Silver Protein Staining of Nerve Fibers in Celloidin Sections

Celloidin sections from formalin-fixed brain and spinal cord of primates are stored in 70% alcohol after cutting, soaked in 2% pyridine in 50% alcohol for 6-8 hr at 37 C, and transferred to 1% concentrated NH₄OH in 50% alcohol 15-18 hr at 20-25 C. After washing and flattening, the sections are transferred to 1% silver protein solution containing 30 ml of 0.2 M H₃BO₃/100 ml. Impregnation is accomplished in 50 ml screw-top jars, 50 mm in diameter, which are filled to a depth of 35 mm, and have 1 gm of copper foil, 0.002 inch thick added. The foil is folded in loose accordion-fashion, pierced and threaded, cleaned in 5% HNO₃, rinsed in distilled water, and suspended in the solution just above the sections by fastening the thread to the jar lid. The sections are impregnated for 24 hr at 37 C, rinsed in distilled water, reduced in a solution of 5% Na₂SO₃ and 1% hydroquinone for 10 min, washed in distilled water and toned in 0.2% gold chloride for 5 min. After rinsing in distilled water, the sections are transferred to 1% oxalic acid for 45-60 sec, washed in distilled water and placed in 5% Na₂S₂O₃ for 5 min. Sections are then washed, dehydrated to 95% alcohol, cleared in terpineol, followed by 3 changes in xylene, and mounted.     

Staining Paraffin Sections with Protargol 6. Impregnation and Differentiation of Nerve Fibers in Adrenal Glands of Mammals

A series of experiments with protargol staining of nerve fibers in mammalian adrenal glands has yielded the following procedure: Fix-1-2 days in a mixture of formamide (Eastman Kodak Company) 10 cc, chloral hydrate 5 g., and 50% ethyl alcohol 90 cc. Wash, dehydrate and embed in paraffin. Cut sections about 15 and mount on slides. Remove the paraffin and run down to distilled water. Mordant 1-2 days in a 1% aqueous solution of thallous (or lead) nitrate at 56-60°C. Wash thru several changes of distilled water and impregnate in 1% aqueous protargol (Winthrop Chemical Company) at 37-40°C. for 1 to 2 days. Rinse quickly in distilled water and differentiate 7-15 seconds in a 0.1% aqueous solution of oxalic acid. Rinse thru several changes of distilled water for a total time of 0.5 to 1.0 rain. Reduce 3-5 rain, in Bodian's reducer: hydroquinone 1 g., sodium sulfite 5 g., distilled water 100 cc. Wash in running water 3-5 min. and tone 5-10 min. in a 0.2% gold chloride solution. Wash 0.5 min. or more and reduce in a 2% oxalic acid solution to which has been added strong formalin, 1 cc. per 100. (Caution. This last reduction is critical and over-reduction can spoil an otherwise good stain; 15-30 seconds usually suffices, and the sections should show only the beginning of darkening to a purplish or gray color.) Wash, fix in hypo, wash, dehydrate and cover.     

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.     

The Stainability of Nerve Fibers by Protargol With Various Fixatives And Staining Technics

The influence of the commonly used tissue fixing reagents, individually and in various combinations, on subsequent staining by protargol was studied. The reagents used were formalin, formamide, picric acid, acetic acid, paranitrophenol, pyridine and chloral hydrate. Parraffin sections from intestine and peripheral nerve of cat, dog, monkey and rat were stained with protargol after fixation in various experimental mixtures of the fixing reagents. Satisfactory nerve stains of intestine were not obtained with regularity after any one fixing and staining procedure. (Good fixation and staining appeared to be influenced by properties inherent in the tissue itself and showed marked variations from animal to animal even in the same species.)Stains of nerve fibers in peripheral nerve trunks were much more easily obtained than in the intestine where good stains were sporadic and unpredictable. The use of a mixture of 0.5% protargol and 0.1% fast green FCF, is proposed as a silver-dye staining medium.     

Staining Paraffin Sections with Protargol 3. The Optimum pH for Reduction. 4. A Two-Hour Staining Method

Further work on conditions affecting the reduction of paraffin sections impregnated with protargol showed that the optimum pH for sulfite-amidol mixtures was between 6.5 and 7.5. A staining method which requires about two hours to complete consists of the following steps: (1) One hour impregnation at 60° C. in 10% AgNO₃. (2) Wash in distilled water 3 changes of 30 sec. each. (3) Put into protargol (Winthrop Chem. Co., New York, N. Y.) 0.2% aq. for another hour at room temperature. (4) Rinse 2 sec. (5) Reduce one to two min. in amidol 0.2 g., Na₂SO₃ 8 g., NaHSO₃ I g., and water 100 cc. (6) Wash thoroly. (7) Tone with 0.1% gold chloride. (8) Wash. (9) Reduce with a 0.5% aq. soln. of amidol (no sulfite). (10) Wash, dehydrate and cover. The method stains neurofibrillae and unmyelinated fibers and has worked well on most tissues of vertebrates. The stain follows acid alcoholic fixation.     

