A Protargol Method for Staining Nerve Fibers in Frozen or Celloidin Sections

Extensive experimentation with protargol staining of neurons in celloidin and frozen sections of organs has resulted in the following technic: Fix tissue in 10% aqueous formalin. Cut celloidin sections IS to 25 μ, frozen sections 25 to 40 μ. Place sections for 24 hours in 50% alcohol to which 1% by volume of NH₄OH has been added. Transfer the sections directly into a 1% aqueous solution of protargol, containing 0.2 to 0.3 g. of electrolytic copper foil which has been coated with a 0.5% solution of celloidin, and allow to stand for 6 to 8 hours at 37° C. Caution: In this and the succeeding step the sections must not be allowed to come in contact with the copper. From aqueous protargol, place the sections for 24 to 48 hours at 37° C. directly into a pyridinated solution of alcoholic protargol (1.0% aqueous solution protargol, 50 ml.; 95% alcohol, 50 ml.; pyridine, 0.5 to 2.0 ml.), containing 0.2 to 0.3 g. of coated copper. Rinse briefly in 50% alcohol and reduce 10 min. in an alkaline hydroquinone reducer (H₃BO₃, 1.4 g.; Na₂SO₃, anhydrous, 2.0 g.; hydroquinone, 0.3 g.; distilled water, 85 cc; acetone, 15 ml.). Wash thoroly in water and tone for 10 min. in 0.2% aqueous gold chloride, acidified with acetic acid. Wash in distilled water and reduce for 1 to 3 min. in 2% aqueous oxalic acid. Quickly rinse in distilled water and treat the sections 3 to 5 min. with 5% aqueous Na₂S₂O₃+5H₂O. Wash in water and stain overnight in Einarson's gallocyanin. Wash thoroly in water and place in 5% aqueous phosphotungstic acid for 30 min. From phosphotungstic acid transfer directly to a dilution (stock solution, 20 ml.; distilled water, 30 ml.) of the following stock staining solution: anilin blue, 0.01 g.; fast green FCF, 0.5 g.; orange G, 2.0 g.; distilled water, 92.0 ml.; glacial acetic acid, 8 ml.) and stain for 1 hour. Differentiate with 70% and 95% alcohol; pass the sections thru butyl alcohol and cedar oil; mount.     

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.     

Differential Staining of Inner Ear Fluids by Protargol

Present day techniques for processing temporal bones involve celloidin embedding. With a few modifications in Bodian's silver staining procedure the celloidin of the endolymphatic spaces stains darker than that of the perilymphatic spaces providing there is no break in the anatomical barrier between them. Essentially the routine procedure of Bodian is used except that metallic copper is omitted from the staining solution, impregnation time is reduced to 3 hr, reduction time is extended to 10 min and no oxalic acid is used for gold toning.     

Staining Nerve Fibers After Sublimate-Acetic and After Bouin'S Fluid

A silver staining method for paraffin sections of material fixed in HgCl₂, sat. aq., with 5% acetic acid is as follows. Process the sections through the usual sequence of reagents, and including I-KI in 70% alcohol, thiosulfate (5% aq.), washing and back to 70% alcohol containing 5% of NH₄OH (conc. aq.). After 3 minutes in the ammoniated alcohol, wash through tap water and 2 changes of distilled water and silver 5-10 minutes at 25°C. in 15% AgNO₃ aq. to which 0.02 ml. of pyridine per 100 ml. has been added. Blot the slide, but not the section and do not rinse. Reduce at 45°C. in 0.1% pyrogallol in 55% alcohol, then rinse in 55% alcohol and wash in water. The remainder of the process consists of gold toning, intensifying in oxalic acid, fixing in 5% Na₂S₂O₃, washing, dehydrating, clearing and covering. When the specimen contains much smooth muscle, the I-KI solution is acidified before use by adding 2 ml. of 1N nitric acid per 100 ml., and the sections treated for 3 minutes instead of the usual 2 minutes. Formalin should not be added to sublimate-acetic, but specimens that do not contain strongly argyrophilic nonneural tissue may be fixed in formalin or, preferably, Bouin's fluid. Sections of tissue after the latter type of fixation will not require the I-KI and thiosulfate but can go from 95% alcohol to the ammoniated alcohol. The advantages of fixing in HgCl₂-acetic acid are suppression of the staining of connective tissue and intensifying the staining of nerve fibers.     

