Notes of Technic

The leaching of water-soluble and exchangeable calcium in histoautoradiog-raphy of oat tissue can be prevented by using acetone as the dehydration fluid (freeze substitution technique) and by keeping the tissue sections, while stretching on water, embedded in the methacrylate matrix. Ca⁴⁵ was either added to the mineral solution on which the oat plants were grown (75 μc), or applied on the leaf surface (8 μc). After freezing in melting isopentane, specimens of 1-2 mm dimensions are fixed for 24 hr in an acetone-OsO₄ (1%) solution at—80 C. Dehydration is obtained by transferring the material every day for 6 successive days to a fresh acetone solution at—80 C. The material is infiltrated by a three-time renewed monomer methacrylate mixture (methylmethacrylate I, butylmethacrylate 4) at—50 C. The specimens are embedded in the polymerizing methacrylate mixture at room temperature. Sections of 4-8 μ are easily cut with a rotating microtome. If the methacrylate is not removed from the sections, they can be stretched on water without leaching of calcium. The presence of methacrylate in no way hinders microscopic observation nor effective histoautoradiography.     

A Plastic Embedding Medium for Thin Sectioning in Light Microscopy

Tissue blocks with surface areas up to 2 cm² can be sectioned at 1 or 2 μ after embedding in a medium consisting of: methyl methacrylate, 27 ml; polyethylene glycol distearate MW 1540, 6 gm; dibutyl phthalate, 4 ml; and Plexiglas molding powder A-100, 9 gm (added last). The methacrylate mixture is polymerized at 50° C by benzoyl peroxide, 0.8 gm/ 100 ml of methacrylate. The polymerized matrix is transparent and the blocks can be cut on a rotary microtome with a steel knife. The plastic can be removed from sections with acetone prior to staining. Artifacts caused by embedding and sectioning are negligible     

A Tribasic Stain for Thin Sections of Plastic-Embedded, OsO4-Fixed Tissues

Thin (0.5-1 μ) sections of plastic-embedded, OsO₄-fixed tissues were attached to glass slides by heating to 70 C for 1 min. A saturated solution combining toluidine blue and malachite green was prepared in ethanol (8% of each dye) or water (4% of each dye). Methacrylate or epoxy sections were stained in the ethanol solution for 2-5 min. The water solution was more effective for some epoxy sections (10-80 min). Epoxy sections could be mordanted by 2% KMnO₄, in acetone (1 min) before use of the aqueous dye, reducing staining time to 5-10 min and improving contrast. Aqueous basic fuchsin (4%) was used as the counter-stain in all cases; staining time varied from 1-30 min depending upon the embedding medium and desired effects, methacrylate sections requiring the least time. In the completed stain, nuclei were blue to violet; erythrocytes and mitochondria, green; collagen and elastic tissue, magenta; and much and cartilage, bright cherry red. Sections were coated with an acrylic resin spray and examined or photographed with an oil-immersion lens.     

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.     

Histochemical Demonstration of Carbonic Anhydrase Activity

Fresh tissue slices fixed in chilled acetone for 1 hour and washed in distilled water for 10-30 minutes were incubated for 30-45 minutes at 37°C. in the freshly prepared incubating mixture: filtrate of a mixture of 8% sodium bicarbonate, 100 ml., and MnCl₂·4H₂O, 1 g. After washing in distilled water for 1 hour, they were dehydrated and embedded in paraffin. Sections were cut 15-20μU, deparaffinized, rinsed in absolute alcohol and placed in a 0.1% solution of potassium periodate for 48 hours at 37°C. The mounted sections were counterstained (if desired), dehydrated in alcohol, cleared in xylene (not carbol-xylene) and mounted in balsam. Many brown granules were produced on the sites of enzyme activity by this procedure. The results obtained seem to be in good agreement with previous findings by biochemical determinations.     

Autoradiography of Mobile, C14-Labeled Herbicides in Sections of Leaf Tissue

Fresh leaf tissue containing a soluble, C¹⁴-labeled herbicide was mounted in cold 1% gelatin on a holder, quick frozen in a cryostat, and cross sectioned at 16 μ with single-edge, stainless steel razor blades. The sections were transferred (without thawing) to cold (—10 C) microscope slides which had been partly covered with double-coated Scotch tape #665. The tissue was freeze-dried in a vacuum desiccator at—20 C then secured to the tape with pressure. Autoradiography was accomplished in a darkroom by covering the slides with dry, nuclear track emulsion films. These films were made by dipping 2 inch diameter wire loops into liquid emulsion, letting the film dry, and applying it by blowing it as it was placed against the tissue. After a 19 day exposure in light-tight boxes at 25-27 C the preparations were processed in the usual manner. The method-was used successfully to trace the movement of soluble, C¹⁴-labeled herbicides in leaf tissue without the loss of labeling material or artifacts caused by its diffusion. High resolution autoradiograms with low backgrounds were obtained.     

