A rapid technique for preparing microorganisms for transmission electron microscopy.

A rapid and efficient method of preparing microorganisms for transmission electron microscopy is reported. In developing the method Salmonella, streptococcal, and protozoal specimens were fixed with glutaraldehyde. After fixation cells are collected on a membrane filter, washed with buffer, postfixed with osmium tetroxide, then washed with distilled water and stained en bloc with uranyl acetate. Specimens are dehydrated using a graded series of acetone and then infiltrated with graded mixtures of acetone and Spurr embedding medium. Finally the membrane filter is cut into small pieces and embedded in fresh embedding medium polymerized in polyethylene capsules. By collecting and processing the specimens on membrane filters, numerous centrifugations are eliminated from standard procedures. The use of a low viscosity embedding medium allows for rapid infiltration and embedding of the specimen. Using this technique microbial specimens can be sectioned after less than 4 hours preparation.     

A Method for the Preparation of Undecalcified Bone Sections for Light Microscopy and Microradiography

A method which gives good quality 1-2 μm thick sections of undecaldfied cancellous and thin cortical bones for light miuoscopy is described. Formalin fixed material is dehydrated in graded acetones and embedded in a modiEed formula of Spurr's low viscosity embedding medium. After a 16 hour polymerisation period at 60 C, sections are cut at 1-2 μm thickness on a Porter-Blum JB4A rotary microtome Using glass knives. Sections are attached to clean glass slides with heat, the resin degraded in bromine vapour and removed in acetone. This allows comparative ease of staining. The technique is rapid, does not interfere with tetracycline fluorescence and the same specimens can be used to prepare thick sections for microradiography.     

In Situ Multiple Sampling of Attached Bacteria for Scanning and Transmission Electron Microscopy

An in situ electron microscope sampling technique for characterizing cells attached to smooth surfaces is demonstrated with an ultraviolet-induced mutant of Streptococcus mutans. The sterilized sampling unit consists of a 9 cm plastic Petri dish containing a glass slide, a 12 mm round coverglass, and a coverglass with Formvar-carbon coated copper grids. After the bacterial culture in a liquid medium is incubated in the Petri dish, the slide with attached bacteria is washed in double-distilled water, air-dried, coated with platinum and carbon, and processed for replicas and shadowed specimens for transmission electron microscopy. The coverglass is similarly washed, fixed in 2% glutaraldehyde, air- or freeze-dried, coated with palladium/gold, and examined in the scanning electron microscope. The coverglass with grids is rinsed in double distilled water, the grids are transferred to a filter paper and stained with a loopful of 2% phosphotungstic acid at pH 5.5. The bacteria growing on the surface of the plastic Petri dish are fixed, dehydrated, and embedded in situ with Epon. Sectioned and stained specimens are then examined in the transmission electron microscope. This procedure also appears useful with such other attached systems as normal or infected tissue culture cells.     

A Simple Stain for Myelin in Frozen Sections: A Modification of Mahon's Method

A simple and rapid method for demonstrating myelinated nerve fibers in frozen sections of the central and peripheral nervous system is described. Material fixed by perfusion with mixed aldehydes gives the best results but the method also works on specimens fixed by immersion in formaldehyde. Frozen sections varying in thickness from 15-50 μm are mounted on slides subbed with chrome alum-gelatin. After hydration (60-140 min), Sections are mordanted (20-40 min) in 2.5% iron alum and rinsed briefly in three changes of distilled H₂O (total 2 min). Staining is for 60-180 min in 40 cc freshly made 10% alcoholic hematoxylin diluted with 165 cc distilled H₂O to which 15 cc saturated Li₂CO₂is added. the sections are washed in distilled H₂O (5-15 min) and dehydrated in graded alcohols without differentiation in mordant, and covered. Myelin stains a dark blue-purple against a light grey background. Fiber tracts, as well as individual myelinated fibers, are clearly demonstrated.     

Histochemical Demonstration of Acid Phosphatase in Hard Tissues

The action of the following decalcifying solutions for the demonstration of acid phosphatase has been studied: buffer solution acetic-acetate 0.05 M, pH 5; 2, 5, 10, 20, and 50% formic acid and 20% sodium citrate in equal parts (pH 2.6, 3, 3.8, 4.2, and 5); 0.5 M citric acid-NaOH, pH 4.2; Versene solution, 5%, pH 7. A comparative study of fixatives has been made also (neutral formalin, 10-20%; formalin-chloral hydrate (Fishman), acetone and 80% alcohol). The best results were obtained with fixation at 4°C in 10-20% neutral formalin or formalin-chloral hydrate, for a period of 24 hr, and decalcification with 20% sodium citrate, 5% formic acid, in equal parts, pH 4.2, which can act on both specimens or sections for a period up to 2 wk with very little loss of enzyme. It is not necessary to reactivate the enzyme after decalcification; frozen sections should be used and should be washed in distilled water before proceeding with the demonstration of the enzyme (Gomori's method or azo dye coupling). Other fixatives (acetone and alcohol) and paraffin embedding produce a greater loss in enzymes and very irregular results.     

