Impregnation of Oligodendroglia in Nervous Tissue Kept in Formalin for Many Years

A method for impregnating oligodendroglia in nervous tissue (monkey) fixed and preserved in formalin for many years is described. This tissue is reconditioned by placing 12 to 30μ frozen sections of it in concentrated ammonia (sp. gr. 0.90) and by washing them slowly for 24 hours with a 1 mm. stream of water. The fluid is then poured off the sections; the jar is refilled with concentrated ammonia; and washing is repeated for another 24 hours. The sections are then plunged into concentrated ammonia for 7 minutes.

After treatment in ammonia, the sections are incubated for one hour at 38^oC. in Globus' 5% hydrobromic acid solution. They are washed again, in distilled water, and then impregnated in a “medium” strength ammoniacal silver carbonate solution (5 ml. of 10% AgNO₃ added to 15 ml. of 5% Na₂CO₃. The precipitate is dissolved in concentrated ammonia and diluted to SO ml. with distilled water). Impregnation is followed by reduction in 1% formalin without agitation; fixation in 5% Na₂S₂O₃; dehydration, and mounting in clarite.

Typical oligodendroglia (Fig. 1) were made visible by use of the method outlined in this paper.     

A Golgi-electron microscope method for insect nervous tissue.

Golgi's light microscope method of selective silver impregnation for nervous tissue combined with electron microscopy appears to offer a promising method for working out the detailed anatomy of individual neurons and their connections. Insect nervous tissue is fixed in a mixture of 2% paraformaldehyde and 2 1/2% glutaraldehyde in Millonig's buffer (pH 7.2) before postfixation for 12 hours in a solution brought to pH 7.2 with KOH containing 2% potassium dichromate, 1% osmium tetroxide and 2% D-glucose. The tissue is then transferred to a solution of 4% potassium dichromate for 1 day; and for 1-2 days to a 0.75% silver nitrate solution. After dehydration and embedding in Araldite, 50 mum sections are made. Areas of interest are cut from these sections and re-embedded in silicone molds. Ultrathin sections are then cut and stained with uranyl acetate and lead citrate. The Golgi method described here gives good results at the level of both light and electron microscopy.     

A Golgi-Electron Microscope Method for Insect Nervous Tissue

Golgi's light microscopic method of selective silver impregnation for nervous tissue combined with electron microscopy appears to offer a promising method for working out the detailed anatomy of individual neurons and their connections. Insect nervous tissue is fixed in a mixture of 2% paraformaldehyde and 21/2% glutaraldehyde in Millonig's buffer (pH 7.2) before postfixation for 12 hours in a solution brought to pH 7.2 with KOH containing 2% potassium dichromate, 1% osmium tetroxide and 2% D-glucose. The tissue is then transferred to a solution of 4% potassium dichromate for 1 day; and for 1-2 days to a 0.75% silver nitrate solution. After dehydration and embedding in Araldite, 50μm sections am made. Areas of interest are cut from these sections and re-embedded in silicone molds. Ultrathin sections are then cut and stained with uranyl acetate and lead citrate. The Golgi method described here gives good results at the level of both light and electron microscopy.     

Differentiation of the Secretory Cells of the Pars Nervosa of the Hypophysis Cerebri

A method was found by means of which two types of granular cells in the pars nervosa of the hypophysis cerebri could be preserved in permanent preparations so as to retain the appearance these cells presented in fresh tissue and in tissue cultures. The essential points of the technic were the fixation for 24 hours of the hypophysis in a solution composed of 2 parts of 3% potassium bichromate plus one part of a one-half saturated solution of bichloride of mercury in 95% alcohol. Sections of this material were prepared using dioxan in place of the higher alcohols and xylene. The sections were stained by means of Mallory's connective tissue stain leaving the sections in the fuchsin solution for 30 minutes and in the mixture of aniline blue, orange G and phosphotungstic acid for 1 to 24 hours.     

High-efficiency liquid-scintillation counting of 14C-labelled material in aqueous solution and determination of specific activity of labelled proteins

1. Inexpensive scintillation mixtures are described which enable the detection of as little as 40μμc of ¹⁴C in aqueous solution with an efficiency of counting of over 80%. 2. A rapid method for the counting of alkaline, acidic and neutral aqueous solutions of up to 1ml. volume is described. Ethanol or 2-ethoxyethanol is used as blending agent. 3. The scintillation counting of alkaline solutions is applied to the accurate determination of the specific activity of ¹⁴C-labelled proteins from plant tissues. 4. Attention has been paid to the importance of a standardized washing procedure for the removal of all traces of radioactive material from glassware.     