Improvements in Histological Techniques for Epoxy-Resin Embedded Bone Specimens

A procedure is presented in which some of the processing difficulties with fixation, embedding and cutting whole mouse bones and large bone pieces from other species are considered. The bone specimens are fixed in acetone or by a Karnovsky-formol-saline process which preserves intact endosteal surface-to-cortex layers. After fixation the bones are embedded in a hard mixture of epoxy resin to provide blocks with face sizes up to 3.5 × 3.0 cm. Mineralized sections are cut at 4 μm; demineralized at 3 μm. Sections are fastened to gelatin-subbed slides with pressure plates which produce flat, secure sections. After removal of the plastic, an unmodified Mayer's hematoxylin and a polychromatic eosin staining method is applied to demineralized sections, and a slightly modified method to mineralized sections.     

Differentiated Staining of Osmium Tetroxide-Fixed, Vestopal-w-Embedded Tissue in thin Sections

Tissues were fixed at 20° C for 1 hr in 1% OsO₄, buffered at pH 7.4 with veronal-acetate (Palade's fixative), soaked 5 min in the same buffer without OsO₄, then dehydrated in buffer-acetone mixtures of 30, 50, 75 and 90% acetone content, and finally in anhydrous acetone. Infiltration was accomplished through Vestopal-W-acetone mixtures of 1:3, 1:1, 3:1 to undiluted Vestopal. After polymerisation at 60° C for 24 hr, 1-2 μ sections were cut, dried on slides without adhesive, and stained by any of the following methods. (1) Mayer's acid hemalum: Flood the slides with the staining solution and allow to stand at 20°C for 2-3 hr while the water of the solution evaporates; wash in distilled water, 2 min; differentiate in 1% HCl; rinse 1-2 sec in 10% NH,OH. (2) Iron-trioxyhematein (of Hansen): Apply the staining solution as in method 1; wash 3-5 min in 5% acetic acid; restain for 1-12 hr by flooding with a mixture consisting of staining solution, 2 parts, and 1 part of a 1:1 mixture of 2% acetic acid and 2% H₂SO₄ (observe under microscope for staining intensity); wash 2 min in distilled water and 1 hr in tap water. (3) Iron-hematoxylin (Heidenhain): Mordant 6 hr in 2.5% iron-alum solution; wash 1 min in distilled water; stain in 1% or 0.5% ripened hematoxylin for 3-12 br; differentiate 8 min in 2.5%, and 15 min in 1% iron-alum solution; wash 1 hr in tap water. (4) Aceto-carmine (Schneider): Stain 12-24 hr; wash 0.5-1.0 min in distilled water. (5) Picrofuchsin: Stain 24-48 hr in 1% acid fuchsin dissolved in saturated aqueous picric acid; differentiate for only 1-2 sec in 96% ethanol. (6) Modified Giemsa: Mix 640 ml of a solution of 9.08 gm KH₂PO₄ in 1000 ml of distilled water and 360 ml of a solution of 11.88 gm Na₂HPO₄-2H₂O in 1000 ml of distilled water. Soak sections in this buffer, 12 hr. Dissolve 1.0 gm of azur I in 125 ml of boiling distilled water; add 0.5 gm of methylene blue; filter and add hot distilled water until a volume of 250 ml is reached (solution “AM”). Dissolve 1.5 gm of eosin, yellowish, in 250 ml of hot distilled water; filter (solution “E”). Mix 1.5 ml of “AM” in 100 ml of buffer with 3 ml of “E” in 100 ml of buffer. Stain 12-24 hr. Differentiate 3 sec in 25 ml methyl benzoate in 75 ml dioxane; 3 sec in 35 ml methyl benzoate in 65 ml acetone; 3 sec in 30 ml acetone in 70 ml methyl benzoate; and 3 sec in 5 ml acetone in 95 ml methyl benzoate. Dehydrated sections may be covered in a neutral synthetic resin (Caedax was used).     

