A solvent-free coating-procedure for the improved preparation of cryostat sections in light microscope histochemistry

Summary A new technique is presented for the external stabilization of cryostat sections by spraying the specimen surfaces with an aqueous solution of poly(vinyl alcohol) before each sectioning stroke. The spray freezes upon the surface and forms a tough coating which facilitates subsequent sectioning and handling especially of difficult material. The sections are affixed upon cold glass slides covered with an improved formulation of pressure-sensitive adhesive. During further processing of the affixed sections, the PVA-coating and any surrounding supporting medium dissolve without traces in the first aqueous incubation or staining solution.     

A simple method for preparing thin (10 microM) histological sections of undecalcified plastic embedded bone with implants

A simple method for preparing undecalcified thin sections of bone with implants has been developed. After exposing a surface of bone and implant in a plastic block by sawing thick sections, the surface is stained prior to making a thin section. A glass coverslip is affixed with a thin layer of cement to the stained surface to stabilize the tissue and implant during sectioning. A mixture of glycerine and water is used as a coolant and lubricant. The orientation in situ is preserved allowing demonstration of bone architecture and cells, and the tissue-implant interface.     

A Simple Method for Preparing Thin (10 μm) Histological Sections of Undecalcified Plastic Embedded Bone with Implants

A simple method for preparing undecalcified thin sections of bone with implants has been developed. After exposing a surface of bone and implant in a plastic block by sawing thick sections, the surface is stained prior to making a thin section. A glass coverslip is affixed with a thin layer of cement to the stained surface to stabilize the tissue and implant during sectioning. A mixture of glycerine and water is used as a coolant and lubricant. The orientation in situ is preserved allowing demonstration of bone architecture and cells, and the tissue-implant interface.     

The preparation of microtome sections of unaltered soil for the study of soil organisms in situ

Summary A new technique for study of small soil organisms in situ in unaltered soil is described.The soil samples are cooled in a refrigerator at — 10°C to kill the animals. A small portion taken from a frozen soil sample, is slowly immersed in a solution of gelatin. When the specimen is infiltrated with gelatin and the whole cooled it is fixed in formalin to enable it to withstand treatment with hydro-fluoric acid for removal of sand grains. Subsequently the specimens are immersed in gelatin solution for a second time after which the specimens are affixed to wooden blocks which can be clamped in the microtome. Before sectioning, the embedded specimen affixed to the wooden block is hardened in methylalcohol after which it is possible to cut sections 7,5–10µ thick.The most satisfactory staining procedure proved to be the quadruple staining method of Johansen. By this method nematodes, fungi, bacteria and amoebae are easily distinguishable from the soil particles.     

Dichromate-Chlorate Perfusion Prior to Staining Degeneration in Brain and Spinal Cord

A method of fixation compatible with both the Nauta-Gygax and Swank-Davenport procedures for degenerating nerve fibers, which shortens the time required by the former procedure, is as follows: The central nervous system is perfused with a 0.9% aqueous solution of NaCl followed by an aqueous solution containing 5% K₂Cr₂O₇ and 2.5% KClO₃. The central nervous system is then hardened in 10% formalin for 1-3 days. Tissue for Marchi-type staining can be taken at this stage. For silver staining, the processing is continued by immersion overnight in 10% formalin in 20% alcohol, and frozen sections cut the next day. Sections, up to 50μ in thickness, are collected in 10% formalin and impregnated by the Nauta-Gygax technique. Best results are obtained by impregnating within 24-48 hr after sectioning.     

Staining Methods For Studying Ingredient Distribution In Baked Cakes

A method is described for preparing cake crumb for sectioning and staining. Previous to embedding, the fat was stained and fixed by exposing small blocks of cake to the fumes from a 5%, freshly-prepared, aqueous solution of osmic acid (OsO₄). This was followed by dehydration in ethyl alcohol and tertiary butyl alcohol, removal of air under vacuum and infiltration with paraffin.

Sections were cut 20 and 9Op thick and mounted with water.

Wax was removed by immersion in xylene. The sections were rehydrated in a series of ethyl alcohol dilutions, from concentrated to dilute, then transferred to distilled water.

