Formic Acid Corrosion for the Demonstration of Pulmonary Elastic Tissue

By a revised technique, human pulmonary elastic tissue can be isolated in a form suitable for examination under the stereoscopic microscope. Fresh human lungs from autopsy are fixed by intrabronchial infusion with 10% formalin for 24 hr. Slabs 1.5 cm thick are cut and the formalin removed in running water. One such slab is embedded under intermittent vacuum in an aqueous mixture containing 15% gelatin, 10% glycerol, and 1% phenol; then allowed to gel. Frozen sections 2 mm thick are cut on a large-section MSE sledge microtome. Squares 3 × 3 cm from such a section are corroded for 4-5 days in 88% formic acid at 45 C, washed once with distilled water, and mounted in glychrogel containing 6% gelatin. The elastic tissue network of the lung will have been freed from surrounding elements. The preparation should be stored in a refrigerator. Blocks for thin sections and large thick un-corroded sections can be prepared from the same lung as part of an over-all procedure.     

A Sudan Black B Myelin Stain for Peripheral Nerves

Blocks of fresh issue were fixed 2 or more days in: cobalt sulfate (or nitrate), 1 gm; distilled water, 80 ml; 10% calcium chloride, 10 ml; and formalin, 10 ml. The fixed tissue was washed thoroughly in tap water, embedded in gelatin, frozen sections cut, and mounted on slides with gelatin adhesive. The sections were stained 15-30 min in a saturated, filtered solution of Sudan black B in 70% alcohol, differentiated in 50% alcohol under microscopic observation, and a cover glass applied with glycerol-gelatin. In thick (50-100 μ) sections, myelin stained green to gray-green and this allowed easy differentiation between nerves and other tissue elements.     

A Modified Verhoeff-Van Gieson Elastin Histochemical Stain to Enable Pulmonary Arterial Hypertension Model Characterization

Optimal histochemical staining is critical to ensure excellent quality stained sections to enable light microscopic and histomorphometric image analysis. Verhoeff-van Gieson is the most widely used histochemical stain for the visualization of vascular elastic fibers. However, it is notoriously difficult to differentiate fine elastic fibers of small vasculature to enable histomorphometric image analysis, especially in organs such as the lung. A tissue fixation procedure of 10% neutral buffered formalin with subsequent fixation in 70% ethanol further compounds the problem of small vessel staining and identification. Therefore, a modified Verhoeff’s elastin stain was developed as a reliable method to optimally highlight the internal and external elastic laminae of small arteries (50-100 µm external diameter) and intra-acinar vessels (10-50 µm external diameter) in 3 µm thick lung tissue sections from models of pulmonary arterial hypertension. This modified Verhoeff’s elastin stain demonstrated well-defined staining of fine elastic fibers of pulmonary blood vessels enabling subsequent histomorphometric image analysis of vessel wall thickness in small arteries and intra-acinar vessels. In conclusion, modification of the standard Verhoeff-van Gieson histochemical stain is needed to visualize small caliber vessels’ elastic fibers especially in tissues fixed in 10% neutral buffered formalin followed by additional fixation in 70% ethanol.Key words: Histochemical stain, histomorphology, lung, Verhoeff-van Gieson, elastin     

Controlled Differentiation of Cells and Connective Tissue Fibers in Silver Staining