Embedding in Epoxy Resins for Ultrathin Sectioning in Electron Microscopy

Fixed tissue is dehydrated with tertiary butyl alcohol overnight. The following day it is cleared in toluene, infiltrated and embedded in Araldite resin-hardener-accelerator mixture without dibutyl phthalate, and polymerized at 60° C. More rapid than previous techniques, this method gives blocks which do not fracture unduly on trimming and provides sections of soft tissues at 1 μ for phase contrast microscopy, as well as ultrathin sections which cut as easily with glass knives as sections of methacrylate. Araldite manufactured in the U.S.A. and in England are different. Satisfactory proportions for the American are: hardener DDSA, 3.5 ml; casting resin 6005, 5.0 ml; accelerator B, 0.12 ml. For the British product, these are: hardener 964 B, 5.0 ml; casting resin M, 5.0 ml; accelerator 964 C, 0.25 ml. The use of 2% agar for orienting small specimens in Araldite is feasible. Mallory's borax-methylene blue has been applied to the staining of Araldite sections as thin as 0.5 μ mounted on glass slides.     

Thin Histological Sections Prepared from Large Thick Sections: a New Technique

A method is described for obtaining thin (1 μm) sections for light microscopy from large area thick (100 μm) sections of low viscosity nitrocellulose embedded specimens of human spinal osteoligamentous material.     

Hematoxylin Toluidine Blue-Phloxinate Staining of Glycol Methacrylate Sections of Retina and Other Tissues

For a detailed study of the developing chick retina a technique has been developed using glycol methacrylate embedding and a hematoxylin toluidine blue-phloxinate stain. After removal of the vitreous body, one half-segment of the eye is dehydrated through graded ethyl alcohols to 95%, infiltrated and embedded in glycol methacrylate, and sectioned at 2 μm. The sections are stained in alum hematoxylin and then in a mixture containing toluidine blue-phloxinate from a stock solution of the dye that has matured for 2-3 weeks. Differentiation is not required and there is only slight staining of the plastic matrix. The quality and clarity of the sections contrasts markedly with that of similarly stained 5 μm paraffin wax sections of the retina. This technique has also been applied to skin, spinal cord, dorsal root ganglia, pancreas and small intestine. The stained sections from these tissues have proved very useful in revealing structural components.     

Histochemical Demonstration of Enzyme Activities in Plastic and Paraffin Embedded Tissue Sections

Histochemical staining for enzymes is usually performed on frozen sections. This report lists the longer incubation times required to demonstrate esterase, acid phosphatase, β-galactosidase, and cytochrome oxidase in plastic embedded and routine paraffin embedded tissues. The sections embedded in plastic, i.e. water soluble methacrylate (Polyscience's JB-4) and cut at 2 μm, were far superior to frozen Sections and paraffin embedded sections both in tissue detail and in the localization of the histochemical reaction product.     

Double Emulsion Autoradiography for Identifying Tritium-Labelled cells in sections

The sensitivity of tissue autoradiography can be doubled and the number of false negative cells nearly eliminated by interposing thin tissue sections between two layers of photographic emulsion. A mouse was given 50 μc of tritiated thymidine (SA 2,500 c/M) intraperitoneally and killed 1.5 hr later. A portion of the small bowel was removed, fixed and embedded in methacrylate in the usual way. Sections 2 μ thick were cut and allowed to flatten on water at 40° C. Some sections were used to make conventional single emulsion auto-radiographs and other sections were interposed between two layers of emulsion by first coating slides with NTB 3 emulsion, picking up the sections from a water bath at 18° C, drying, soaking 1 min in benzene, drying, and then dipping again in NTB 3 emulsion. They were exposed at 4° C in a low humidity, 100% CO₂ atmosphere for 10 days, developed and covered in the usual way. There was an average of 20.16 ± 1.4 grains per labelled cell in the double emulsion group compared with 10.6 ± 0.9 grains in the single emulsion group. In the double emulsion autoradiographs there were 55.1 ± 1.65 labelled cells per unit area as compared with 39.8 2 2.0 in the single emulsion group.     

