A gelatin film procedure for the localization of proteolytic activity in Leucochloridiomorpha constantiae (Trematoda) adults

A gelatin film procedure was used to localize proteolytic activity in the lumen of the intestinal ceca of Leucochloridiomorpha constantiae (Trematoda) adults. Slides were coated with a 7·5% gelatin solution and then fixed in 10% neutral buffered formalin (NBF). Cryostat sections of isolated worms or those attached to the bursa of Fabricius of the domestic chick were affixed to the gelatin film. Experimental and control slides were incubated in a humid chamber for 30 min at 37·5 and 4°C, respectively. Slides were again fixed in NBF, and then stained for protein with mercuric bromphenol blue (MBB). In experimental slides, the lumen of the intestinal ceca was lysed and did not stain, whereas worm and host tissue and the gelatin were protein-positive. Control sections stained uniformly positive for protein. In this procedure tissue is retained on the slide and proteolytic activity can be correlated with a tissue site on the same slide.     

Histochemical Use Of Lead Tetra-Acetate I. Cleavage Of 1-2 Glycols

Lead tetra-acetate acts specifically to split the carbon-carbon single bond of the 1,2-glycol linkage to produce aldehyde radicals which may then be demonstrated by means of leucofuchsin, 2,4-dinitrophenlyhydrazine, or p-nitrophenylhydrazine. Routinely prepared slide sections from tissues fixed in 10% formalin are run down to 95% alcohol, rinsed in glacial acetic acid and then treated for 2 minutes in a saturated solution of lead tetra-acetate in glacial acetic acid with 5 g. of potassium acetate added for each 100 ml. of reagent. The sections are then washed in distilled water and placed in leucofuchsin for 10 minutes, or in a saturated 30% alcoholic solution of p-nitrophenylhydrazine for 5 minutes or 2,4-dini-trophenylhydrazine for 30 minutes. After staining, the sections are rinsed in 30% alcohol if the nitrophenylhydrazines were used, or in the standard dilute sulfite bath followed by running tap water for 5 minutes if leucofuchsin were used. Sections are routinely dehydrated, cleared, and covered. On examination, the sites of 1,2-glycol linkages will be stained violet by leucofuchsin or yellow by the nitrophenylhydrazines.     

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.     

A Durable Nissl Stain for Frozen and Paraffin Sections

Frozen sections, 25-50 /j. thick, of formalin-fixed nervous tissues are mounted following the Albrecht gelatin technic. Paraffin sections, 15 p., are deparaffinized and transferred to absolute ethanol. The slides are then coated with celloidin. Both frozen and paraffin sections subsequently follow the same steps: absolute ethanol-chloroform (equal parts) for at least 20 min, 95% ethanol, 70% ethanol (1-3 min), then rinsed in distilled water. Sections are stained in Cresylechtviolett (Chroma) 0.5% aqueous solution containing 4 drops of glacial acetic acid per 100 ml, rinsed in distilled water, agitated in 70% ethanol until excess stain leaves the slide, and rinsed in 95% ethanol. Sections are then dehydrated in absolute ethanol, followed by butanol, cleared in xylene, and enclosed in permount.     

New Fuchsin-Hematoxylin-Eosin; A Stain for Acid-Fast Bacilli and Surrounding Tissue

This is a staining technique for histopathologic evaluation of tissue reaction in the environs of acid-fast tubercle bacilli (avian and bovine) in sections. Fresh tissue is fixed in 10% neutral formalin and processed in the usual manner for embedding in paraffin. Sections are cut approximately 6 μ. thick, dewaxed, hydrated, and stained with Harris' hematoxylin. They are rinsed in tap water, differentiated in add alcohol, washed in tap water, given a distilled water rinse and stained at 20-30° C in a 1% solution of new fuchsin in 5% phenol. Each slide is then handled individually by placing it directly into a saturated aqueous solution of Li₂CO₃ and agitated gently for a few seconds. This is followed by differentiation with 5% glacial acetic acid in absolute or 95% ethyl alcohol until the color stops running. Two rinses in absolute or 95% ethyl alcohol follow. The sections are then counterstained in the color add of eosin Y prepared according to the method of Schleicher (Stain Techn., 28, 119-23, 1953) and used as an 0.025% solution in absolute alcohol. Following passage through 2 changes of absolute alcohol, the sections are cleared in xylene, then mounted in Permount or similar synthetic resin. The add-fast barilli are emphasized by their bright retractile red color within a contrasting background of hematoxylin and eosin.     