Controlled silver staining of connective tissue fibers and sometimes of these fibers and cells simultaneously can be obtained. 1. Fix in 10% formalin. Embed in paraffin and cut sections as usual, but do not mount them on slides. Deparaffinize and hydrate through xylene, alcohols and distilled water and henceforth treat them the same as frozen sections. Real frozen sections can also be used. 2. Treat with a freshly prepared 1% solution of KMnO₄, usually 15-60 sec, sometimes up to 10 min. 3. Wash in distilled water, 5-10 sec. 4. Decolorize in 2% potassium metabisulfite, 10-20 sec. 5. Place in distilled water, 1 min. 6. Sensitize with 2% iron alum, 1 min. 7. Place in distilled water, 1 min. 8. Impregnate in Gomori's silver oxide solution, 2 min. 9. Wash in a 1.5% aqueous solution of pyridine, about 15 sec. 10. Reduce in a mixture containing 0.25% gelatin and 2% formalin 1 min. 11. Repeat steps 7 to 10 once or several times until the connective tissue fibers are completely stained. For cell staining (which may fail) proceed as follows: After the first insufficient staining of the connective tissue fibers, rinse in distilled water, dip for 1 sec in Gomori's solution and reduce immediately in gelatin-formalin without previous washing in pyridined water. This step can be repeated. 12. If the staining is too strong, decolorize as needed in 2% iron alum. 13. Toning in 0.2% gold chloride, 5 min or more, followed by fixation in 5% sodium thiosulfate, 1 min, is optional. Counterstain as desired. 14. Wash in tap water, dehydrate, clear in xylene and mount in balsam. The same technique applied to sections attached to slides gives good results but inferior to that obtained in paraffin sections processed in the loose, unmounted condition.     

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.     

A Method for Detecting Visceral Malformations in Gelatin-Embedded Rat Fetuses Using an Automatic Slicing Apparatus

A method for observing whole rat fetal viscera embedded in gelatin using an automatic slicing apparatus is described. Fetuses were immersed in Bouin's solution. Part of the thoracic and abdominal skin of each fetus was removed, and fetuses were immersed consecutively in sodium bicarbonate 30% in 70% ethanol, gelatin 15% in water, gelatin 30% in water, then embedded in fresh 30% gelatin. The gelatin blocks containing the fetuses were immersed in 10% formalin. After fixation, the block was sliced into 200 μm serial transverse sections using a rotor-slicer at a rotation speed of 120 rpm and a cutting speed of 25 mm/sec. Complete slicing of a single fetus required about 20 min. The advantages of the method presented here include: complete fetal serial sections are produced, thin and uniform sections are obtained easily, viscera can be identified easily, and observation can be carried out at any time after slicing. The method presented can be used to detect whole fetal visceral malformations in developmental toxicity tests.     

Suppression of Connective Tissue Impregnation in a Silver Technique for Demonstrating Nerve Fibers

A tissue pretreatment technique is introduced which effectively suppresses the silver impregnation of connective tissue and nompecific background elements in peripheral nerve. The result is a selective impregnation of nerve fibers. The procedure utilizes fresh frozen sections and can be used with the Holmes (1947) or Bodian (1936) techniques. Fresh frozen sections are cut at 10 microns, mounted on slides and air dried for 5 minutes. They are fixed for 30 minutes in formol-sublimate (10% formalin saturated with mercuric chloride) and then placed into 0.5% iodine in 70% alcobol for 5 minutes followed by bleaching in 2.5% sodium thiosulfate for 2 minutes. After washing in running tap water for 10 minutes and a brief rinse in distilled water, impregnation is accomplished by the Holmes (1947) or Bodian (1936) procedure beginnins with the step containing the aqueous silver solution. The results show an absence of impregnation of connective tissue and nonspecific background. The technique is simple, rapid, and, by utilidng fresh hrozen sections, can be used for other histological and histochemical purposes. Several experiments were done to determine the causes of the connective tissue and background suppression. The air drying step was omitted; the sections were fixed in formalin without mercuric chloride; and the formol-sublimate fixation time was increased. The results suggest that connective tissue impregnation H suppressed by the use of mercuric chloride in the fixative and that the background supprgsion is related to the short fixation time with formol-sublimate.     

Suppression of connective tissue impregnation in a silver technique for demonstrating nerve fibers.