Protective Paraffin Infiltration of Soft Tissues in Insects to Facilitate Softening of Hard Exoskeletons

This technique can produce serial sections as thin as 5 μ from hard chitin-covered materials of insects or other arthropods. Procedures: Fix with alcoholic Bouin's fluid for 3 hr. Henceforth subject material to partial vacuum in each step to ensure a final proper embedding. Wash with 80% ethanol 2 or 3 times for 2 hr or until the picric acid is largely removed. Dehydrate to 90% ethanol and give 2 changes of n-butanol 2 hr each, and one of a 1:1 n-butanol-paraffin mixture in 56-57° oven for 12 hr. Finally, use 2 baths of pure paraffin, 3 hr each, to complete the infiltration. After the last bath, withdraw the specimen from the paraffin, and remove the superficial paraffin, first mechanically and then with a xylene bath for 4 min. Rinse first with n-butanol, and afterwards with absolute ethanol, 2 min each. The compound eyes are protected with a paraffin covering, the specimen is hydrated with a 1% aqueous solution of detergent for 1 hr and then washed with running tap water. The material is treated with a concentrated sulfuric-nitric mixture (H₂SO₄:HNO₃) for 4 hr to eliminate the exoskeleton. After this treatment, the specimen is washed with running tap water for 12 hr, dehydrated with acetone and then bathed in a 2% solution of celloidin in ethyl acetate to form a protective artificial cuticle. This coating is hardened with 2 quick baths of chloroform, the specimen reembedded in paraffin, and the block cast for sectioning.     

Immunofluorescence Microscopy of Cyanurated Tissues

Test tissues consisted of: (1) popliteal lymph nodes of rabbits, removed 6 hr after injection of hind footpads with 0.2 ml of 125 mg/ml solution of 5× crystallized chicken ovalbumin, and (2) lungs from guinea pigs, passively sensitized with rabbit antiovalbumin serum, then anaphylactically shocked by intracardial injection of a 1% chicken ovalbumin solution. Similar control tissues from normal rabbits, and lungs of passively sensitized guinea pigs, but shocked with histamine instead of ovalbumin, were included. Pieces of fresh tissue not exceeding 2 mm³ were fixed as follows: (1) Cyanuration—lymph nodes, 1 hr; lung, 0.5 hr; both at 23-27 C—in anhydrous methanol containing 0.5% w/v cyanuric chloride and 1% v/v N, N-diethylaminoethanol. (2) Control fixatives—all specimens 18-24 hr at 4—6 C—absolute methanol; 95% ethanol; neutral buffered 10% formalin; and an FAA mixture (formalin, conc., 6; glacial acetic acid, 2; 30% ethanol, 92). Freeze-dried material was either left unfixed (a control) or fixed in xylene or toluene containing 0.5% w/'v cyanuric chloride and 1% v/v N, N-diisopropylaminoethanol; time and temperature as for fresh tissues. All tissues were routinely dehydrated, cleared, and vacuum embedded in an ester wax, diethylene glycol distearate, or in paraffin at 52 C. Sections 2-4 μ thick were attached to gelatin-coated slides, the wax removed in petroleum ether, and stained 20 min at 23-27 C in a 0.10% solution of fluorescein isothiocyanate-conjugated rabbit antiovalbumin globulin, washed in phosphate buffered saline 10 min, dehydrated, cleared and covered in a nonfluorescent medium. With ultraviolet illumination, brightly immunofluorescent, anti-genically specific staining was obtained in cyanurated fresh and freeze-dried lymph node and lung tissues. In contrast, specific staining was diminished or absent in comparable tissues reacted in the control fixatives.     

Orcein-Hematoxylin in Iodized Ferric Chloride as a Stain for Elastic Fibers, With Metanil Yellow Counterstaining

Autopsy and biopsy specimens of human skin were fixed overnight in alcoholic Bouin's solution, embedded in paraffin, cut at 7 μ, deparaffinized, hydrated to 70% alcohol, and treated as follows—stained 2 hours in a mixture consisting of: 0.2% orcein in 70% alcohol and 1% HC1 (conc.), 125 ml; 5% hematoxylin in absolute alcohol, 40 ml; 6% FeCl₃ in water, 25 ml; and aqueous I₂-KI (1:2:100), 25 ml—rinsed in distilled water until the excess stain was removed—differentiated in 1.2% FeCl₃, 5-15 sec—washed in running water, 5 min—differentiation completed in 0.01% HC1 acid-alcohol, 1 min—a dip in 95% alcohol—distilled water, 2 min—0.25% aqueous metanil yellow, 5-10 sec—a 95% alcohol dip—dehydrated in absolute alcohol, xylene, and mounted in a resinous medium. The technic combines the orcein of Pinkus' stain and the hematoxylin mixture of Verhoeff into a single staining solution and gives sharp and reliable results for both coarse and extremely delicate elastic fibers. These stain purple; nuclei, violet; and background, yellow. The stain allows the use of formalin, Bouin's fluid and Zenker-formol fixation. The results have been consistent in other primates as well as in man.     