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.     

Syntheses and thermal characterization of the magnesium phthalocyanine complexes with 2-methoxy- and with 2-ethoxyethanol. X-ray structure of (2-ethoxyethanol)magnesium phthalocyanine

The syntheses of the magnesium phthalocyanine complexes with dry 2-methoxy-ethanol and 2-ethoxyethanol have been performed by recrystallization method using both anhydrous MgPc and aquated magnesium phthalocyanine, MgPcH₂O, as a starting material. It has turned out that in the temperature range below ca. 140 °C, the bi-axially ligated complexes are formed, i.e., MgPc(2-methoxyethanol)₂ and MgPc(2-ethoxyethanol)₂ with 4 + 2 coordination of Mg, whereas at higher temperatures, up to about 200 °C, the mono-axially ligated complexes are stable, i.e., MgPc(2-methoxyethanol) and MgPc(2-ethoxyethanol) with 4 + 1 coordination of Mg.The single crystal structure of MgPc(2-ethoxyethanol) complex has been determined. The central Mg atom is displaced towards the hydroxyl group of the ligand by about 0.37 Å from the (N-isoindole)₄ plane of the Pc ring. Hydrogen bonds of the type O-H?N between the hydroxyl groups of 2-ethoxyethanol and one of the azamethine N-atoms of the Pc ring link the molecules related by a centre of symmetry. Such a packing arrangement in the crystal leads to a dimerization with the ligand-to-Pc connections. The syntheses, thermogravimetric results and structure characteristics are compared with the known MgPc complexes with O- and N-donating molecules.     

A silver carbonate method for cell counts of neurons and glial elements on paraffin embedded brain tissue.

A modification of the Del Rio-Hortega method for the demonstration of central nervous system elements is presented. This silver impregnation technique is particularly useful for the classification of cell types for quantitative differential cell counts. Formalin fixed paraffin sections are immersed in formol-ammonium bromide for 1 1/2 hours; this solution is an excellent mordant for various silver nitrate stains. The samples are stained for 20 to 60 minutes in a silver carbonate solution (25 ml of 25% silver nitrate combined with 200 ml of 5% sodium carbonate) and then reduced in a 1% formaldehyde solution to which 20 drops of acetic acid have been added. Finally, the slides are fixed in sodium thiosulfate, rinsed in tap water, dehydrated, cleared, and mounted. This procedure will enable this investigator to identify neurons, oligodendroglia, and astrocytes on the basis of their nuclear staining as well as to demonstrate the laminae of brain tissue since the method allows differentiation of cell layers and fiber tracts.     

Alizarin Red S and Toluidine Blue for Differentiating Adult or Embryonic Bone and Cartilage

This technic has been successfully employed by the author for staining, in toto, the bones and cartilage of mature specimens of Urodela and the developing bone and cartilage of the embryonic human, cat, pig and rat. The differential staining is accomplished by using a modification of Dawson's method of staining bone with alizarin red S following a toluidine blue solution specific for cartilage. Specimens are fixed in 10% formalin, stained one week in a solution of .25 g. of toluidine blue in 100 cc. of 70% alcohol, macerated 5 to 7 days in a 2% KOH solution, counterstained for 24 hours in a 0.001% solution of alizarin red S in 2% aqueous KOH, dehydrated in cellosolve and cleared in methyl salicylate. In the adult and embryonic forms thus treated the soft tissues are cleared while the osseous tissue is stained red, the cartilage blue.     

Postembedding adjuvants in routine microtomy of paraffin-processed tissues.

Soaking the exposed face of routine paraffin-processed tissue blocks from human autopsies in solutions containing either 60% ethyl alcohol-glycerol 9:1; glycerol-aniline 9:1; 5% aqueous phenol; 5% aqueous Photo-flo, or 10% aqueous ammonium hydroxide for 2--4 hours at 0--4 C resulted in greater ease of ribboning and overall improvement of slide quality in the majority of sections compared to the results with the same blocks soaked only in water at 0--4 C. The use of postembedding adjuvants, especially 10% aqueous ammonium hydroxide, presents a simple, convenient, inexpensive means of increasing section quality with routine as well as refractory paraffin-embedded tissue without sacrificing tissue adhesion, staining qualities or cellular detail.     