Protein was then stained pink by immersion of the slides in an acidified 0.04% water solution of eosin Y, or starch was stained blue with a dilute aqueous solution of iodine. Ten grams iodine and 10 g. KI were dissolved in 25 ml. distilled water. This stock solution was diluted for use one to two hundred times.

The relationship between protein and starch was demonstrated by staining the sections with eosin, differentiating in 50% alcohol and staining with iodine.

When slides of cake crumb were prepared in this way, the fat was stained black, the protein bright pink and the starch granules a dark blue.     

A TECHNIQUE FOR ULTRACRYOTOMY OF CELL SUSPENSIONS AND TISSUES

Ultracryotomy of fixed tissue has been investigated for a number of years but, so far, success has been limited for several reasons. The simple technique herein reported allows the ultracryotomy not only of a variety of tissues but also of single cells in suspension, with a preservation and visualization of ultrastructural detail at least equivalent to that obtained with conventional embedding procedures. In this technique, sucrose is infused into glutaraldehyde-fixed tissue pieces before freezing for the purpose of controlling the sectioning consistency. By choosing the proper combinations of sucrose concentration and sectioning temperature, a wide variety of tissues can be smoothly sectioned. Isolated cells, suspended in a sucrose solution, are sectioned by sectioning the frozen droplet of the suspension. A small liquid droplet of a saturated or near-saturated sucrose solution, suspended on the tip of an eyelash probe, is used to transfer frozen sections from the knife edge onto a grid substrate or a water surface. Upon melting of the sections on the surface of the sucrose droplet, they are spread flat and smooth due to surface tension. When the section of a suspension of single cells melts, individual sections of cells remain confined to the small area of the droplet surface. These devices make it possible to cut wide dry sections, and to avoid flotation on dimethyl sulfoxide solutions. With appropriate staining procedures, well-preserved ultrastructural detail can be observed. The technique is illustrated with a number of tissue preparations and with suspensions of erythrocytes and bacterial cells.     

Notes on Technic: “Clearing” Steel Knife Epon Sections in a Polystyrene Film Sandwich

Serial sectioning epoxy embedments by steel knife permits rapid light microscope survey of large tissue volumes, and preselection of areas of interest for electron microscopy. Acetate film (Hollander 1970) and Turtox plastic slides (West 1972) have been suggested as substrates upon which the sections may be “cleared” with an added layer of cured epoxy. In our experience, these substrates are excessively adherent to Epon, and “cleared” sections thinner than 40-50 μm cannot be released from them reliably. The following method is suitable for processing Epon sections 10 or more microns thick.     

Direct Staining of Reticular Fibers with Gold Chloride

Reticular fibers are selectively stained in paraffin sections of formalin-fixed or Bouin's-fixed tissue as follows: 1% aqueous solution of gold chloride for 20 min, followed by a 10 min immersion in an aqueous solution containing 5% Na₂CO₃ and 0.5% KOH. The sections then are placed in a 5% aqueous solution of KI for 2 min. Counterstaining with a 0.25% aqueous solution of methylene blue chloride is optional. The reticular fibers stain dark pink; the collagen bundles are a light pink to straw color without the counterstain, or a light blue color when the methylene blue is used.     

Novel Fixation of Plant Tissue, Staining through Paraffin with Alcian Blue and Hematoxylin, and Improved Slide Preparation

Onion (Allium cepa) root tips were fixed in a proprietary solution without aldehyde, toxic metals or acetic acid. Fixed specimens were embedded in paraffin, sectioned on a rotary microtome and mounted on detergent-washed slides without adhesive. Slides with ribbon segments affixed were immersed in 0.2% aqueous alcian blue 8GX in screw-capped Coplin jars in a water bath at 50 C for 1 hr. Excess alcian blue was rinsed off under cold running tap water and the slides were immersed in quick-mixed hematoxylin at room temperature for 15 min. Stained slides were deparaffinized, rinsed with isopropanol, air dried, and coverslips were affixed with resin. Thus, the traditional paraffin microtechnique has been modified at all steps from fixation to finishing slides with coverslips.     