A tissue pretreatment is introduced which effectively suppresses the silver impregnation of connective tissue and nonspecific background elements in peripheral nerve. The result is a selective impregnation of nerve fibers. The procedure utilizes fresh frozen sections and can be used with the Holmes (1947) or Bodian (1936) techniques. Fresh frozen sections are cut at 10 microns, mounted on slides and air dried for 5 minutes. They are fixed for 30 minutes in formol-sublimate (10% formalin saturated with mercuric chloride) and then placed into 0.5% iodine in 70% alcohol for 5 minutes followed by bleaching in 2.5% sodium thiosulfate for 2 minutes. After washing in running tap water for 10 minutes and a brief rinse in distilled water, impregnation is accomplished by the Holmes (1947) or Bodian (1936) procedure beginning with the step containing the aqueous silver solution. The results show an absence of impregnation of connective tissue and nonspecific background. The technique is simple, rapid, and, by utilizing fresh frozen sections, can be used for other histological and histochemical purposes. Several experiments were done to determine the causes of the connective tissue and background suppression. The air drying step was omitted; the sections were fixed in formalin without mercuric chloride; and the formol-sublimate fixation time was increased. The results suggest that connective tissue impregnation is suppressed by the use of mercuric chloride in the fixative and that the background suppression is related to the short fixation time with formolsublimate.     

Silver Impregnation of the Golgi Apparatus, with Subsequent Nitrocellulose Embedding

The appearance of silver impregnation of the Golgi apparatus can be enhanced by the use of nitrocellulose as an embedding medium. Fixation of 1.5 mm thick pieces of fresh tissue for 8 hr in: glycine, 1.7 gm; 15% formalin, 100 ml; HNO₃, conc., 0.5 ml, at pH 2.6 followed by rinsing in water, 4 hr in 1.5% AgNO₃, another rinse, and 2 hr reduction in 1.5% hydroquinone in 15% formalin. This staining procedure yields consistently good results for rat, rabbit, and human tissues. Low-viscosity nitrocellulose embedding is done by infiltrating at 56 C in 7% nitrocellulose for 0.5 hr, 15% for 4 hr, and 27% for 1 hr. The nitrocellulose is hardened 2 hr in chloroform, after which, sections as thin as 5 μ can be cut on a sliding microtome. Gold toning and counterstaining can be done with the tissue affixed to the slide. The Golgi apparatus is stained dark brown to black, and there is better preservation of cellular detail than in tissues processed in paraffin.     

Zinc Chromate Modification of the Golgi Technic

A modification of the Golgi technic is described in which the reaction takes place in well fixed formalin material. Thin slices (whole sections of adult monkey, cat and rat cerebrum) 2 to 3 mm. thick, from brains fixed 3 to 4 months in 10% formalin, are chromated for two days in 3 g. of zinc chromate dissolved in 98 ml. of distilled water and 2 ml. of formic acid. Slices are then removed, blotted dry and immersed, suspended by a thread, in 0.75% silver nitrate solution for two days. Solution should be changed after the first day. After silvering, the slices are dehydrated rapidly (total time about one hour) in 95% and absolute alcohol, placed in xylene 10 minutes, in low melting point paraffin 10 minutes and embedded in low melting point paraffin. Only surface infiltration is necessary since sections are cut 90 to 100 u. Sections are collected in 95% alcohol, dehydrated in absolute alcohol, cleared in several changes of xylene and mounted in Fisher's Permount. Results with fetal and new born material were not good.     

A One-Solution Triacid General Stain Which Differentiates Oxygenated from Non-Oxygenated Red Blood Corpuscles

