Plastic sectioning for light microscopy of Pyrenomycetes.

In preparation for light microscopy, ascocarps of Sordaria fimicola Ces. & DeNot. were embedded in Spurr's medium and sectioned at 1-1.5 mum on an ultramicrotome. Sections were floated on Giemsa staining solution at 60 C for 10-30 min, washed in distilled water, affixed to slides by drying, and mounted in immersion oil. Best preservation of the delicate sterile tissues of the centrum was obtained by fixation in 1% KMnO4 for 2.5-3 hr, followed by the Giemsa stain. This method is suggested for future studies on the morphology of perithecial ascomycetes.     

Sudan Black and Osmic Acid as Staining Agents for Testicular Interstitial Cells

Two standard cytological techniques have heen modified to stain specifically the interstitial cells of the testis. In Method 1, the tissue is fixed in Zenker-formol or Regaud's fluid for several hours or overnight and subsequently postchromed in 3% K₂Cr₂O₇ for 72 hr at 37°C. After paraffin embedding, sections are cut at 5μ, dewaxed, brought down to 70% alcohol and stained in an unfiltered saturated solution of Sudan black in 70% alcohol for 10-30 min. Sections are washed briefly in 70% alcohol to remove all excess dye, differentiated, if necessary, in 50% alcohol, downgraded to water and mounted in Farrants' medium or glycerol jelly. Interstitial cells: deep blue black; remainder of testicular tissue: light blue. Method 2 is essentially the Champy-Kull technique but specific staining for mitochondria is omitted and the sections are downgraded to water; then they are mounted in Farrants' medium or glycerol jelly without further treatment. In this way osmicated lipoids are preserved. Interstitial cells: conspicuous due to the variable number of black granules in their cytoplasm; the remainder of the tissue: yellow.     

Plastic Sectioning for Light Microscopy of Pyrenomycetes

In preparation for light microscopy, ascocarps of Sordaria fimicola Ces. & DeNot. were embedded in Spurr's medium and sectioned at 1-1.5 μm on an ultramicrotome. Sections were floated on Giemsa staining solution at 60 C for 10-30 min, washed in distilled water, affixed to slides by drying, and mounted in immersion oil. Best preservation of the delicate sterile tissues of the centrum was obtained by fixation in 1% KMnO₄ for 2.5-3 hr, followed by the Giemsa stain. This method is suggested for future studies on the morphology of perithecial ascomycetes.     

Localization of Arbutin in Pyrvs by Means of Alkaline P-Phenylenediamine

A reagent composed of 0.2% p-phenylenediamine in 2 N NH₄OH was used for the cytochemical demonstration of arbutin in plant tissue. Sections of fresh tissue were cut at 25-50 μ, mounted in a drop of the reagent, and allowed to stand uncovered 15-20 min before applying a coverslip. Arbutin stained dark blue to dark purple and was easily distinguished from other constituents of the cell, such as chlorogenic acid, isochlorogenic acid, caffeic acid, and quinic acid, which stained yellow, yellow-green, red or brown in color. The limit of sensitivity of the p-phenylenediamine-arbutin reaction was 1:100,000, as determined by spot-plate tests.     

Fluorescence in Paraffin Sections of Dye-Labelled Protein Administered in vivo: Effect of HgCl2 Fixation

Lung and liver slices, 2-3 mm thick, from guinea pigs injected intravenously with fluorescent dye-protein conjugate are fixed for 15-30 min in saturated aqueous HgCl₂, dehydrated in ethanol, cleared in xylene and embedded in paraffin at 60 C. Mercurial deposits are removed with I₂KI from 5 μ sections taken to water, and the iodine then removed with 5% Na₂S₂O₃. Sections are mounted from xylene into permanent nonfluorescent mounting medium. This procedure gives optimal fluorescence which is not decreased by the technic of removing mercurial precipitates. Longer fixation, fixation in phosphate-buffered formalin, or in an HgCl₂-formalin mixture gives inferior results.     

