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.     

The Use of Hydrophobic Adhesive Tape to Produce Miniature Wells on Microscope Slides

Hydrophobic adhesive tape was used to produce miniature wells on microscope slides for staining several sections of tissue with minimal amounts of cytochemical reagents. The wells could be tailored to individual specifications and the method allowed coverslips to be mounted close to the sections using either aqueous or xylene based mounting media. This method was especially useful for multiple immunolabelling of serial semithin cryosections.     

The Benzidine Staining Method for Blood Vessels

Two modifications of the method are described: A. Living specimens of sabellid and serpluid polychaetes, earthworms, small tadpoles, or fish larvae are immersed in an approximately saturated solution of benzidine for 30 minutes and then 3% hydrogen peroxide is added until bubbles of gas appear. When the blood vessels appear dark blue, the specimens are fixed in acidified 70% alcohol, dehydrated, cleared and either mounted in Canada balsam as whole mounts, or embedded in paraffin, sectioned at 100 to 250µ and mounted. B. Material fixed in 10% formalin in sea-water, or in formalin hypertonic saline, is incubated at 37°C. for one hour in an aqueous mixture containing sodium nitroprusside, 0.1%; benzidine, acetic acid 0.5%, followed by a weak (0.01–0.02%) hydrogen peroxide solution for a further hour, embedded in paraffin, cut into thick sections and mounted.     

The Benzidine Staining Method for Blood Vessels

Two modifications of the method are described: A. Living specimens of sabellid and serpluid polychaetes, earthworms, small tadpoles, or fish larvae are immersed in an approximately saturated solution of benzidine for 30 minutes and then 3% hydrogen peroxide is added until bubbles of gas appear. When the blood vessels appear dark blue, the specimens are fixed in acidified 70% alcohol, dehydrated, cleared and either mounted in Canada balsam as whole mounts, or embedded in paraffin, sectioned at 100 to 250µ and mounted. B. Material fixed in 10% formalin in sea-water, or in formalin hypertonic saline, is incubated at 37°C. for one hour in an aqueous mixture containing sodium nitroprusside, 0.1%; benzidine, acetic acid 0.5%, followed by a weak (0.01-0.02%) hydrogen peroxide solution for a further hour, embedded in paraffin, cut into thick sections and mounted.     

A One-Solution Stain for Spermatozoa

Fresh semen is allowed to liquefy 30-60 minutes and thin, even smears of it made on clean slides or cover glasses. The smears are fixed 3 minutes with an equal-parts mixture of alcohol and ether, then air dried. They are stained 5-7 minutes in an aqueous solution made by mixing 2 volumes of 5% aniline blue (water soluble), 1 volume of 5% eosin B and 1 volume of 1% phenol. Staining at 40-60°C. is recommended. After staining, the smears are washed with distilled water, air dried and mounted in balsam or synthetic resin. The method was used on over 2000 samples of dog semen and some human specimens. Good preservation and differentiation of cytological structures was obtained uniformly, but tests were not made with other species.     

Improved transmission electron microscopy technique for the study of cytologic material

A modification of the technique of Coleman et al for the preparation of single cells in cytologic specimens for electron microscopy (EM) is described. By employing materials in the initial cytologic processing that are useful for EM, such as a paraformaldehyde-glutaraldehyde fixative, lactated Ringer's solution as a rinsing medium, glycerol as a mounting medium and cacodylate buffer for removal of coverslips, the use of alcohol fixatives and standard mounting media could be avoided. This preserved the cytoplasmic detail, which is usually degenerated in cells removed from cytologic specimens and processed for EM.     

Evaluation of LR white resin for histology of the undecalcified rat tibia

Histology of plastic embedded undecalcified bone represents a challenging problem to the histotechnologist. We outline here an exploration of LR White resin as a suitable medium for histologic study of undecalcified rat tibia. A procedure was developed for light microscopy of rat tibia embedded in LR White and sectioned by sawing-grinding technics. The specimens were fixed in 10% neutral buffered formalin or alcohol-acetic acid-formol, dehydrated in ethanol, defatted in chloroform followed by resin infiltration and heat-curing of embedded blocks. The procedure of dehydration, defatting, infiltration, and polymerization can be completed within 10 days. Cold curing with accelerator provided by the manufacturer did not yield superior results compared to blocks cured with heat. Thick sections were obtained using a diamond wire saw, attached to plexiform slides, then ground and polished. Surface staining with Von Kossa silver reagent or toluidine blue revealed satisfactory morphological preservation of the mineralized bone sections. Artifacts like small bubbles appeared occasionally and could not be avoided despite prolonged infiltration or cold curing of blocks. Our method is relatively simple for base-line histologic study of rat tibia. The method offers advantages such as easy adaptability, reliable stainability, contrast, and resolution of bone architecture and marrow cells. Two other embedding media, Micro-Bed resin and Unicryl, were also tested, but produced inferior results.     