Tissues from representative mammals, amphibia and invertebrates were fixed for 5-24 hr in either an aqueous solution of 8% p-toluene sulfonic acid (PTSA) or in 10% formalin to which 5 gm PTSA/100 ml had been added, and processed through embedding in polyethylene glycol 400 distearate in the usual manner. Sections cut at 4-6 μ were floated on 0.2% gelatin containing 1.25% formalin, and spread and dried on slides at a temperature not exceeding 25 C. Wax was removed with xylene, and the sections brought to water through ethanol as usual. The working staining solution was made from three stock solutions: A. Chlorantine fast blue 2RLL, 0.5%; B. Cibacron turquoise blue G-E, 0.5%; C. Procion red M-P, 0.5%—each of which was dissolved in 98.5 ml of distilled water to which 0.5 ml of glacial acetic acid and 0.5 ml of propylene glycol monophenyl ether (a fungicide) had been added. For use, the three solutions were mixed in the proportions: A, 3; B, 4; and C, 3 volumes. Staining time was uncritical, 10-30 min usually sufficing for 6 μ, sections. The chief feature of the staining is the differentiation of oxygenated and nonoxygenated red blood corpuscles, in reds and blues respectively. Connective tissue stained blue or blue-green and mucin, green. Nuclei and cytoplasm stain according to their condition at the time of fixation. The mixed stain keeps well, remaining active after 2 yr of storage.     

Triple Silver Impregnation for Selective Staining of Avian Nerves

Frozen sections of avian tissue fixed 7 days or longer in 10% formalin or formol-saline are cut at 20-50 μ, left in distilled water for 2 hr, and placed in 0.002% aqueous AgNO₃ for 3-4 days. Subsequent procedure is essentially that of Weddell and Glees. Sections are placed in 20% AgNO₃ for 30 min, then carried through 3 baths of 3% formalin in less than 10 min. Immediately thereafter they are washed 1-2 sec in a 0.1% solution of NH₄OH (cone) and placed in the ammoniacal silver solution (made with 20% AgNO₃) until the nerves become distinct, as seen under a microscope; usually, in about 15 min. After washing briefly, the sections are fixed in 5% Na₂S₂O₃ for 3-10 min, dehydrated, cleared, and mounted in the usual way.     

Improved Lipid Visualization with a Modified Osmium Tetroxide Method Using Ultrasonic Treatment and Intensification with Imidazole or Triazole

Formalin fixed autopsy tissue containing lipids were cut into 1-5 nun thick blocks, washed well, then postfixed in 2% OsO₄ in 0.03 M veronal acetate buffer for 30, 60, 90, 120, or 180 min with or without ultrasonic treatment. Tissues exposed to ultrasound for 90 min showed superior penetration of OsO₄ and well preserved histological architecture. Tissues also were immersed for 1 hr in veronal acetate buffer (pH 7.4) containing 0.5% imidazole or triazole and compared with untreated controls. Paraffin sections, 4 μm thick, were examined under a light microscope with an image analyzer. Both intensity and percentage area of osmium blackening were significantly higher in samples immersed in imidazole or triazole than in untreated controls. No difference was observed between imidazole- and triazole-immersed samples. The OsO₄ method, modified by ultrasound treatment and imidazole- or triazole-immersion, can be applied to routine formalin fixed autopsy materials for improved lipid visualization.     

Demonstration of surfactant phospholipids in frozen sections of the lung

Saturated phospholipids are known to be the only surface active compounds present in the surfactant system of the lung. Using light microscopy, the identification in situ of pulmonary surfactant has always been hampered by the lack of satisfactory fixatives and dyes which act on saturated phospholipids fast enough to prevent the complete loss of surfactant in the solutions. In this study we adopted the tricomplex flocculation proposed by Elbers et al. (1965) to fix surfactant phospholipids on frozen sections obtained from human, pig and rat lungs. Small pieces of lung tissue were quickly frozen in freon 22 kept at -75 C.; eight micron sections were cut in a cryostat, air dried and immersed for 30s-5m in a 0.05 N Pb(NO3)2 + K3Fe (CN)6 solution in 10% formalin. Lead ions bound to the choline portion of phospholipid molecules were subsequently revealed in a 30 mM ammonium sulfide solution. This procedure delineates a dark brown filmy structure in the respiratory parenchyma, which is very loosely attached to the alveoli and appears to be related to lung surfactant. Preliminary lung lavage or pretreatment of sections with saline, aldehyde fixatives and several organic solvents, fully or partially abolish the stain.     

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.     