Oriented Embedding of Biological Materials and Accurate Localization for Ultrathin Sectioning

Gelatin capsules with rounded ends clipped off and open ends moistened, affixed to a glass slide and sealed with a 15% gelatin solution are used to embed blocks of tissue in plastic. The surface of the slide serves as an orientation plane for structures of the tissue. The plane end of capsules of polymerized plastic containing no tissue is used in embedding frozen tissue sections. The plastic-infiltrated section is flattened against the capsule end under the weight of a 3/4 inch square of plate glass so that larger sections may be cut and surveyed. Embedding cultured cell monolayers grown on coverslips is accomplished in a comparable manner, but the square of plate glass is not needed as a weight. Block-face localization methods depend on the type of material embedded. With blocks of tissue it is achieved by moistening the face with xylene to develop relief. Thin tissue sections are examined by transmitted light, while cell monolayers are stained on the capsule end with methylene blue.     

Preparing Epoxy Sections of Insect Tissues for Light Microscopy

Sections of aldehyde-fixed and osmium-stained insect tissues embedded in various epoxy resins were affixed to glass slides by use of a slide cover and hotplate combination. A high concentration of solvent vapor over the sections was thus maintained while they dried down on the slides, resulting in excellent flatness and adhesion. Sections were then stained at an elevated temperature with a mixture of equal parts of 3 dye solutions: 1% toluidine blue O, 1% safranin O, and saturated auramine O, all made up in 1% solution of borax in water. The method resulted in excellent differentiation of all insect tissue components including lightly chitinized structures.     

A rapid method for detecting malformations in rat fetuses

A rapid method for examining rat fetuses is presented. The technique consists of fixing the fetuses in Bouin's solution, serially sectioning the head, neck and lower trunk with a razor blade and doing sagittal sections of the heart after opening the thoracic cavity. Examples of sections from normal 20 day rat fetuses are given as well as some with the following abnormalities: cleft palate produced by chlorcyclizine and eye and heart malformations resulting from anti-adult rat kidney serum.     

The Use of a Chrome Alum-Gelatin (Subbing) Solution as a General Adhesive for Paraffin Sections

The adhesion obtained from a chrome alum-gelatin solution has been found far superior to results given by widely used general adhesives (Haupt's gelatin and Mayer's egg albumen) for paraffin sections. The subbing solution, which consists of 5.0 gin gelatin and 0.5 gm chrome alum per liter of water, is easier to apply and gives more consistent results. Sections affixed to subbed slides are resistant to removal by acids and bases: 1.0 and 0.1 N HCl or H₂SO₄, 1 M H₃PO₄, 5% oxalic and trichloroacetic acids, 1% and 10% lactic acid, 1.0 and 0.1 N NaOH or NH₄OH, and other fluids and solutions such as organic solvents, water, hypochlorite, KMnO₄ and thiosulfate. The applied adhesive is virtually unstained by many stains, including hematoxylin, eosin, fast green, safranin, PAS, Sudan IV and Mallory's triple stain. The only treatment yet found to detach affixed section in less than 6 hr is immersion in 5% trichloroacetic acid for 15 min at 100 C. The concentration of gelatin and chrome alum in the solution recommended is much lower than in previously described adhesives, but this does not seem to lessen its ability to affix the sections. If the concentrations of gelatin and chrome alum are decreased from those described, adhesive qualities are also decreased. An increase in the concentration of the ingredients causes the adhesive to become stained. The described solution therefore gives optimum adhesion and “resistance” to staining.     

Cryostat technique for central nervous system histofluorescence

Summary This paper offers a technique for obtaining monoamine histofluorescence in the CNS by means of formaldehyde perfusion followed by cryostat sectioning. No freeze-drying is involved. Cryostat sections are exposed to formaldehyde vapor to complete the fluorophore formation. The fluorescence thus obtained is bright, well localized, and does not require loading the animals with precursors. The anatomical distribution of the pathways is identical to that obtained with the classical technique. Furthermore, the fluorescence is reversible by sodium borohydride, and exhibits the expected changes in intensity with pharmacological manipulations. The sections can be exposed to a cold aqueous medium for as long as 15 min with minimal diffusion of fluorophore; this suggests potential for combining monoamine histofluorescence with other visualization techniques.     

Cryostat technique for central nervous system histofluorescence.