Fluorescent Staining of Juxtaglomerular Cells in Frozen Sections

Enzymatic investigations of the juxtaglomerular apparatus often creates the need for visualisation of granulated juxtaglomerular cells (JGC) in preparations subjected to histochemical procedures. In our investigations, Pitcock and Hartroft's (1958) modification of Bowie's method and the Endes et al. (1969) combined trichrome staining proved to be inadequate when applied to fresh cryostat sections, or to formol- or glutaraldehyde-fixetl, gum sucrose-impregnated frozen sections. Friedberg and Reid's (1966) crystal violet procedure for waxembedded kidneys also failed to give uniformly reproducible results. In attempting to find a satisfactory technique for both enzyme and granule staining, we noted Janigan's (1965) and Haratla's (1969) observations on paraffin-embedded JGC, and tested the following fluorochromes: thioflavine T—Fluka, C. I. 49005; auramine O—Merck, C. I. 41000; acridine orange—E. Gurr, C. I. 46005; berberine sulfate—Fluka, C. I. 75160 on 10 μ sections of albino mouse kidneys prepared in 4 different ways as follows:     

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 Rapid lmmunostaining Method for Frozen Sections

A simple and rapid one-step method for demonstrating immunohistochemical markers (leukocyte common antigen, cytokeratin, etc.) is described, which can help define the nature of poorly differentiated neoplasms for diagnosis using frozen section. Microwave irradiation was used to speed immunohistochemical analysis using “Enhanced Polymer One-step Staining” (EPOS) reagents on cryostat sections from a variety of pathologic samples. Reproducible results were obtained using EPOS reagents for leukocyte common antigen and cytokeratin. The overall procedure takes less than 10 min and can be completed during surgery.     

Silver Staining of Nerve Fibers in Cardiac Tissue

Fresh hearts of dog were perfused through the coronary vessels with 1000 ml. of fixative (chloral hydrate, 5 g. per 100 ml. of 70% ethyl alcohol) and blocks of tissue 2 × 5 mm. from epicardium to endocardium fixed 48 hours in the same fixative. The blocks were placed in 95% alcohol containing 0.3% addition of strong ammonia for 4 hours, followed by 2 changes of plain 95% alcohol of 1 hour each, then cleared and infiltrated with paraffin. Mounted sections 12-15 µ thick were incubated in 1% silver proteinate (obtained from Serumvertrieb, Marburg, Germany)² at 38° C. for 48 hours in the presence of 10 g. of 15 gauge copper wire per 200 ml. of solution. The slides were rinsed gently in 3 changes of distilled water for 2 minutes, 1 minute and 1 minute, respectively, and reduced in 1% hydroquinone and 5% sodium sulfite for 5 minutes. They were washed 5 minutes in tap water and 5 minutes in 2 changes of distilled water and toned 3-5 minutes in 0.25% gold chloride, rinsed in distilled water 10 seconds, reduced 10 seconds in 1 % oxalic acid, rinsed 1 minute, fixed in 5% sodium thiosulfate 5 minutes, washed in tap water through 3 changes, dehydrated, cleared and covered. All solutions were made with distilled water except where otherwise specified. The results gave good impregnation of fine nerve fibers without the usual confusing staining of reticular tissue.     

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.     

An Efficient Staining Method for Ultrathin Sections

A staining method to handle simultaneously as many as 20 electron microscope grids is described. The devices used are easily constructed of readily obtained inexpensive materials. The volumes of stain and wash water required are very small and drying grids is simplified.     