A new method for identification of cement lines in undecalcified, plastic embedded sections of bone

A gallocyanin method for demonstrating cement lines in thin, undecalcified sections of bone has been developed that is compatible with prestaining with osteochrome before plastic embedding. After sectioning at 5 microns on the Jung K heavy duty microtome, the sections are attached to a microslide using Haupt's adhesive mounting medium, placed on a slide warmer at 37 C until completely dry, and deplasticized in xylene at 45 C for 16-24 hr. Sections are stained with 0.15% gallocyanin-5% chrome alum solution for 30 min, followed by staining in buffered Villanueva blood stain for 1-1 1/2 hr, quickly dehydrated, differentiated in equal parts xylene and 100% ethanol, cleared, and mounted in Eukitt's medium. Reversal lines appear as thin, scalloped, blue or purple lines approximately 0.3 micron wide, and arrest lines as thick, homogeneous, straight or evenly curved, dark blue or purple lines approximately 2 microns wide. The method also demonstrates abnormal halo volumes around osteocytes, old and new bone matrix, osteoid seams, and the granular mineralization front at the osteoid-bone interface. It promises to be valuable in the study of age-related bone loss, osteoporosis, and metabolic bone disease.     

A New Method for Identification of Cement Lines in Undecalcified, Plastic Embedded Sections of Bone

A gallocyanin method for demonstrating cement lines in thin, undecalcified sections of bone has been developed that is compatible with prestaining with ostcochrome before plastic embedding. After sectioning at 5 pm on the Jung K heavy duty microtome, the sections are attached to a microslide using Haupt's adhesive mounting medium, placed on a slide warmer at 37 C until completely dry, and deplasticized in xylene at 45 C for 16-44 hr. Sections are stained with 0.15% gallocyanin-5% chrome alum solution for 30 min, followed by staining in buffered Villanueva blood stain for 1-1 1/2 hr, quickly dehydrated, differentiated in equal parts xylene and 100% ethanol, cleared, and mounted in Eukitt's medium. Reversal lines appear as thin, scalloped, blue or purple lines approximately 0.3 pm wide, and arrest lines as thick, homogeneous, straight or evenly curved, dark blue or purple lines approximately 2 pm wide. The method also demonstrates abnormal halo volumes around ostcocytes, old and new bone matrix, osteoid seams, and the granular mineralization front at the osteoid-bone interface. It promises to be valuable in the study of age-related bone loss, osteoporosis, and metabolic bone disease.     

Frozen Section Micro-Incineration

A method for micro-incineration of frozen sections is described. Material containing diffusible or soluble salts is cut on the freezing microtome and the sections are placed into xylol and mounted out of xylol onto Corex D slides previously filmed with glycerin-gelatin medium. Material containing non-diffusible or insoluble salts can be fixed in 10% formalin before sectioning. Sections of the fixed material are dehydrated thru 50, 70, and 95% ethyl alcohol and mounted out of absolute alcohol onto Corex D slides previously fumed with glycerin-gelatin medium. After mounting by either procedure the sections are incinerated in an electric furnace and the temperature of incineration is dependent on the type of tissues to be incinerated and the character of the salts present. The method is time saving and when no fixation is required the whole procedure can be carried out in one hour.     

The Peracetic-Orcein-Halmi Stain: A Stain for Connective Tissues

A selective stain useful for the study of connective tissues is described. The stain demonstrates elastic and oxytalan fibers as well as fibrils in mucous connective tissues previously undescribed. Reticular fibers are not stained. The stain may be used on sections that have been fresh frozen or fixed in formalin or ethanol. Sections are deparaffinized, washed in absolute ethanol, oxidized in peracetic acid 30 min, washed in running water, stained in Taenzer-Unna orcein 15 min, 37°C, differentiated in 70% ethanol, washed in running water, stained in Lillie-Mayer alum hematoxylin 4 min, blued in running water, and counterstained 20 sec in a modified Halmi mixture of 100 ml distilled water, 0.2 gm light green SF, 1.0 gm orange G, 0.5 gm phosphotungstic acid and 1.0 ml glacial acetic acid. Sections are rinsed briefly in 0.2% acetic acid in 95% ethanol, dehydrated and mounted.     

Gallocyanin as A Nuclear Stain

Gallocyanin has been used successfully as a nuclear stain. Sections are cut by the freezing method of either fixed or unfixed tissue. The tissues are warmed (not exceeding 70°C.) for 2-4 minutes in the gallocyanin solution. A counterstain may be used if desired. The most effective are Biebrich scarlet, phloxine, or eosin Y. The sections are then dehydrated and mounted in clarite. The nuclear pattern is clearly demonstrated and the sections are permanent.     