False Cellular Localization of Aminopeptidase Caused by Resinous Mounting Media

Fresh frozen sections of rat submandibular gland were processed for leucine aminopeptidase localization with L-lencyl-β-naphthylamide hydrochloride and L-leucyl-β-naphthylamide. When the first substrate was utilized stained sections were dehydrated in an ethyl alcohol series, cleared in xylene, and mounted in water-insoluble (resinous) media. Sections were also removed at each stage of dehydration and cleared and mounted with glychrogel. When the second substrate was used, tissues were partially decolorized in 40% ethyl alcohol and mounted directly in glychrogel. Comparison of all sections mounted in glychrogel indicated that there was no variation in cellular localization, regardless of substrate used or degree of dehydration. Nuclei were unstained. After mounting in balsam or synthetic resin the nuclei exhibited an intense stain and the parenchymal reaction was stronger, but diffuse. Progressive staining of the nuclei was observed, microscopically, immediately after applying the resinous mounting medium. The use of an aqueous mounting medium appears to be mandatory in this procedure—glychrogel is recommended.     

Enzyme histochemistry on freeze-dried, resin-embedded tissue

We have developed a method for histochemical demonstration of a wide range of enzymes in freeze-dried, resin-embedded tissue. Freeze-dried tissue specimens were embedded without fixation at low temperature (4 degrees C or -20 degrees C) in glycol methacrylate resin or LR Gold resin. Enzyme activity was optimally preserved by embedding the freeze-dried tissue in glycol methacrylate resin. All enzymes studied (oxidoreductases, esterases, peptidases, and phosphatases), except for glucose-6-phosphatase, were readily demonstrated. The enzymes displayed high activity and were accurately localized without diffusion when tissue sections were incubated in aqueous media, addition of colloid stabilizers to the incubating media not being required. Freeze-drying combined with low-temperature resin embedding permits the demonstration of a wide range of enzymes with accurate enzyme localization, high enzyme activity, and excellent tissue morphology.     

Sandwich Embedding of Monolayers of Cells in Epoxy Resin for Ultrathin Sectioning

In this procedure for embedding monolayers of cells, the usual glass slides are replaced by plates of resin 1-1.5 mm thick. Unlike the open-face embedding technique, the present procedure uses only a few drops of unpolymerized resin, which are applied to the fixed and dehydrated cells. During polymerization this small amount of liquid resin spreads across a relative large area, leaves the cells covered by a very thin layer, and permits phase contrast observations through it. Ultrathin sections of a particular cell encircled by a rotary scriber can be obtained by sectioning the resin slide, which has been trimmed and mounted directly in the specimen holder of the ultramicrotome.     

Observation on Rice Embryo Sac Development with Confocal Laser Scanning Microscopy

An easy and rapid procedure has been launched for observafion of embryo sac development of Oryza sativa L. Following fixation and dehydration, whole mounted ovaries were cleared in methyl sulieylate, finally sealed under coverslips with clove oil. With the confocal laser scanmng microscopy procedure, the embryo sac exhibited autofluorescence and could be clearly observed. FAA and 4% glutaraldehyde as fixative produced different results but the latter seemed to be better.     