This paper offers a technique for obtaining monoamine histofluorescence in the CNS by means of formaldehyde perfusion followed by cryostat sectioning. No freeze-drying is involved. Cryostat sections are exposed to formaldehyde vapor to complete the fluorophore formation. The fluorescence thus obtained is bright, well localized, and does not require loading the animals with precursors. The anatomical distribution of the pathways is identical to that obtained with the classical technique. Furthermore, the fluorescence is reversible by sodium borohydride, and exhibits the expected changes in intensity with pharmacological manipulations. The sections can be exposed to a cold aqueous medium for as long as 15 min with minimal diffusion of fluorophore; this suggests potential for combining monoamine histofluorescence with other visualization techniques.     

The Mounting of Carbowax Sections with Lighter-Fluid and Rubber Cement

Serial sections cut from plant tissues embedded in Carbowax have been affixed to slides with rubber cement. A rather thick layer of undiluted rubber cement was first spread on the slides. The Carbowax ribbons were added next. Lighter-fluid, essentially petroleum ether which can be substituted for it, was then run under the sections to dissolve the rubber cement and to float the ribbons. This notation medium did not dissolve the Carbowax and the ribbons could be manipulated in it for accurate location. The slides were dried on a 45° C warming table which also helped to flatten the sections. Adhesion was best when drying times were held to 4 hr or less. All excess rubber cement was washed away with xylene immediately prior to covering and the cover slips were carefully applied with a very thin resinous mounting medium to prevent dislodging the sections. Both aqueous and alcoholic stains have been used successfully and the slides have been left in them for as long as 3 days without loss of sections. The method was developed for fluorescence microscopy but serves equally well for visible light microscopy. Slides stained with a safranin-fast green combination have been used for both purposes, the safranin staining and fluorescing in a manner similar to rhodamine B.     

Block-Surface Staining

A method is described by which the tissue exposed on sectioning a specimen embedded in paraffin can be visualized in situ. The fixed specimen is impregnated with lead acetate, dehydrated in dioxane, infiltrated with paraffin and embedded. Tissues exposed on sectioning are developed by applying to the cut surface of the block a solution of potassium sulphide in water. Concentrations of the reagents used and the time intervals for the procedure are dependent upon the size of the specimen and upon the degree of contrast required. The method is described as it was applied to the study of a small human fetus in cross section. Representative photographs are included to show the results obtained.     

On the preparation of cryosections for immunocytochemistry

The key preparation steps in the Tokuyasu thawed frozen section technique for immunocytochemistry, namely freezing, sectioning, thawing, and drying, were studied. A spherical tissue culture cell was used as a model system. The frozen hydrated section technique indicated that glutaraldehyde-fixed, 2.1 M sucrose-infused pellets of cells were routinely vitrified by immersion in liquid nitrogen but water was crystallized when lower sucrose concentrations (0.6-1 M) were used. Quantitative mass measurements showed that the fixed cells are freely permeable to sucrose. The frozen hydrated sections were severely compressed but cell profiles regained their circular appearance upon thawing. The average section thickness of our frozen-hydrated sections was 110 nm: this was reduced to 30-50 nm upon thawing, washing, and air-drying. This change was accompanied by severe drying artifacts. By using the methyl cellulose drying technique, this collapse upon air-drying could be significantly reduced, but not completely prevented, giving an average thickness of 70 nm.     

An evaluation of the form of vacuoles in thin sections and freeze-etch replicas of root tips

Summary A comparison is made of the form of vacuoles in thin sections and freeze-etch replicas of root tips. In sections, vacuoles exhibit a diversity of shapes, the greatest irregularity being found with fixation in aqueous KMnO₄. Vacuoles of frozen-etched roots are mainly spherical. They are not found with narrow extensions or angular irregularities but retain a turgid appearance with a smoothly contoured tonoplast, except in some prefixed and poorly frozen fresh cells. As freeze-etching avoids artifacts of sectioning techniques it is considered that results obtained from freeze-etching give a more accurate picture of the shape of vacuoles. The irregular shapes of vacuoles in thin sections are apparently caused by shrinkage during fixation. When shrinkage is severe, portions of the tonoplast become apposed and superficially resemble profiles of endoplasmic reticulum.