Fink-Heimer Silver Impregnation of Degenerating Axons and Terminals in Mounted Cryostat Sections of Fresh and Fixed Brains

The silver impregnation method of Fink and Heimer (Brain Res., 4: 369-74, 1967) has been used on cryostat sections of both fresh frozen and formalin-fixed brain tissue mounted on slide. The fixed brains were soaked in 25% sucrose for 2-3 days before freezing. The slides used for mounting were dipped in a 0.5% aqueous gelatin solution containing 50 mg of chrome-alum per 100 ml, drained and allowed to dry in a vertical position. Sections of fresh tissue were fixed for 16 hr in a 10% formalin solution buffered with phosphates to pH 7.0. Staining was carried out according to the Fink-Heimer procedure II and gave results comparable to those obtained on unmounted frozen sections of formalin-fixed material     

Fink-Heimer Silver Impregnation of Degenerating Axons and Terminals in Mounted Cryostat Sections of Fresh and Fixed Brains

The silver impregnation method of Fink and Heimer (Brain Res., 4: 369-74, 1967) has been used on cryostat sections of both fresh frozen and formalin-fixed brain tissue mounted on slide. The fixed brains were soaked in 25% sucrose for 2-3 days before freezing. The slides used for mounting were dipped in a 0.5% aqueous gelatin solution containing 50 mg of chrome-alum per 100 ml, drained and allowed to dry in a vertical position. Sections of fresh tissue were fixed for 16 hr in a 10% formalin solution buffered with phosphates to pH 7.0. Staining was carried out according to the Fink-Heimer procedure II and gave results comparable to those obtained on unmounted frozen sections of formalin-fixed material     

A gelatin film procedure for the localization of proteolytic activity in Leucochloridiomorpha constantiae (Trematoda) adults

A gelatin film procedure was used to localize proteolytic activity in the lumen of the intestinal ceca of Leucochloridiomorpha constantiae (Trematoda) adults. Slides were coated with a 7·5% gelatin solution and then fixed in 10% neutral buffered formalin (NBF). Cryostat sections of isolated worms or those attached to the bursa of Fabricius of the domestic chick were affixed to the gelatin film. Experimental and control slides were incubated in a humid chamber for 30 min at 37·5 and 4°C, respectively. Slides were again fixed in NBF, and then stained for protein with mercuric bromphenol blue (MBB). In experimental slides, the lumen of the intestinal ceca was lysed and did not stain, whereas worm and host tissue and the gelatin were protein-positive. Control sections stained uniformly positive for protein. In this procedure tissue is retained on the slide and proteolytic activity can be correlated with a tissue site on the same slide.     

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.     

Freeze-Sectioning of Plant Tissues

Techniques are described for freeze-sectioning a wide range of both fresh and fixed plant tissues. Gelatin-antifreeze media are used to support but not infiltrate the tissue during sectioning. At cryostat temperatures of -10 to -15 C, 15% gelatin (w/v) containing 0.8% dimethyl sulfoxide (DMSO), or 1.5% ethanediol (ethylene glycol), or 2% glycerol is used. Lower concentrations of gelatin and higher concentrations of antifreezes are required for sectioning at -24 C. Petri plates of media are stored at 2 C, and used by simply melting a hole in the medium. Fresh tissues can be placed directly in the hole, or prefrozen at temperature of liquid nitrogen, or equilibrated in antifreeze solution, before freeze-sectioning in the gelatin antifreeze medium. Many plant tissues have highly vacuolated cells and need equilibration in antifreeze solutions prior to freeze-sectioning. Fixed tissues are rehydrated and washed in water or buffer for 15-24 hr before equilibrating in a 10% solution of either DMSO, ethanediol or glycerol (named in order of rapidity of equilibration). Pretreatment in 10% DMSO is usually for 1-6 hr at 2 C for histochemical studies; or in 10% ethanediol or glycerol for 15-24 hr at either room temperature or 37 C for morphological studies. These methods permit serial cryostat sections free from freezing and thawing artifacts to be cut as thin as 2 μ.