New Fuchsin-Hematoxylin-Eosin; A Stain for Acid-Fast Bacilli and Surrounding Tissue

This is a staining technique for histopathologic evaluation of tissue reaction in the environs of acid-fast tubercle bacilli (avian and bovine) in sections. Fresh tissue is fixed in 10% neutral formalin and processed in the usual manner for embedding in paraffin. Sections are cut approximately 6 μ. thick, dewaxed, hydrated, and stained with Harris' hematoxylin. They are rinsed in tap water, differentiated in add alcohol, washed in tap water, given a distilled water rinse and stained at 20-30° C in a 1% solution of new fuchsin in 5% phenol. Each slide is then handled individually by placing it directly into a saturated aqueous solution of Li₂CO₃ and agitated gently for a few seconds. This is followed by differentiation with 5% glacial acetic acid in absolute or 95% ethyl alcohol until the color stops running. Two rinses in absolute or 95% ethyl alcohol follow. The sections are then counterstained in the color add of eosin Y prepared according to the method of Schleicher (Stain Techn., 28, 119-23, 1953) and used as an 0.025% solution in absolute alcohol. Following passage through 2 changes of absolute alcohol, the sections are cleared in xylene, then mounted in Permount or similar synthetic resin. The add-fast barilli are emphasized by their bright retractile red color within a contrasting background of hematoxylin and eosin.     

Hematein-its Advantages for General Laboratory Usage

The advantages of hematein over hematoxylin are that it is easy to prepare, easy to use, and saves time; while it gives equally good results. The writer has been employing for some time a pre-war imported hematein and until within the last few months has been unable to locate a satisfactory product of recent manufacture, either domestic or foreign. At the request of the Biological Stain Commission, however, an American manufacturer has at last put on the market a C. P. hematein which gives splendid results. The technic is as follows:

Paraffin or celloidin sections of Bouin or Zenker-formol material are run down to water and stained about 5 minutes in Mayer's hemalum (0.5 g. hematein ground up in a glass mortar with 10 cc. 95% alcohol and added to 500 cc. of 5% aqu. sol. potassium alum.) Rinse 1 to 3 seconds in tap water. Dip 1 to 3 seconds in eosin B (1 part 0.5% sol. in 20% alc. added to 2 pats dist. water; filtered from time to time). Wash several minutes in running water or in several changes of tap water. Dehydrate and mount; with unattached celloidin sections this may be done by running up to 95% alcohol, spreading on slide, blotting, wetting with absolute alcohol, draining and mounting in euparal.     

A STAINING TECHNIQUE FOR AUTORADIOGRAPHS OF NONDIFFUSIBLE ISOTOPES

Nondeparaffinized radioactive tissue sections are stained with hematoxylin and eosin by being floated on aqueous solutions for 1 hr each. The sections are then thoroughly washed, dried and exposed to autoradiographic plates or emulsions for predetermined periods of time. When desirable, both stained and unstained adjacent tissue sections can be mounted on a single slide of autoradiographic plate for exposure. Kodak DK-19 and 30% Na₂S₂O₃.5H₂O solutions are used for subsequent developing and fixing. The finished autoradiographs show excellent resolution and cytologic detail, since the gelatin remains unstained while the tissue retains its stain. Stains other than hematoxylin and eosin can be applied to this technique, provided they withstand the developmental and fixation processes.     

Notes on Selling'S Green-Light Method for Critical Microscopy (Continued)

Kill root tips in 1 part glacial acetic acid to 3 parts absolute alcohol for 12 or more hours. Remove from killing fluid and place for 5 to 10 minutes in a solution consisting of 1 part 95% alcohol to 1 part concentrated HCl. Transfer to Carnoy's fluid for 5 minutes or longer. Cut a small piece (0.5 mm. or less) off the tip of the root and place on a clean slide in a small drop of iron-aceto-carmin stein. Press directly on the piece of root with a small flat scalpel; the cells will now separate and float free in the stain. Place cover slip over the drop of stain and apply gentle pressure. Heat carefully by passing the slide 3 or 4 times thru the flame of an alcohol lamp. Seal with heated mixture of 1 part Parowax to 1 part gum mastic. Make permanent by the McClintock permanent method.     

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.     

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.     