Identification of normal or neoplastic murine mesenchymal cells in chimeric tissues or heterotransplants

An improved method for identifying murine mesenchymal cells in chimeric tissues or heterotransplants using Hoechst dye 33258 is described. Following fixation in formalin-saline, tissues are embedded in JB-4 plastic. Sections 3 micron thick are then stained in a 10 microgram/ml solution of Hoechst 33258 in Hanks' balanced salt solution for 5-10 min at 4 C. After rising, the sections are coverslipped using a modified polyvinyl alcohol mounting medium. This approach offers several advantages over existing techniques: 1) uniform section thickness is more easily obtained than with paraffin or cryostat microtomy, thereby allowing improved resolution and more reliable identification of mesenchymal cells with small nuclei such as skeletal muscle myocytes or fibroblasts, 2) the preparations are stable over long periods and can be repeatedly viewed or photographed, and 3) calcified tissues can be examined without prior decalcification. An example is shown of species identification using rat chondrosarcoma cells grown in nude mice.     

Identification of Normal or Neoplastic Murine Mesenchymal Cells in Chimeric Tissues or Heterotransplants

An improved method for identifying murine mesenchymal cells in chimeric tissues or heterotransplants using Hoechst dye 33258 is described. Following fixation in formalin-saline, tissues are embedded in JB-4 plastic Sections 3 μm thick are then stained in a 10 μg/ml solution of Hoechst 33258 in Hanks' balanced salt solution for 5-10 min at 4 C. After rinsing, the sections are coverslipped using a modified polyvinyl alcohol mounting medium. This approach offers several advantages over existing techniques: 1) uniform section thickness is more easily obtained than with paraffin or cryostat microtomy, thereby allowing improved resolution and more reliable identification of mesenchymal cells with small nuclei such as skeletal muscle myocytes or fibroblasts, 2) the preparations are stable over long periods and can be repeatedly viewed or photographed, and 3) calcified tissues can be examined without prior decalcification. An example is shown of species identification using rat chondrosarcoma cells grown in nude mice.     

The Demonstration of a new nerve-cell Organoid

A method is described for the demonstration of a new nerve-cell organoid— the binary spheroid systems (Baker bodies). Zenker-formol or acetic-osmium-bichromate materials are postchromed at 37°C. and embedded in paraffin, sectioned and mounted in the ordinary manner. The sections are colored in a 70% alcohol solution of Sudan black B, rinsed in 70% alcohol, counter-stained in Mayer's carmalum and mounted in Farrant's medium. After examination the cover may be removed and the Sudan black extracted in 96% alcohol. The sections can then be restained by the azan method of Heidenhain, and other cytoplasmic inclusions can be correlated accurately to the functional states of the spheroid systems.     

Staining Neural End Feet and Mitochondria After Postchroming and Carbowax Embedding

Spinal cord of cat and rabbit was fixed by perfusion with 10% formalin in physiological salt solution followed by a 2-day immersion in 10% aqueous formalin. Further treatment (postchroming) consisted of a 5-day immersion in: K₂Cr₂O₇, 5 gm; CrFl₃-4H₂O, 2 gm; distilled water, 100 ml; followed by 5% aqueous K₂Cr₂O₇ at 38–40°C for 2–4 wks. After thorough washing, blocks were embedded by infiltration first with polyethylene glycol 1000 M. E. and then with Nonex 63B (Gemec Chemicals Co., London, E. C. 2), and casting in the Nonex. Sections were stained, either mounted or unmounted, by modifications of the Bielschowsky-Gros method, and mounted sections by Weigert-Pal's hematoxylin or by Silver's Protargol method. All 3 methods gave apparently complete staining of pericellular end feet and showed also an abundance of mitochondria. Cytologic preservation was much better than that seen after the usual procedures for this type of staining. Retention of lipoid material in the sections is considered to be the cause of efficient staining of end feet and mitochondria.     

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.     

