Embedding with Low Viscosity Nitrocellulose

The gelation and cutting of embedding masses of low viscosity nitrocellulose (L. V. N.) and of celloidin were compared. L. V. N. forms a firmer mass which can be cut into thinner sections than celloidin. It tolerates considerable water (up to 6%) in a solvent system of alcohol-water-ether, thereby permitting the use of 95% alcohol instead of absolute for making up the solution. The fluidity of its solutions permits transfer of tissue directly from alcohol-ether to a 20% embedding solution. Faults of L. V. N. are: the nitrated cotton lint contains some grit (hence its solution should be allowed to settle), the sections cut from it are somewhat more easily torn than celloidin sections, and it is sufficiently soluble in absolute alcohol to preclude the use of this fluid in handling sections.     

A method for obtaining histological sections from tissue previously studied by scanning electron microscopy

An accelerated method of paraffin embedding of tissue specimens previously examined with scanning electron microscopy is proposed aimed to obtain sections for routine histological examination. The tissue is passed through acetone, absolute alcohol, alcoholic-oil celloidin solution, chloroform to be eventually mounted into paraffin. The method allows obtaining good quality sections within 24 hours.     

Alcoholic Thionin Counterstaining Following Golgi Dichromate-Silver Impregnation

Brains of cats that had been fixed 2 months or longer in 10% formalin were cut into 3-6 mm. slices and impregnated by Golgi's dichromate-silver procedure (6% dichromate solution, 4-6 days; 1.5% silver nitrate solution 2 days). Sections 100 µ thick were cut after embedding in low melting point paraffin. Three changes of xylene and three of absolute alcohol were followed by staining 3-5 minutes in a saturated solution of thionin in absolute alcohol. The sections were dipped quickly in absolute alcohol and cleared in xylene, then differentiation was effected by an equal-parts mixture of absolute alcohol and xylene. A final clearing in three changes of xylene and mounting in Permount completed the process. Counter-staining was most successful when applied to freshly cut sections.     

Basic Dye Counterstaining of Sections Impregnated by the Golgi-Cox Method

Celloidin blocks of Golgi-Cox impregnated material are cut at 50 μ, the sections collected in 70% alcohol, transferred to a 3:1 mixture of absolute alcohol and chloroform for 2 min, and then stored in xylene or toluene for at least 3 min, or up to 2 wk until processed further. Mounting is done on glass slides which have been coated with fresh egg albumen diluted in 0.2% ammonia water (or a 0.5% solution of dry powdered egg albumen) and then dried at 60°C overnight. For attachment to these coated slides, sections are first soaked for 2-3 min in a freshly prepared mixture of methyl benzoate, 50 ml; benzyl alcohol, 200 ml; chloroform, 150 ml; and then transferred quickly to the slides by means of a brush. After 2-3 min the chloroform evaporates and the celloidin softens. The slides are then immersed in toluene which hardens the celloidin and anchors the sections to the slides. Alcohols of descending concentrations to 40% are followed by alkalinizations, first in: absolute alcohol, 40 ml; strong ammonia water 60 ml, for 2 min, then in: absolute alcohol, 70 ml; strong ammonia water, 30 ml, for 1 hr. Excess alkali is then removed by 70% and 40% alcohol, 2 min each, and a 10 min wash in running tap water. Bleaching in 1% Na₂S₂O₃, for 10 min and washing again in tap water for 10 min completes the process preliminary to staining. The preparations are then stained for 90 min in an aqueous solution of either 0.5% cresylecht violet, neutral red, or Darrow red, buffered at pH 3.6. Dehydration and differentiation in ascending grades of alcohol, clearing with toluene or xylene, and applying a cover glass with a mounting medium having a refractive index of about 1.61 completes the process.     

Notes on Technic: A Device for Centering Tissue Blocks on the Porter-Blum Microtome

A method of double embedding fixed tissues in 3% low viscosity nitrocellulose and paraffin is described. Five percent phenol in 80% alcohol during dehydration and 5% glycerin in the nitrocellulose solutions enhance cutting qualities. A modified Ruyter's solution is used to flatten sections. After a section is aflixed to a slide, it is passed through chloroform and acetone to remove the paraftin and celloidin. A 1% celloidin dip insures adherence of the seaion to the slide. Slides are stored in 70% alcohol until they are to be stained. Following staining and dehydration in graded alcohols, clearing should be done in a 1: 3 mixture of terpineol and toluene.     