Selective Staining of Neurosecretory Material in Semithin Epoxy Sections by Gomori's Aldehyde Fuchsin

The epoxy resin was removed from semithin (1 μm) sections by immersing them for 30 sec in sodium methoxide (Mayor et al., J. Biophys. Biochem. Cytol., 9: 909-10, 1961) and then processed as follows: (1) left for 1-3 hr at 60 C in a mixture of formalin, 25 ml; glacial acetic acid, 5 ml; CrO₃, 3 gm; and distilled water, 75 ml: (2) oxidized 10 min in a 1:1:6 v/v mixture of 2.5% KMnO₄, 5% H₂SO₄ and distilled water: (3) bleached in 1% oxalic acid, and (4) stained for 15 min in aldehyde fuchsin, 0.125% in 70% alcohol, or in a 1% aqueous solution of toluidine blue. The neurosecretory material is selectively stained.     

Preparation of Large Epoxy Sections for Light Microscopy as an Adjunct to Fine-Structure Studies

Tissue blocks 2 × 2 × 0.4 cm were fixed 6-24 hr in phosphate-buffered 5% glutaraldehyde then sliced to 2 × 2 × 0.1 cm and soaked in 0.1 phosphate-buffer (pH 7.3) for at least 12 hr. Fixation was continued for 2 hr in phosphate-buffered 1-2% OsO₄. The slices were dehydrated, infiltrated with Araldite, and embedded in flat-bottomed plastic molds. Sectioning at 1-8 μ with a sliding microtome was facilitated by addition of 10% dibutylphthalate to the standard epoxy mixture. The sections were spread on warm 1% gelatin and attached to glass slides by drying, baking at 60 C, fixing in 10% formalin or 5% glutaraldehyde and baking again. Sections were mordanted in 5% KMnO₄ (5 min), bleached with 5% oxalic acid (5 min) and neutralized in 1% Li₂CO₃ (1 min). Several stains could then be applied: azure B, toluidine blue, azure B-malachite green, Stirling's gentian violet, MacCallum's stain (modified), tribasic stain (modified) and phosphotungstic acid-hematoxylin. Nuclei, mitochondria, specific granules, elastic tissue or collagen were selectively emphasized by appropriate choice of staining procedures, and cytologic detail in 1-3 μ sections was superior to that shown by conventional methods. Selected areas from adjacent 4-8 μ sections could be re-embedded for ultramicrotomy and electron microscopy.     

Acceleration of Mallory's Phosphotungstic Acid-Hematoxylin Staining for Skeletal Muscle Fixed in Formalin

The staining time for mammalian skeletal muscle fixed in neutral phosphate-buffered formalin was shortened from 12-24 hr to 10-30 min. The permanganate-oxalate sequence was omitted although oxidation by periodic acid or with iodine was found to be necessary. The material was embedded in paraffin and cut 6 μ or less. Deparaffinized sections were treated with 1% alcoholic iodine for 10 rain followed by 5% Na₂S₂O₃ for 2 min and placed in an oven at 60 C for 10-30 min to stain in a preheated mixture of 50 ml of ripened Mallory's phosphotungstic acid-hematoxylin and 1 ml of 2% phosphomolybdic acid. Experiments with fixation showed that the staining procedure followed Zenker's fluid successfully but not Bouin's fluid. Oxidation by KMnO₄ was effective only after Zenker fixation; oxidation by CrO₃ was unsuccessful.     

Method for Histological Preparation of Bone Sections Containing Titanium Implants

A thin sectioning technique involving hand grinding has been developed to produce 20—40-μn-thick sections of bone-titanium implant sites. Components include: 1) surface staining of sections prior to mounting on slides so bone labels (oxytetracycline-HCI and 2,4-bis(N,N-dicarbometnyl) aminomethylfluorescein (DCAF)) can be seen in sections viewed with transmitted light, 2) a pneumatic sample press for bonding sections to slides with a thin, uniform glue line and without trapped air bubbles, and 3) bonding methyl methacrylate embedded sections to clear acrylic slides with methyl methacrylate monomer to provide enhanced bond strength and grinding properties compared to those obtainable with glass slides. Sample cracking and distortion is minimized and the tissue-implant interface can be kept intact The expense of start-up equipment for this technique is minimal.