Epidermal and Cuticular Mounts of Plant Material Obtained by Maceration

Mounts of leaf and stem epidermises or bare cuticles, useful in both general anatomical and specialized phylogenetic studies, can be prepared by a maceration process using Jeffrey's solution (equal volumes of 10% aqueous CrO₃ and 10% HNO₃). Leaves, including those of conifers, and stems with cuticles thick enough to maintain integrity when isolated are amenable to this process. Dried specimens are hydrated by boiling in water; fluid-preserved specimens are washed thoroughly in water; fresh specimens need no pretreatment. Specimens are cut to a convenient mount size and trimmed so as to allow adequate and even penetration of the macerating fluid. Laminar leaf segments are left with one edge untrimmed so that upper and lower epidermises remain contiguous. Cylindrical leaf and stem segments are slit lengthwise through about half their thickness. Specimens are macerated in Jeffrey's solution for 1 to 4 days until unwanted tissues are loosened and easily freed from the epidermis (or bare cuticle, if that is desired). The macerating fluid is then washed out completely with changes of water and specimens are stained in a 0.5% aqueous solution of safranin. Dehydration is accomplished with several changes of tertiary butanol. All unwanted tissues not removed by agitation during previous steps of the process are removed by teasing prior to mounting. Specimens are then mounted on slides in Canada balsam and dried in the usual manner.     

On the status of Eimeria nieschulzi oocysts embedded in resin eleven years ago: a permanent method for preserving coccidian oocysts

Sporulated oocysts of Eimeria nieschulzi that were fixed and mounted on glass slides in polymerized resin in 1976 are examined. Size, shape, and integrity of oocysts and sporocysts are compared to similar observations we made in 1977 and reported in 1978 (Journal of Parasitology 64: 163-164). Our conclusion is that the methods we reported on in 1978 provide one opportunity to produce permanent specimens of sporulated oocysts that could be made available for deposit in nationally accredited museums.     

A Rigid Illuminator Giving Transmitted Light to Facilitate Microhardness Testing of Calcified Tissues

The structural form of calcified tissues necessitates their examination with transmitted light to identify various morphological features which are not evident with incident illumination alone. To achieve stability for indentation, the specimens must be rigidly mounted. A Leitz Model 514095 base illuminator was fitted with a bottom plate of 3 mm brass for attaching it to the graduated stage and a perforated, 3 mm thick brass top plate, surmounted by a 6 mm thick piece of plate glass. The 25 mm hole in the top plate coincided with a ground glass disc to transmit diffused light through the plate glass to the specimen. Thus, the specimen could be examined on a rigid microscope stage, morphological features identified, and subsequent interpretation of indents made with the usual incident illumination.     

The Use of Snail Gut Cytase in Fluorescent Banding Studies on Plant Chromosomes

The fluorescent bands produced in chromosomes by quinacrine derivatives, although highly acid-labile, have been shown to be resistant to digestion by snail gut cytase. This enzyme may therefore be used to soften plant root-tips and so facilitate the production of flat, unbroken mitotic-metaphase plates for fluorescent-banding studies. The roots are fixed in 3:1 absolute alcohol:glacial acetic acid then thoroughly washed in water. The washed roots are then dipped in undiluted cytase and digested for either 2 hrs at room temperature or overnight in a domestic refrigerator. Next they are squashed on a slide and stained in 0.5% aqueous quinacrine hydrochloride for 10-15 minutes at pH 6.2. After washing in three changes of water they are mounted in water. The coverslips are sealed on with clear nail varnish.     

Dehydrogenase enzyme histochemistry on freezedried or fixed resin-embedded tissue

Summary A method has been developed for the histochemical demonstration of a variety of dehydrogenases in freeze-dried or fixed resin-embedded tissue. Seven dehydrogenases were studied. Lactate dehydrogenase, NADH dehydrogenase and NADPH tetrazolium reductase were all demonstrable in sections of paraformaldehyde-fixed resin-embedded tissue. Freeze-dried specimens were embedded, without fixation, in glycol methacrylate resin or LR Gold resin at either 4°C or –20°C. All the dehydrogenases except succinate dehydrogenase retained their activity in freeze-dried, resin-embedded tissue. Enzyme activity was maximally preserved by embedding the freeze-dried tissue specimens in glycol methacrylate resin at –20°C. The dehydrogenases were accurately localized without any diffusion when the tissue sections were incubated in aqueous media. Addition of a colloid stabilizer to the incubating medium was not required. Freeze-drying combined with low-temperature resin embedding permits accurate enzyme localization without diffusion, maintenance of enzyme activity and excellent tissue morphology.     

Effect of combined recompression and air, oxygen, or heliox breathing on air bubbles in rat tissues.