Staining Myelin Sheaths of Optic Nerve Fibers with Osmium Tetroxide Vapor

OsO₄ solution in water, long regarded as the best fixing and staining agent for myelin sheaths, has poor penetrating power. This peculiarity has limited its use to very small pieces of tissue. The vapor from an aqueous solution is known to have a much greater penetrating power for non-neural tissues than the solution itself but nothing has been recorded about its advantages for fixing and staining myelin sheaths of nerve fibers. Difficulties in securing adequate staining of the myelin sheaths in vertebrate optic nerves were overcome largely by the use of the vapor of OsO₄. The technic is carried out as follows: 1) suspend a portion of the nerve above a 2% solution of OsO₄ for 12-24 hours in an air-tight container at room temperature; 2) wash 4-6 hours in distilled water, dehydrate in ethyl alcohol (50% for 2 hours, 70% for 2 hours, and finally 95% overnight), and transfer to n butyl alcohol (2 changes of 2 hours each); 3) embed in paraffin, section, mount and cover in balsam in the customary manner.     

Staining Nerve Fibers with Silver in Tissue Fixed with Bouin's Fluid

Nerve fibers, in organs fixed with Bouin's fluid, are usually refractive to the Davenport silver technic. The axons, however, can be successfully stained if the sections, on slides, are given a preliminary treatment with concentrated pyridine (1 hour), then a 24-hour bath of ammoniated alcohol (99 cc. 80% alcohol, 1 cc 28% ammonium hydroxide) and an interval in 40% aqueous silver nitrate (6-8 hours) before being immersed in the acidified alcoholic silver solution of Davenport. Following the silvering, reduction and toning of the axons, according to the procedure of Davenport, the surrounding non-nervous tissue elements can be counterstained with a combination of either azocarmine, light green and orange G, or azocarmine, aniline blue and orange G.     

Silver Staining of Insect Central Nervous Systems by the Bodian Protargol Method

Improved fixation of ganglia of the central nervous system of Periplaneta americana and Schistocerca gregaria for silver staining by Power's (1943) modification of the Bodian protargol method is given by alcoholic Bouin aged for at least 40 days at 60° C. During impregnation of sections, increased copper and decreased pH give paler staining, more selective for nerve fibres. Prolonging impregnation from 24 to 48 hours weakens the stain and decreases selectivity. The intensity of the stain depends chiefly upon the amount of unreduced (developable) silver combined with the tissues; selectivity is determined mainly by the number and distribution of the reduced silver particles (‘nuclei’). In development, increased sodium sulphite gives more differentiation, increased hydroquinone gives less. Optimum developer composition depends upon impregnation, and thick sections need more differentiation than thinner ones. Within limits, change in one of the factors that control staining can be balanced by changes in others, but by suitable adjustment of the conditions the result can be varied from almost total staining of nerve fibres, for general neuroanatomy, to highly selective staining for tracing individual fibres.     

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.     

Differential Staining Of Tissue In The Block With Picric Acid And The Feulgen Reaction

The authors have found a modification of the Feulgen reaction to be a satisfactory stain for tissue in the block.

Pieces of fresh mammalian tissue not thicker than 5 mm. are fixed for approximately 48 hours at 25° C. in a mixture of equal parts of 5% aqueous sulfosalicylic acid and saturated aqueous picric acid. They are washed for 30 minutes in three ten-minute changes of distilled water and placed in Feulgen's staining solution diluted to one-half strength with distilled water. The staining solution is allowed to act for 24 hours (2 to 3 mm. thick blocks) up to 48 hours for 5 mm. thickness. After staining, the specimens are transferred to a mixture of sodium bisulfite, 0.5 g. and N hydrochloric acid, 5 ml. in' 100 ml. of distilled water. Two changes of IS to 30 min. each in the acid sulfite are given and these are followed by dehydration through 50%, 70% and 95% alcohol. One to two hours are allowed for each change except the last 95%, in which the stained tissue is allowed to remain overnight. The dehydration is completed in two changes of absolute alcohol with subsequent clearing in xylene and embedding in paraffin. Sections may be cut 10 μ or other thickness desired, mounted on slides, paraffin removed, and covered in the usual manner. Nuclei stain reddish violet against a lemon yellow background when the stain is typical. Orange G, 200 mg. per 100 ml. may be added to the fixing fluid if a more polychromatic effect is desired.     