Studies on the optimalisation and standardisation of the light microscopical succinate dehydrogenase histochemistry

The present investigation was undertaken in order to establish an optimal tissue pretreatment and an optimal incubation medium for the histochemical demonstration of succinate dehydrogenase (E.C. 1.3.99.1). The investigations were performed on steroid producing (testicle, adrenal gland) and steroid dependent (Fallopian tube) tissues. We studied the influences fo formalin fixation, acetone, magnesium ions, cyanides, electron carries (phenazine methosulfate, menadione coenzyme Q10), osmolarity, substrate concentration and inhibitors (oxalacetate, oxalate, malonate, 4-chloromercuribenzoic acid). The following procedure yields blameless morphological integrity and enzyme localization as well as optimal SDH-activity: Freezing of tissue cubes (diameter less than 5 mm) in propane cooled with liquid nitrogen or in melting freon. Incubation of 5 micrometer cryostat sections in narrow jars in the following medium (38.5 ml):--10 ml of 0.2 M sodium phosphate buffer pH 7.6 (52 mM).--18 mg tetranitro-BT in 0.5 ml dimethylformamide and aqua bidest. ad 10 ml (0.5 mM).--2.6 mg KCN in 16 ml aqua bidest. (1 mM).--540 mg succinate (disodium salt, hexahydrate) in 2 ml aqua bidest. (52 mM).--3 mg PMS (phenazine methosulfate) in 0.5 ml aqua bidest. (0.25 mM). The incubation medium has an osmolarity of 440 mosm. The incubation is carried out for 10 min at 37 degree C in darkness. To avoid non specific formazan deposits in lipid containing tissues a preincubation of the cryostat sections in 100% acetone at--22 degree C or--40 degree C for 7--10 min and an incubation time of 20--30 min is recommended. Control incubations adduced proof at the specificity of the SDH demonstration. Parallel incubation without PMS in order to determine indirectly the content of endogenous CoQ10 is further recommended.     

A modified semipermeable membrane technique for carbonic anhydrase histochemistry of the nervous system

We present a modification of Hansson's method for the demonstration of carbonic anhydrase activity. Using a semipermeable membrane together with a fluid incubation medium, frozen sections of aldehyde-fixed tissue were incubated without floating or dipping. Thin sections (thickness, 20-40 microns) were mounted on the outer surface of a tubular-shaped, semipermeable cellophane dialysis membrane containing the incubation fluid. After incubation for 25-30 min at room temperature, the sections were rinsed in buffer and treated with 0.5% (NH4)2S solution. The histochemical reaction was fully inhibited by 10(-4) M acetazolamide.     

An Ultraviolet-Schiff Reaction for Unsaturated Lipids

Tissues are fixed 12-18 hr in cold, neutral, 10% aqueous formalin, sectioned on a freezing microtome and the sections placed in a shallow, open dish of water under an ultraviolet light source. Sections are then treated with Schiff's reagent for 15 min, rinsed in 3 changes of sulfurous acid (3 min in each) and in distilled water and mounted in an aqueous mounting medium. The time of irradiation necessary to produce a maximum reaction is determined empirically. The magenta color indicating the site of unsaturated lipids fades and runs in many preparations in a few days. Evidence is presented that this reaction is most likely the histochemical equivalent of an iodine number determination.     

Mature Cereal Grain Endosperm: Rapid Glass Knife Sectioning for Examination of Proteins

A simple, rapid procedure was developed by which the quality of endosperm protein in cereal grains could be evaluated by microscopic observation of the subcellular protein structure. Kernels with vitreous endosperm were sectioned without pretreatment at 3-4 μ with a glass knife. Floury endosperm tissues were fixed in 10% glutaraldehyde, pH 7.4, for 16 hr at 5 C, and boiled to gelatinize the starch. After drying, this tissue was sectioned as for vitreous endosperm. Sections were mounted on gelatin-coated slides and destarched with a-amylase. To identify sites of prolamine, alcohol-soluble protein was extracted for 1 hr at about 70 C with 80% ethanol. The subcellular proteins were stained with either iodine vapor or various organic dyes. Proteins were differentiated by a combination of staining and mounting in selected high refractive index liquids.     

A Basic Fuchsin and Alkalinized Methylene Blue Rapid Stain for Epoxyembedded Tissue

Sections 1 μ thick of epoxy-embedded, OsO₄-fixed tissues were stained with 4% aqueous basic fuchsin at 70 C for 1 min, rinsed well and destained, also at 70 C, for 1 min. A 2% aqueous methylene blue solution, alkalinized to pH 12.5 by mixing 1 N NaOH with the dye on the slide in the proportion of about 2:1, was then allowed to act for 2 min at 23-27 C. The stain was rinsed off the slide, and the preparation air dried before applying a mounting medium and cover glass. The mounting medium consisted of immersion oil sealed with epoxy household cement. Stains had not faded after 1 yr. The method is simple, rapid (total time 4-5 min), and provides sharp contrast between cellular and connective tissue components.