Use of Hot Formaldehyde Fixative in Processing Plant-Parasitic Nematodes for Electron Microscopy

A preparative technique is formulated for processing plant-parasitic nematodes of the order Tylenchida for electron microscopy. A population of Dolichodorus heterocephalus is used as test objects. One and a half grams of paraformaldehyde are dissolved in 25 ml of water at 60 C. Five drops of 1 N sodium hydroxide are added to clear the solution, which is then cooled to room temperature. Two and a half milliliters of 25% glutaraldehyde are added with 23 ml 0.1 M phosphate buffer, pH 7.3, and 0.2 M with respect to sucrose. The final solution contains 3% formaldehyde and 1% glutaraldehyde and is pH 7.2. It is heated to 70 C, poured over specimens, and allowed to cool to 4 C in 2 hr. The nematodes are then incised in a fixative containing 2% glutaraldehyde and 5% dimethyl sulfoxide at 4 C for 16-24 br. Five milliliters of 25% glutaraldehyde and 2.5 ml of dimethyl sulfoxide are combined in 17.5 ml of water. Twenty-five milliliters of phosphate buffer (supplemented as above) are added. The final pH is 7.2. The glutaraldehyde, aided by dimethyl sulfoxide, uniformly and permanently fixes the nematode tissues. The specimens are embedded in agar. Following a 30-min buffer wash (4 C) they are postfixed in buffered 2% osmium tetroside for 2 hr at room temperature, washed, and dehydrated through an ethanol series and two acetone baths. Dehydration includes a 2-hr stop in 75% ethanol containing 2% uranyl acetate. After embedding in Spurr's epoxy resin, specimens are sectioned and poststained in 0.5% aqueous uranyl acetate for 6 min and saturated aqueous lead citrate 3-4 min.

This technique reduces killing time to less than 2 sec, straightens specimens for easier orientation, and eliminates the typically high internal pressure of nematodes which causes displacement of internal structures observed with other fixation techniques.     

Histochemical detection of horseradish peroxidase activity in the rat by the vibratome-paraplast method

A method is described for the histochemical detection of horseradish peroxidase in Paraplast Plus embedded brain sections. The procedure uses 150-micron-thick Vibratome-cut slices of glutaraldehyde-paraformaldehyde-fixed brain tissue. Tetramethylbenzidine stabilized by diaminobenzidine/cobalt/H2O2 is used as chromogen. The Vibratome-cut slices are dehydrated through a graded series of acetone, cleared in toluol and flat-embedded in Paraplast Plus embedding medium. Serial sections can be cut as thin as 5-7 micron. The method is universal in its application and permits optimal visualization of labeled neurons with great morphological detail at the light-microscopic level.     

Microcount Method for Petrolatum-Based Topical Ointments Containing Waxes

A practical solvent system for the detection of microorganisms in topical ointments has been developed. The method involves dissolving 0.5 g of topical ointment in 50 ml of a solvent mixture (92 parts isopropyl myristate, 6 parts carbon disulfide, and 2 parts xylene) and filtering it through a 0.45-mum membrane filter. Residual solvent is then washed from the filter pad with 200 ml of sterile 0.5% Brain Heart Infusion broth containing 0.1% Tween 80. The filter pad is then removed and placed on a petri plate containing Trypticase Soy Agar medium. The petri plates thus prepared are then incubated at 37 C for 7 days, and the colonies produced are then counted. The toxicity of the solvent mixture was determined against Staphylococcus aureus, Escherichia coli, Pseudomonas aeruginosa, Salmonella newington, and spores of Bacillus subtilis and was found to be less toxic than the heat-sterilized isopropyl myristate and comparable to the filter-sterilized isopropyl myristate.     

Sample preparation and digestion for proteomic analyses using spin filters

We describe the use of commercially available microcentrifugation devices (spin filters) for cleanup and digestion of protein samples for mass spectrometry analyses. The protein sample is added to the upper chamber of a spin filter with a > or = 3000 molecular weight cutoff membrane and then washed prior to resuspension in ammonium bicarbonate. The protein is then reduced, alkylated, and digested with trypsin in the upper chamber and the peptides are recovered by centrifugation through the membrane. The method provides digestion efficiencies comparable to standard in-solution digests, avoids lengthy dialysis steps, and allows rapid cleanup of samples containing salts, some detergents, and acidic or basic buffers.     