Paradichlorobenzene and Other Chemicals in Chromosome Work

The action of different prefixing agents including water and a dilute CaCl₂ solution on plants with high chromosome number showed that all the solutions tried exhibited marked effect on the physical state of the plasma. Also the plasmatic constitution of the pretreated tissue differed from the appearance of one fixed directly in dye-acid mixture for a few seconds without any pre-treatment. For the clarification of the karyotypes, the best results have been obtained with paradichlorobenzene. It was used as a prefixing agent with a short schedule for temporary smears of difficult materials, particularly those with high chromosome number. The method comprised treatment of the root tips in a saturated aqueous solution of the chemical for 3 hours at 12-16°C., followed by 4-5 seconds heating just to boiling in orcein-hydrochloric acid mixture, then smearing in 1% aceto-orcein solution.     

Aerosol ot Solution—An Effective Softener of Herbarium Specimens for Anatomical Study

Dried plant parts are cut into convenient sizes and soaked in a solution containing 2.5-3.3% Aerosol OT in distilled water for 5 or more hours until well penetrated by the Aerosol. After brief washing in distilled water the material can be embedded, can be sectioned freehand or, if the nature of the material permits, with a sliding microtome, without embedding. Although it is a good wetting agent, Aerosol is chemically neutral; therefore, microchemical tests can be performed successfully on material treated with it. Refractory plant tissues embedded in paraffin can be successfully softened if one face of the block is trimmed to expose the tissue, then soaked in an Aerosol solution before sectioning.     

Changes in laminin immunoreactivity as a marker for the state of liver preservation

Summary In order to examine the role of the extracellular matrix glycoprotein laminin as a marker for the preservation of liver tissue, dog livers were perfused and then preserved for 5 min, 1, 2, 4, 6, 8, 10, 12, 22 and 26 hours with HTK (histidine-tryptophan-ketoglutarate) solution at 5°C and at 25°C and with UW (University of Wisconsin) solution at 5°C. The tissue was processed for the immunohistochemical demonstration of laminin using an anti-P1 and an anti-E8 antibody. The peroxidase-antiperoxidase method was used for the visualization of the immunohistochemical reaction. At the beginning of the preservation, immunostaining was observed for both fragments of laminin around bile ducts and blood vessels of the portal spaces, under all preservation conditions. Clear immunostaining was also visible in the wall of the terminal arterioles located between the liver lobules. In the 5°C-preserved tissues, immunostaining for both laminin fragments occurred for preservation times between 4 and 6 hours in the form of isolated perisinusoidal deposits at the transition point where the sinusoids sprout from the terminal venules. In the 25°C-preserved tissue, such a staining pattern was already visible after 1 to 2 hours, preservation time. Our results show that the occurrence of laminin immunoreactivity in the sinusoids can be taken as a marker for the state of liver preservation. A hypothesis for the presence and the role of this glycoprotein in the perisinusoidal space is presented.     

Chlorazol Black E As An Aceto-Carmine Auxiliary Stain

In making chromosome counts on plants and plant parts treated with colchicine it was found that in cases where aceto-carmine alone is not satisfactory—as in axillary buds of apple, pear, plum, peach, apricot, and cherry—the following method was effective : Dissect out the meristematic parts of the axillary bud under a binocular (or cut free-hand sections) and transfer the dissected tissue immediately to a solution of 3 volumes alcohol to 1 volume acetic acid for killing and fixing. Let the fixative act at least 10 minutes; a longer time, 12-24 hours, improves the staining quality. Wash in at least 3 changes of 70% alcohol to remove most of the acid. Stain for 5-25 minutes in 1% chlorazol black E² in 70% alcohol. Rinse in 3 changes of 70% alcohol to remove excess stain. Transfer the material to a slide, cover with a drop of aceto-carmine, and if necessary, dissect further under a binocular. Cover with cover glass, heat, flatten and seal, or run Zirkle's fluid under the cover for permanent mounting. For smears of sporocytes, chlorazol black E may also be employed alone, or in combination with aceto-carmine, if a dark purple nuclear stain is desired.     

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 new method for easy demonstration of argyrophil cells.