A Comparative Survey of Section Adhesives and of Factors Affecting Adhesion

Equal-size pieces of spleen, liver, cerebrum, tonsil and myocardium were taken from human postmortem tissue and sections of the following kinds were made: cryostat, CO₂ freezing microtome, paraffin, and double-embedded celloidin-paraffin. Fixation was in 10% formol-saline with the exception of the cryostat material which was fresh-frozen. The sections thus prepared were attached to identical sets of slides with glycerol-albumen, soluble starch, amylopectin starch, human plasma, 0.2% gelatin, 0.2% gelatin-formol, by flattening with 50% alcohol, and including a control slide without adhesive. Identical batches of the mounted sections were then dried overnight at 18-20, 37 and 56 C followed by washing in running tap water, 10% NH₄OH at 18-20 C and 1% NaOH and 10% NaOH solution at 56 C over a period of 4 days. Sections separating from the slides during successive intervals of soaking showed that plasma was the overall best adhesive and 56 C the most effective drying temperature. Agar, in the limited tests applied, was found to be quite effective but had a tendency to take up nuclear dyes.     

Histochemical Use of Sodium Bismuthate

Histochemical 1,2-glycoI cleavage, similar to that obtained with periodic acid and lead tetraacetate, may be obtained with sodium bismuthate. Routinely prepared slide sections, from tissues fixed in 10% formalin, are run down through xylene and graded alcohols to water and then oxidized for three minutes in a 1% sodium bismuthate 20% aqueous phosphoric acid solution. The oxidizing solution must be freshly prepared and used immediately. Following oxidation, sections are rinsed 15 sec. in IN HC1 to remove bismuth pentoxide precipitate, a by-product of the reaction. The sections are then washed in distilled water and placed in leuco-fushsin for 10 min., or in a saturated 30%) alcoholic solution of p-nitrophenylhydrazine for 5 min. or 2,4-dinitrophenylhydrazine for 30 minutes. After staining, the sections are rinsed in 30% alcohol if the nitrophenylhydrazines were used, or in the standard dilute sulfite bath followed by running tap water for 5 min. if leucofuchsin were used. Sections are routinely dehydrated, cleared, and covered. On examination, the sites of 1,2-glycol linkages will be stained violet by leucofushsin or yellow by the nitrophenylhydrazines.     

Histochemical Use of Sodium Bismuthate

Histochemical 1,2-glycoI cleavage, similar to that obtained with periodic acid and lead tetraacetate, may be obtained with sodium bismuthate. Routinely prepared slide sections, from tissues fixed in 10% formalin, are run down through xylene and graded alcohols to water and then oxidized for three minutes in a 1% sodium bismuthate 20% aqueous phosphoric acid solution. The oxidizing solution must be freshly prepared and used immediately. Following oxidation, sections are rinsed 15 sec. in IN HC1 to remove bismuth pentoxide precipitate, a by-product of the reaction. The sections are then washed in distilled water and placed in leuco-fushsin for 10 min., or in a saturated 30%) alcoholic solution of p-nitrophenylhydrazine for 5 min. or 2,4-dinitrophenylhydrazine for 30 minutes. After staining, the sections are rinsed in 30% alcohol if the nitrophenylhydrazines were used, or in the standard dilute sulfite bath followed by running tap water for 5 min. if leucofuchsin were used. Sections are routinely dehydrated, cleared, and covered. On examination, the sites of 1,2-glycol linkages will be stained violet by leucofushsin or yellow by the nitrophenylhydrazines.     

Autoradiography of Water-Diffusible Substances in Sections of Whole Baby Rats

Rats, 7 days postnatal which had been injected with a radioactive nuclide, were quick frozen and sectioned in the frozen state. An adhesive cellulose tape (Sellotape) was used to support the section during cutting, through freeze-drying, and attaching to slides. Dehydration of the frozen sections consisted of 1 hr in a chilled desiccator containing silica gel, then at reduced pressure of 2-3 mm Hg until quite dry. The exposed side of the section was sprayed with celloidin dissolved in amyl acetate and allowed to dry. This side of the section was attached to a slide, previously coated with 1% gelatin containing 0.1% chrome alum, by means of an adhesive consisting of 4% gelatin and 5% formalin in 60% glycerol. In applying this adhesive it is mandatory that a border of about 3 mm of bare glass be left outside the adhesive, to allow intimate contact between the sticky side of the tape and the glass. The adhesive was allowed to set for 20 min, the slide immersed in water lor 50 sec, and the cellulose layer of the tape peeled off. The rubber base from the tape was removed with chloroform, the slide dried, and the exposed surface of the section coated with celloidin in amyl acetate, by dipping. After this treatment, the slides could be coated by dipping in autoradiographic emulsion without affecting water-soluble radioactive substances in the tissue.     