Staining Pericellular Boutons of Glutaraldehyde-Fixed Nervous Tissue; A Chromation-Silvering Sequence for Frozen Sections

After fixing in phosphate-buffered 5% glutaraldehyde, pH 6.8, by perfusion, brains were sliced to 3-5 mm pieces which were placed in the fixative for 5-7 days. The pieces were washed through several changes of 2.26% NaH₂PO₄ for 12 hr, 30 μ frozen sections cut, and mordanted 2 days in an equal-parts mixture of 3.5% CrO₃ and 5% Na-tartrate, which had been aged at 20-25 C for 20 days prior to use. After washing in distilled water, the sections were put into a solution containing AgNO₃, 20 gm; and KNO₃, 15 gm, in distilled water, 80 ml; at 30 C for 1.5-2 hr, then reduced at 40-45 C in three pyrogallol solutions as follows: 1-2 sec in 1% pyrogallol in 55% alcohol; 3-4 sec in a 0.67% solution in 33% alcohol, and 5-7 sec in a 0.5% solution in 25% alcohol. Gold toning is optional; dehydration, clearing and covering, routine. The technic shows particularly the perisomatic fibers, boutons en passant and boutons termineaux. Fibers in nerve tracts may be visible but lightly stained; cell nuclei may be dark, but the cytoplasm remains pale.     

Staining of Nerve Fibers with Methylene Blue Factors Improving the Staining

Several factors influencing the staining of nerve fibers with methylene blue, especially the influence of chloralhydrate and carbamylcholine chloride (as parasympathicotonics), and of some anesthetics were studied. The intestines of mouse, rat, and guinea pig were used. The following immersion technic is suggested: Tissue from animals anesthetised by chloralhydrate is immersed in: zinc free methylene blue, 0.03%; sodium tartrate, 0.5%; sodium pyruvate, 0.05% carbamylcholine, 0.00005%; 0.2 M Na₂HPO₄, 0.77%; 0.1 M citric acid, 0.18%; NACl, 0.79%; also an anesthetic which varies with the animal selected. Air is kept bubbling through the staining solution and microscopic examination is made at 6 min. intervals. After 0.5-1 hr. the tissue is fixed in: ammonium molyb-date, 10 g.; sucrose, 35 g.; distilled water, 100 ml.; to which is added just before use, 1% platinum chloride, 3 ml.; 2% osmic acid, 3 drops. Washing is in ice cold water and dehydration at 0°C. in Lang's fluids (varying mixtures of ethanol and n-butanol). The tissues thus prepared are stored in liquid paraffin.     

A Differential Staining Method for Elastic Fibers, Collagenic Fibers, and Keratin

A method allowing for the differential presentation of elastic fibers, other connective tissue fibers, epithelial and other types of cytoplasm, and keratin is described. The procedure is based on the affinity of orcein for elastic fibers, of anilin blue for collagenic material, and of orange G for keratin. Bouin-fixed, tissue-mat embedded sections are stained in Pinkus' acid orcein for 1 1/2 hours and rinsed in distilled water. The sections are differentiated in 50% alcohol containing 1% hydrochloric acid, washed in tap and then in distilled water. The sections are next transferred for I to 2 minutes to the anilin blue, orange G, phosphomolybdic acid combination known as solution No. 2 of Mallory's connective tissue stain, diluted 1:1 with distilled water. They are then rinsed in distilled water, quickly passed into 95% alcohol, and dehydrated in absolute alcohol containing some orange G, after which they are cleared and mounted. Within less than two hours sections may be stained and mounted with the following results: elastic fibers — red; collagenic fibers — blue; muscle fibers — yellow; keratin — orange.     

A Urea Silver Nitrate Method for Nerve Fibers and Nerve Endings

A silver nitrate stain for nerve fibers and endings applicable to paraffin sections on the slide utilizes the properties of urea to accelerate the procedure and improve the specificity of the stain. After removal of the paraffin the sections are run through absolute, 95% and 80% alcohol and placed for 60-90 minutes at 50-60°C. in: 1% aqueous silver nitrate, 100 ml.; urea, 20-30 g.; 1g. mercuric cyanide and 1 g. picric acid in 100 ml. of distilled water, 1-3 drops. After the silver bath they are rinsed quickly in 2 changes of distilled water and reduced for 3-5 minutes at 25-30°C. in: water, 100 ml.; sodium sulfite, anhydrous, 10g.; hydroquinone, 1-2g.; urea, 20-30g. They are then washed thoroughly in 4-5 changes of distilled water, passed through graded alcohols into 80% alcohol and examined under the microscope. If nerve fibers are not distinct, the sections are returned to the same urea-silver-nitrate bath for 10-15 minutes, rinsed, reduced, washed and dehydrated as before. This process may be repeated until staining is adequate; then they are dehydrated, cleared, and mounted.

Nerve fibers show a color range from brown to black; nerve cells from yellow to brown; and the background, depending on the type of tissue and its fixation, from yellow to light brown.