A new method for silver impregnation of endocrine cells of the gastrointestinal mucosa is described. It offers great reliability, eveness of impregnation, and, since it can be used on batches of slides, is also suitable for histology class and investigation material. The procedure for paraffin sections of formalin-fixed material is as follows: dewax and transfer to distilled water, leave in 0.5% silver nitrate solution for 2 hours at 60 C. Rinse in distilled water, then treat in Bodian developer (hydroquinone, 1 g; sodium sulphite, 5 g; distilled water, 100 ml) previously heated to 60 C. Rinse in running tap water, distilled water, and then re-impregnate for 10 minutes at 60 C in the same silver solution and reduce in Bodian's solution. Since the background is not impregnated by this method, sections may be counterstained by any basic anilin dye to bring out nuclei. A 0.1% kernechtrot solution was found very satisfactory in this respect. The granulations of argyrophil cells stand out sharply black against a red background.     

A New Method for Easy Demonstration of Argyrophil Cells

A new method for silver impregnation of endocrine cells of the gastrointestinal mucosa is described. It offers great reliability, eveness of impregnation, and, since it can be used on batches of slides, is also suitable for histology class and investigation material. The procedure for paraffin sections of formalin-fixed material is as follows: dewax and transfer to distilled water, leave in 0.5% silver nitrate solution for 2 hours at 60 C. Rinse in distilled water, then treat in Bodian developer (hydroquinone, 1 g; sodium sulphite, 5 g; distilled water, 100 ml) previously heated to 60 C. Rinse in running tap water, distilled water, and then re-impregnate for 10 minutes at 60 C in the same silver solution and reduoc in Bodian's solution. Sma the background is not impregnated by this method, sections may be counterstained by any basic anilin dye to bring out nuclei. A 0.1% kernechtrot solution was found very satisfactory in this respect. The granulations of argyrophil cells stand out sharply black against a red background.     

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.     

Postembedding Adjuvants In Routine Microtomy Of Paraffin-Processed Tissues

Soaking the exposed face of routine paraffin-processed tissue blocks from human autopsies in solutions containing either 60% ethyl alcohol-glycerol 9:1; glycerol-aniline 9:1; 5% aqueous phenol; 5% aqueous Photo-Flo, or 10% aqueous ammonium hydroxide for 2-4 hours at 0-4 C resulted in greater ease of ribboning and overall improvement of slide quality in the majority of sections compared to the results with the same blocks soaked only in water at 0-4 C. The use of postembedding adjuvants, especially 10% aqueous ammonium hydroxide, presents a simple, convenient, inexpensive means of increasing section quality with routine as well as refractory paraffin-embedded tissue without sacrificing tissue adhesion, staining qualities or cellular detail.     

Staining Paraffin Sections with Protargol: 1. Experiments with Bodian's Method. 2. Use of Jvpropyl and N-Butyl Alcohol in Hofker's Fixative:

Paraffin sections of nervous tissue, which had been fixed in Hofker's fluid, stained readily with protargol solution without the addition of metallic copper or other activator. Amidolsulfite mixtures reduced the protargol more rapidly and completely than hydroquinone-sulfite. Intensification of the stain could be secured by reducing with 0.5% amidol (or pyrogallol) solution after gold toning. The completeness of staining of unmyelinated fibers of the dorsal roots of cat spinal nerves was checked by estimating the number of fibers in a root and the cells of its associated ganglion. A fiber cell ratio of 1:1 was found hi 4 specimens, indicating within limits of error that all fibers were stained. An improvement of die original Hofker's mixture as a fixative was obtained by using a mixture of formic acid, 5 cc.; trichloracetic acid, 10 g.; n-propyl alcohol, 20 cc.; and n-butyl alcohol, 60 cc. (instead of the acetic, trichloracetic, ethyl alcohol mixture used hi the original formula). The following arbitrary method is suggested. Fix 12 to 24 hours, pass to water thru graded ethyl alcohol, wash several hours, dehydrate and embed in paraffin. Cut, mount, and remove the paraffin, pass to water and impregnate 2 or 3 days at 27 to 30$$C. in a 0.5% aqueous solution of protargol (Winthrop Chemical Co.). Rinse 2 or 3 seconds and reduce with 0.5% amidol (Agfa brand used) in 5% sodium sulfite solution. Wash, tone with 0.1% gold chloride, wash and reduce with 0.5% amidol (no sulfite), wash, dehydrate and cover. The method works well on spinal nerve roots, cerebrum, cerebellum, and spinal cord, and moderately well on nerve trunks including sympathetic nerves. Tissues from cat and guinea pig were used.