A Modification of the Cresyl Violet Technic for Staining Nerve Cells

Kill root tips in 1 part glacial acetic acid to 3 parba RB Solute alcohol for 12 or more hours. Remove from king fluid a d place for 5 to 10 minutes in a solution consisting of 1 part 95% alcohol to 1 part concentrated HC1. Transfer to Carnoy's fluid for 5 minutes or longer. Cut a small piece (0.5 mm. or less) off the tip of the root Press directly on the piece of root with a small fiat scalpel; the cells will now separate and float free in the stain. Place cover slip over the drop of stain and apply gentle pressure. Heat carefully by paseing the slide 3 or 4 times thru the flame of an alcohol lamp. Seal with heated mixture of 1 part Parowax to 1 part gum mastie. Make permanent by the McClintock permanent method. and place on a clean slide in a small drop of iron-ace-sinin.     

Polystyrene embedding: a new method for light and electron microscopy.

Polystyrene embedments of histological specimens can be obtained with a solution of 1:14 polystyrene-toluene, 5% benzyl alcohol and 1% dibutyl phthalate, allowing the solvent to evaporate in polyethylene containers for 2-3 days at 58 C. The resulting blocks are easily cut into truncated pyramids, each containing a piece of tissue, which are then glued to a Plexiglas support. Drying is completed at 80 C for 20 hr. The pyramids can then be sectioned to produce thick sections with a steel knife or to produce semi- or ultrathin sections with a glass knife. A 10% paraldehyde solution is used to mount the light microscopy sections on a slide heated on a hot plate to 80 C; these can be treated with the same techniques used with paraffin sections. The results are of high quality. Semithin sections of tissues fixed for electron microscopy can be stained directly after mounting, or by a wider range of stains once the polystyrene has been removed by organic solvents. In electron microscopy, the ultrathin sections obtained with the usual techniques are highly electron beam-resistant and given acceptable results.     

An Abrasive Paper Technique for Preparing Sections of Wood for Microscopic Study

Dry wood specimens are sawed to 2mm thickness and impregnated with resin such as Lewisol 28 (Hercules Powder Co.). One side of the specimen is ground by hand on abrasive papers of grades #100, #180, #240, and #320. This side is then cemented to a petrographic glass slide with stick shellac and the other side similarly ground. Scratches can be eliminated by scraping the ground surface with the sharp edge of an ordinary glass microscopic slide. The section is removed by heating the slide, dissolving the shellac with alcohol and the resin of the embedding matrix with xylene. The sections can be stained in a hot saturated alcoholic solution of safranin O, counterstained with 0.01% fast green in an equal parts mixture of clove oil, methyl cellosolve, and absolute alcohol, and mounted in balsam.     

Polystyrene Embedding: A New Method for Light and Electron Microscopy

Polystyrene embedments of histological specimens can be Obtained with a solution 1 : 4 polystyrene-toluene, 5% benzyl alcohol and 1% dibutyl phthalate, allowing the solvent to evaporate in polyethylene containers for 2-3 days at 58 C. The resulting blocks are easily cut into truncated pyramids, each containing a piece of tissue. which are then glued to a Plexiglas support Drying is completed at 80 C for 20 hr. The pyramids can then be sectioned to produce thick sections, with a steel knife or to produce semi- or ultrathin sections with a glass knife. A 10% paraldehyde solution is used to mount the light microscopy dons on a slide heated on a hot plate to 80 C; those can be treated with the same techniques used with paraffin sections. The results are of high quality. Semithin sections of tissues fired for electron microscopy can be stained directly after mounting, or by a wider range of stains once the polystyrene has been removed by organic solvents. In electron-microscopy, the ultrathin sections obtained with the usual techniques are highly electron beam-resistant and give acceptable results.     

Formic Acid Corrosion for the Demonstration of Pulmonary Elastic Tissue

By a revised technique, human pulmonary elastic tissue can be isolated in a form suitable for examination under the stereoscopic microscope. Fresh human lungs from autopsy are fixed by intrabronchial infusion with 10% formalin for 24 hr. Slabs 1.5 cm thick are cut and the formalin removed in running water. One such slab is embedded under intermittent vacuum in an aqueous mixture containing 15% gelatin, 10% glycerol, and 1% phenol; then allowed to gel. Frozen sections 2 mm thick are cut on a large-section MSE sledge microtome. Squares 3 × 3 cm from such a section are corroded for 4-5 days in 88% formic acid at 45 C, washed once with distilled water, and mounted in glychrogel containing 6% gelatin. The elastic tissue network of the lung will have been freed from surrounding elements. The preparation should be stored in a refrigerator. Blocks for thin sections and large thick un-corroded sections can be prepared from the same lung as part of an over-all procedure.