Epoxy resin as fixative during freeze-substitution

An alternative protocol for freeze-substitution is described. Araldite/Epon embedding medium (20% in acetone) is first used as a stabilizer (as e.g., OsO(4)) and then as embedding medium. The major components of the Araldite/Epon resin formulation react with proteins and lipids and provide for an excellent preservation and reasonable visualisation of the ultrastructure. The ultrastructural appearance can be deliberately influenced with the standard freeze-substitution procedure [Van Harreveld, A., Crowell, J., 1964. Electron microscopy after rapid freezing on a metal surface and substitution fixation. Anat. Rec. 149, 381-386.] using OsO(4) as stabilizing agent by protocols which degrade cytoplasmic and membrane proteins. Epoxy stabilized and embedded samples may become an important tool to get information about the effects of different reagents and protocols used in freeze-substitution. We believe that an in-depth understanding of the procedures is required to correctly interpret images and to complement studies of dynamic processes by light microscopy with reliable, highly detailed ultrastructural information. The block face of epoxy stabilized samples after ultrathin sectioning is highly suited for the analysis of the ultrastructure by AFM.     

A new method for transfer of polyethylene glycol-embedded tissue sections to silanated slides for immunocytochemistry

Polyethylene glycol (PEG) is an excellent embedding medium for immunohistochemical studies. It provides structural preservation superior to frozen sections and increased sensitivity of antigen detection compared with paraffin sections. One limitation of PEG embedment is that PEG sections are difficult to handle and adhere poorly to glass slides. Here we present a simple and effective method for embedding tissues in PEG and transferring the resultant sections onto silanated glass slides. In addition, a method for silver enhanced colloidal gold immunostaining was combined with common dye staining to demonstrate the excellent structure preservation and sensitive antigen detection. Bovine chorionic membrane was fixed with Bouin's fixative, embedded in polyethylene glycol (PEG) 1500, cut into 5-microns sections, flattened over agarose blocks (10 x 10 x 2 mm3), and blotted onto Digene silanated slides. Slides were then washed in PBS, which removed the PEG and agarose blocks. Tissue sections were immunocytochemically stained with dilute antiserum raised in a rabbit against purified bovine placental retinol binding protein (bpRBP). Sections were washed and incubated with 1-nm colloidal gold-labeled goat anti-rabbit IgG. The immunogold particles were enhanced by silver staining (IGSS). Specimens were observed and photographed with an Olympus epipolarization microscope. The new method offered excellent morphological preservation of cell structure and the epipolarization microscopy provided high sensitivity for detection of specific immunogold-silver particles.     

An Embedding Method for Monolayer Cell Cultures for Light and Electron Microscopy

Plastic containers are widely used for monolayer cell culture. In situ embedding, the obvious method of choice for subsequent ultrastructural study, has been achieved by Brinkley et al. (1967) using Epon as a final embedding medium and water soluble acrylates as intermediates. Although the results are satisfactory, the method has two drawbacks: firstly, the water soluble acrylates are difficult to get, and secondly, removal of the plastic container is possible only with blocks already cut off for electron microscopy.     

Preparation of methacrylate-embedded cytologic smears from prefixed cells

A new method of preparing smears of alcohol-fixed cytologic material by using methacrylate embedding medium to make the cells adhere on plain glass slides is presented. After centrifugation, the cytologic material was mixed with Lowicryl K4M embedding medium and smeared on slides. The polymerization process was achieved by exposing the slides to ultraviolet light. The morphology in such smears was similar to that of specimens prepared by the filter technique. The methacrylate method does not have the most common disadvantages of the filter technique--the development of air bubbles over time and the visually disturbing presence of the filter.     

Purifying Pararosanilin for Use in Colorless Schiff Reagent

Pararosanilin hydrochloride or pararosanilin base was purified by suspending 20 g. of the dye in 400 ml. of water, acidifying with 50 ml. of 2N HCl and adding 4—5 g. of decolorizing charcoal. The mixture was then heated to boiling and boiled for 2 minutes. The entire mixture was transferred to a large, covered, fluted filter, and the filtrate allowed to stand overnight while the pararosanilin hydrochloride precipitated. The pararosanilin hydrochloride was filtered off, resuspended in 100 ml. of ether-alcohol (10:1) and shaken for 3 to 5 min. The ether-alcohol suspension was filtered and the pararosanilin hydrochloride washed repeatedly on the filter with ether until the ether wash was no longer colored. It was then washed several times with distilled water (total volume 400 ml.), dried in vacuuo over concentrated sulfuric acid, ground to a fine powder and stored in a dark brown bottle.     