Staining Flagellate Protozoa by Various Silver-Protein Compounds

Seven samples of silver protein (Protargol type) were tested on flagellate protozoa from the alimentary tract of frogs, golden hamsters, and termites. The samples consisted of one of Protargol (Winthrop's prewar German), one of Protargol S (Winthrop-Stearns, Inc., present manufacture, Commission Certified), and five of the experimental batches (4Z, 20, 20B, 22, and 36) by H. A. Davenport and collaborators (1952). The pre-war Protargol is rated best and batch 22 second best for staining of all flagellates. Protargol S gave uniform, but only fair results, with all organisms, while batch 20B, better than Protargol S in several instances, was poorer in a few. Thus Protargol S and batch 20B are rated third, as about equal. Batch 4Z is rated fourth; and batch 20, which stained some species, fifth. Batch 36 stained no protozoa. The tests show that while the present Protargol S is usable for protozoa, it is still inferior to the old German variety. Since some of the experimental batches, none of which was made by exactly the same process, gave promising results, further study of the relation between manufacturing process and subsequent staining reaction should be fruitful.     

Staining Flagellate Protozoa by Various Silver-Protein Compounds

Seven samples of silver protein (Protargol type) were tested on flagellate protozoa from the alimentary tract of frogs, golden hamsters, and termites. The samples consisted of one of Protargol (Winthrop's prewar German), one of Protargol S (Winthrop-Stearns, Inc., present manufacture, Commission Certified), and five of the experimental batches (4Z, 20, 20B, 22, and 36) by H. A. Davenport and collaborators (1952). The pre-war Protargol is rated best and batch 22 second best for staining of all flagellates. Protargol S gave uniform, but only fair results, with all organisms, while batch 20B, better than Protargol S in several instances, was poorer in a few. Thus Protargol S and batch 20B are rated third, as about equal. Batch 4Z is rated fourth; and batch 20, which stained some species, fifth. Batch 36 stained no protozoa. The tests show that while the present Protargol S is usable for protozoa, it is still inferior to the old German variety. Since some of the experimental batches, none of which was made by exactly the same process, gave promising results, further study of the relation between manufacturing process and subsequent staining reaction should be fruitful.     

Staining Nerve Fibers with Methylene Blue. An Evaluation of Variables Used in An Immersion Technic

A study of the effects of osmotic pressure, pH, the presence of dextrose, acetate, pyruvate or lactate, and agents affecting cell permeability during supravital staining by methylene blue was made by means of an immersion technic. Mesentery and intestine of dogs and cats were used. Penetration of the dye was limited to the mesentery and more superficial layers of intestine. Conditions which facilitated the characteristic differentiation of of nerve fibers were: continuous oxygenation of the staining solution, pH about 5.6 stabilized by phosphate buffer, and the presence of small amounts of acetate and lactate. Young animals' tissue stained better than old. Methylene blue was a much more effective staining agent than less completely methylated thionins.     

Visualizing Peripheral Nerve Regeneration by Whole Mount Staining

Peripheral nerve trauma triggers a well characterised sequence of events both proximal and distal to the site of injury. Axons distal to the injury degenerate, Schwann cells convert to a repair supportive phenotype and macrophages enter the nerve to clear myelin and axonal debris. Following these events, axons must regrow through the distal part of the nerve, re-innervate and finally are re-myelinated by Schwann cells. For nerve crush injuries (axonotmesis), in which the integrity of the nerve is maintained, repair may be relatively effective whereas for nerve transection (neurotmesis) repair will likely be very poor as few axons may be able to cross between the two parts of the severed nerve, across the newly generated nerve bridge, to enter the distal stump and regenerate. Analysing axon growth and the cell-cell interactions that occur following both nerve crush and cut injuries has largely been carried out by staining sections of nerve tissue, but this has the obvious disadvantage that it is not possible to follow the paths of regenerating axons in three dimensions within the nerve trunk or nerve bridge. To try and solve this problem, we describe the development and use of a novel whole mount staining protocol that allows the analysis of axonal regeneration, Schwann cell-axon interaction and re-vascularisation of the repairing nerve following nerve cut and crush injuries.     