The fate of bubbles formed in tissues during the ascent from a real or simulated air dive and subjected to therapeutic recompression has only been indirectly inferred from theoretical modeling and clinical observations. We visually followed the resolution of micro air bubbles injected into adipose tissue, spinal white matter, muscle, and tendon of anesthetized rats recompressed to and held at 284 kPa while rats breathed air, oxygen, heliox 80:20, or heliox 50:50. The rats underwent a prolonged hyperbaric air exposure before bubble injection and recompression. In all tissues, bubbles disappeared faster during breathing of oxygen or heliox mixtures than during air breathing. In some of the experiments, oxygen breathing caused a transient growth of the bubbles. In spinal white matter, heliox 50:50 or oxygen breathing resulted in significantly faster bubble resolution than did heliox 80:20 breathing. In conclusion, air bubbles in lipid and aqueous tissues shrink and disappear faster during recompression during breathing of heliox mixtures or oxygen compared with air breathing. The clinical implication of these findings might be that heliox 50:50 is the mixture of choice for the treatment of decompression sickness.     

Transmission and scanning electron microscopy of cell monolayers grown on polymethylpentene coverslips.

Plastic coverslips made of polymethylpentene serve as excellent substrates for growth of bovine endothelial cells, and are easily processed for both transmission (TEM) and scanning (SEM) electron microscopy. Portions of the same coverslip (monolayer) are used for both SEM and TEM examination and are fixed, postfixed, and dehydrated as a single entity. The portion of the coverslip for SEM is then excised, critical point dried, and mounted for sputter coating prior to viewing. The remaining piece of coverslip used for TEM is Epon-Araldite embedded, polymerized, separated from the coverslip by liquid nitrogen immersion, and sectioned either "en face" or in cross section for viewing. Coated glass coverslips are not required and organic solvents such as propylene oxide, acetone, and amyl acetate can be used for dehydration and infiltration. Furthermore, specimens do not require re-embedding or blocks to be glued onto blank capsules before sectioning. The number of cells needed to achieve a monolayer is significantly reduced compared to the usual culture flasks, but are abundant enough to assess ultrastructural changes accurately. Support films may be required to prevent folding of the ultrathin section which can obstruct viewing of cells located on the edge of the section.     

A Simple Toluidine Blue—Basic Fuchsin Stain for Spermatozoa in Epoxy Sections

Differential staining of cell components of spermatozoa is readily accomplished in Epon or Araldite sections 0.5-1 μ thick from rat and hamster testis and epididymis, and stained as follows: 1% aqueous toluidine blue buffered at pH 6, 0.5-3 min at 90 C; washed in distilled water; 1% basic fuchsin in 50% alcohol, 3-5 min at 20-25 C; differentiated with 70% alcohol; allowed to dry; and mounted in a resin of high refraction (DPX was used). Results: acrosome, bright magenta; nucleus, deep blue; mitochondrial sheath of the middle-piece, pinkish purple; and tail, pale red. This procedure combined with staining of collagen by applying 2% aqueous phosphotungstic acid 1-2 min as a mordant, followed by 1% light green in 50% alcohol containing 1% acetic acid, 1-2 min at 20-25 C, gives polychromatic staining and is useful as a general stain for other epoxy-embedded tissues.     

Transmission and Scanning Electron Microscopy of Cell Monolayers Grown on Polymethylpentene Coverslips

Plastic coverslips made of polymethylpentene serve as excellent substrates for growth of bovine endothelial cells, and are easily processed for both transmission (TEM) and scanning (SEM) electron microscopy. Portions of the same coverslip (monolayer) are used for both SEM and TEM examination and are fixed, postfixed, and dehydrated as a single entity. The portion of the coverslip for SEM is then excised, critical point dried, and mounted for sputter coating prior to viewing. The remaining piece of coverslip used for TEM is Epon-Araldite embedded, polymerized. separated from the coverslip by liquid nitrogen immersion, and sectioned either “en face” or in cross section for viewing. Coated glass coverslips are not required and organic solvents such as propylene oxide, acetone, and amyl acetate can be used for dehydration and infiltration. Furthermore, specimens do not require re-embedding or blocks to be glued onto blank capsules before sectioning. The number of cells needed to achieve a monolayer is significantly reduced compared to the usual culture flasks, but are abundant enough to assess ultrastructural changes accurately. Support films may be required to prevent folding of the ultrathin section which can obstruct viewing of cells located on the edge of the section.