80% and 95% alcohol, followed by dehydration in absolute alcohol, clearing in xylene and applying a cover glass with a synthetic resin (G. T. Gurr's DePeX was used). The sex chromatin is deep blue-violet and sharply contrasted against the lightly colored particulate chromatin of the nucleus. Cytoplasm remains unstained but fibrin and related structures show metachromasia. Chromosomes are well demonstrated if present. The method works on all types of tissues, is simpler and quicker than the Feulgen method, and often yields superior results.     

Deplasticizing of thick Epon sections involving a new adhesive technique and staining of nervous tissue

The present communication deals with a technique developed for the selective staining of neural tissue in thick (10 micron) Epon sections. A new adhesive method was needed, because the known techniques are only applicable to 0.5-2 micron thin sections. The critical step in the procedure is the adhesion of the sections onto the slides. This is accomplished by heating the sections on top of a uniform layer of albumin glycerol on the slide followed by coating with celloidin. The results after deplasticizing and coagulation with this technique are comparable to those obtained by paraffin or frozen section techniques, but in addition have the advantage of Epoxy resin embedding e.g. the possibility of cutting undecalcified hard tissues and sections for serial reconstruction.     

A Rapid Silver-on-the-Slide Method for Nervous Tissue

A progressive silver staining method is described, which permits microscopic examination of the sections during the staining process. After formaldehyde fixation, dehydration and embedding in paraffin or celloidin, fine fibers and synaptic endings may be demonstrated. After formaldehyde fixation and mordanting in 3% K₂Cr₂O₇, myelinated fibers and mitochondria are specifically stained.

The unique feature of this method is, that the silver solution (0.5% protargol) is mixed with the reducing solution: 1.6% Rochelle salts, containing traces of Ag NO₃, MgSO₄, and K₂S (U.S.P.). The sections are placed directly into this mixture, which is then warmed to 45-55° C. Sections are removed when progressive staining is completed, washed in water, dehydrated and mounted.

In the fiber stain, nerve fibers and synaptic endings are dark brown or black, and nuclear chromatin is deep brown, against a pale yellow background. When the myelin sheath procedure is followed, the fiber bundles are deep brown, and the intensity of the staining remains the same for specific tracts, aiding in their identification.     

A Simplified and Convenient Method for the Double-Embedding of Tissues

A method for embedding tissues with a celloidin-paraffin combination is presented. The essential features of the process depend upon (1) a thorough infiltration of the specimen with celloidin of low concentration, and (2) the subsequent impregnation of both the specimen and the celloidin with paraffin.

The methods for sectioning, and the removal of the embedding agent are given.

The chief advantages of this method are: the preservation of all of the advantages of celloidin embedding but with a great saving of time, and greater convenience of storage; the cutting of thin sections (2μ for many types of tissues); it is useful for embedding specimens for which neither pure paraffin nor pure celloidin are entirely satisfactory, i.e. those containing tissues differing in density.