Lymphocyte membrane antigens in glycol methacrylate embedded tissue

The use of formalin or Michel's solution either alone or in combination with acetone, and acetone, methanol or ethanol alone as fixatives, and glycol methacrylate as embedding medium were evaluated for their suitability in procedures to detect lymphocyte membrane antigens by OKT and Leu monoclonal antibodies in human tonsils. No staining was detected in sections fixed in 70% or absolute ethanol and embedded in glycol methacrylate with either the direct immunofluorescence or avidin-biotin methods. Fixation in Michel's solutions plus acetone at room temperature revealed staining by both. Neither method resulted in staining after fixation in Michel's solution plus acetone at 4 C presumably due to the slow action of the fixative. Staining was enhanced using a combination of primary and secondary biotinylated antibodies. Dual staining allowed concurrent detection of two antigens in the same section. Glycol methacrylate embedding is a possible replacement for ultracold storage in the preservation of tissue for immunofluorescent staining.     

Lymphocyte Membrane Antigens in Glycol Methacrylate Embedded Tissue

The use of formalin or Michel's solution either alone or in combination with acetone, and acetone, methanol or ethanol alone as fixatives, and glycol methacrylate as embedding medium were evaluated for their suitability in procedures to detect lymphocyte membrane antigens by OKT and Leu monoclonal antibodies in human tonsils. No staining was detected in sections fixed in 70% or absolute ethanol and embedded in glycol methacrylate with either the direct immunofluorescence or avidin-biotin methods. Fixation in Michel's solutions plus acetone at room temperature revealed staining by both. Neither method resulted in staining after fixation in Michel's solution plus acetone at 4 C presumably due to the slow action of the fixative. Staining was enhanced using a combination of primary and secondary biotinylated antibodies. Dual staining allowed concurrent detection of two antigens in the same section. Glycol methacrylate embedding is a possible replacement for ultracold storage in the preservation of tissue for immunofluorescent staining.     

A Ribonuclease-Gallocyanin Stain for Sexing Skin Biopsies

Skin biopsies for sexing can be fixed best in 10-15% aqueous formalin or this solution saturated with HgCl₂. Bouin's fluid and all chromate mixtures should be avoided. Celloidin-paraffin double embedding is recommended but not essential. Sections are brought to water, mercurial residues removed if necessary, and then washed in distilled water. They are incubated at 37°C in a ribo-nuclease solution: approximately 1 mg of ribonuclease powder (Light's) in 100 ml of glass-distilled water; boiled 3-5 sec after dissolving, and kept in a refrigerator (usable about a week). The sections are rinsed and incubated at 37°C overnight in gallocyanin-chromalum (Einarson, 1951) made as follows: Dissolve 5 gm of chromalum in 100 ml of distilled water, add 0.15 gm of gallocyanin, shake thoroughly, heat slowly and boil 5 min; cool, filter, and wash through the filter with distilled water until the filtrate reaches 100 ml. This solution is usable at once and keeps at least a month. Sections should be dipped in acid alcohol to clean (optional), but no attempt made to differentiate them, and washed in tap water. Dehydration, clearing and covering complete the process. The method is nearly as precise as the Feulgen and more convenient and reliable for routine use on miscellaneous material.     

A Quick Preparative Method for Electron Microscopy Observations of Delicate Objects Using Alginate Embedding Medium

A quick, safe method has been devised for embedding small or fragile specimens and keeping delicate structures intact. Cells or organisms to be embedded are placed in a viscous sodium alginate solution (1-2%), which is then polymerized in 100 mM calcium chloride. The resulting gel is easily dehydrated, embedded in resin and sectioned for electron microscopy. This method, the alginate gel portion of which was originally developed for the immobilization of Euglena, allows direct observation of each element of the specimens in micrographs. If desired, the alginate can be removed after sectioning by sequestration of calcium in a 20 mM solution of sodium citrate or a 10 mM solution of EGTA. Cells and organelles in the sections respond normally to standard staining procedures.     

Direct Epoxy Embedding for Vertical Sectioning of Cells Grown as a Monolayer on Millipore Filter

Cells cultured as a monolayer on MF-Millpore GSWP. 0.22 μ pore size, filter were fixed, dehydrated, and examined by phase-contrast microscopy with the filter immersed in a 1:1 mixture of xylene and the embedding medium. The membrane was cut into 2 × 20 mm strips, and each strip which was selected for desired cells was embedded vertically in a BEEM capsule. Thus direct embedding which allowed edgewise sectioning of cells was obtained without removing them from the culturing support.     

Staining of Bacterial Colonies on the Millipore Filter to Improve Contrast

About 5 ml of 1% blue tetrazolium in 70% ethyl alcohol were poured over mature colonies of Pasteurella pestis and Malleomyces pseudomallei on Millipore filters (MF), contained in the filter holder apparatus, and allowed to drain through with the suction applied. The MF was washed with water and then covered with about 10 ml of 0.001% aqueous trypan blue and drained. This technique provided vivid white colonies sharply defined against a blue background.

Another method utilized 0.1% quinacrine-HCl (Atabrine) to stain colonies yellow and 0.05% vital red to stain the MF pink to light red.