Staining Similarity of Foot's and Hio4-Schiff Technics

A striking similarity is noted in the staining pictures of Foot's method for reticular tissue and the H104-Schiff technic. This raises the question of the type of chemical bonding involved in the two methods. It suggests that there is a carbohydrate or carbohydrate-type radical involved in the silver staining.     

The Effect of the Quantitative Protargol Stain and Lugol's and Bouin's Fixatives On Cell Size: A More Accurate Estimate of Ciliate Species Biomass

ABSTRACT. The quantitative protargol stain (QPS) is used to estimate ciliate biomass and species composition from mixed field samples. Length, width, breadth and volume of live Euplotes sp., Eutintinnus sp., Strobilidium spiralis, Strombidium acutum , and Gymnodinium sanguineum were compared with 0.6% acid Lugol's fixed, 5% Bouin's fixed, and QPS cells. Cells shrank due to treatments (ANOVA and Tukey's test, α= 0.05). Protistan post-fixation cell volume (as a percentage of live volume) was 55%-80% for acid Lugol's fixed, 40%-70% for Bouin's fixed, and 30%-65% for QPS. Each species shrank to a different extent; cytostructural elements apparently alter the effect of fixation. Egestion is likely not the main cause of shrinkage since the autotroph, G. sanguineum , shrank to the same extent as the heterotrophs when stained by QPS. If field studies do not consider fixation effects on cell size, biomass may be underestimated. We recommend, for studies on planktonic ciliates, either acid Lugol's and QPS be used concurrently or QPS be used alone and biovolume values divided by 0.4 to correct for shrinkage. We stress that this is a rough estimate as this value ranges from 0.3 to 0.45 for planktonic protists.     

Thiazin Dyes in Supravital Staining of nerve Fibers

Supravital staining by thiazins of segments of small intestine and mesentery of young dogs was studied with reference to specificity for nervous tissue. Attempts to secure a purer form of methylene blue by alumina adsorption and alcohol elution of the commercial, medicinal dye yielded a product which appeared to be structurally different from the original dye. The treated dye had absorption maxima from 620 to 655 mμ in contrast with 665 for the untreated. Small nerve bundles were stained by the treated dye after 2 to 4 hours of immersion, but staining was always incomplete. Staining by untreated methylene blue was compared with that by the leucobase, thionol, methylene green, toluidine blue, new methylene blue and the azures. It was concluded that the specificity for nerve fibers resides mainly in the =N(CH₃)₂Cl radical, although some specificity appears to be effected by the methyl groups on the trivalent nitrogen, since azure A (dimethyl) and azure C (mono-methyl) stained weakly, but thionin did not. Methylene green showed some specificity but stained weakly. The leucobase was less active than the reoxidized dye obtained from it.     

Silver Impregnation of Degenerating Axons; Comparisons of Postoperative Intervals, Fixatives and Staining Methods

By introducing variations in procedures and comparing density and patterns of degeneration exhibited, we found that: (1) in the rat, cat, and monkey, the optimum survival time was about 1 wk; this period yielding the maximum amount of argyrophilic reaction and the most consistent staining among several different parts of the brain. Survival times of up to 3 wk did not seriously impair the interpretation of argyrophilial patterns; (2) some qualitative changes in the nuclear patterns of degeneration suggested that the use of a range of survival times may provide useful data; (3) staining differences and similarities in different neural systems of known connections were about the same in the 3 species of animals; (4) formalin, buffered formalin, and a 4%:1% paraformaldehyde-glutaraldehyde mixture were satisfactory perfusion solutions; however, staining was less intense when the Nauta-Gygax method followed the last fixative mixture; (5) The Fink-Heimer I method yielded more consistent results than did the Fink-Heimer II or the Nauta-Gygax methods.