Sectioning Insects with Sclerotized Cuticle

Adult insects of different orders including beetles were fixed in a mixture of a saturated solution of picric acid in 90% alcohol, 75 parts; formalin, 25 parts; nitric acid (cone), 8 parts, 4-6 days and even up to 10 days depending upon the hardness of the cuticle (addition of 5% mercuric chloride to this mixture is recommended when prolonged immersion is required), or in Carnoy and Lebruns' fluid 24-48 hours and then transferred to a solution of 3-6 parts of nitric acid in 100 parts of 90% alcohol (3-6 days). After dehydration in different grades of alcohol, the insects were double embedded in celloidin and paraffin, either by (1) clove oil for 1 day, then to a saturated solution of celloidin in clove oil matured for at least 2 months for 20-40 days, or (2) the conventional ether-alcohol-celloidin mixture for 7 days; followed by hardening in chloroform. The difficulty in the proper infiltration of paraffin into celloidin hardened by chloroform around the insect is avoided by keeping the block overnight in a mixture of paraffin, 1 part; chloroform, 5 parts. The rest of the technic is essentially the same as that followed in cutting sections after double embedding.     

New Fuchsin-Hematoxylin-Eosin; A Stain for Acid-Fast Bacilli and Surrounding Tissue

This is a staining technique for histopathologic evaluation of tissue reaction in the environs of acid-fast tubercle bacilli (avian and bovine) in sections. Fresh tissue is fixed in 10% neutral formalin and processed in the usual manner for embedding in paraffin. Sections are cut approximately 6 μ. thick, dewaxed, hydrated, and stained with Harris' hematoxylin. They are rinsed in tap water, differentiated in add alcohol, washed in tap water, given a distilled water rinse and stained at 20-30° C in a 1% solution of new fuchsin in 5% phenol. Each slide is then handled individually by placing it directly into a saturated aqueous solution of Li₂CO₃ and agitated gently for a few seconds. This is followed by differentiation with 5% glacial acetic acid in absolute or 95% ethyl alcohol until the color stops running. Two rinses in absolute or 95% ethyl alcohol follow. The sections are then counterstained in the color add of eosin Y prepared according to the method of Schleicher (Stain Techn., 28, 119-23, 1953) and used as an 0.025% solution in absolute alcohol. Following passage through 2 changes of absolute alcohol, the sections are cleared in xylene, then mounted in Permount or similar synthetic resin. The add-fast barilli are emphasized by their bright retractile red color within a contrasting background of hematoxylin and eosin.     

Double Embedding Again

A method of double embedding fixed tissues in 3% low viscosity nitrocellulose and paraffin is described. Five percent phenol in 80% alcohol during dehydration and 5% glycerin in the nitrocellulose solutions enhance cutting qualities. A modified Ruyter's solution is used to flatten sections. After a section is aflixed to a slide, it is passed through chloroform and acetone to remove the paraftin and celloidin. A 1% celloidin dip insures adherence of the seaion to the slide. Slides are stored in 70% alcohol until they are to be stained. Following staining and dehydration in graded alcohols, clearing should be done in a 1: 3 mixture of terpineol and toluene.     

Pasternack's Paraffin Method Modified for Plant Tissue

This method for preparing paraffin sections of plant material is a modification of Pasternack's one-hour method for animal tissues. Fixation in Randolph's CRAF fixative is hastened by heat and increased vapor pressure obtained by the use of screw top vials. Dehydration with Zirkle's butyl alcohol series likewise is hastened in the same manner. The rapid penetration of paraffin by the use of 1/2 paraffin and 1/2 butyl alcohol in heated screw top vials shortens the embedding process. Sections are held on the slide thru staining by albumen fixative and a coating of 0.2% celloidin in absolute alcohol and ether. Good penetration with freedom from shrinkage or distortion is obtained and root tip chromosome counts can be made in approximately 3 hours.     

Hematein-its Advantages for General Laboratory Usage

The advantages of hematein over hematoxylin are that it is easy to prepare, easy to use, and saves time; while it gives equally good results. The writer has been employing for some time a pre-war imported hematein and until within the last few months has been unable to locate a satisfactory product of recent manufacture, either domestic or foreign. At the request of the Biological Stain Commission, however, an American manufacturer has at last put on the market a C. P. hematein which gives splendid results. The technic is as follows:

Paraffin or celloidin sections of Bouin or Zenker-formol material are run down to water and stained about 5 minutes in Mayer's hemalum (0.5 g. hematein ground up in a glass mortar with 10 cc. 95% alcohol and added to 500 cc. of 5% aqu. sol. potassium alum.) Rinse 1 to 3 seconds in tap water. Dip 1 to 3 seconds in eosin B (1 part 0.5% sol. in 20% alc. added to 2 pats dist. water; filtered from time to time). Wash several minutes in running water or in several changes of tap water. Dehydrate and mount; with unattached celloidin sections this may be done by running up to 95% alcohol, spreading on slide, blotting, wetting with absolute alcohol, draining and mounting in euparal.     

Staining and Mounting Helminths

The following procedure is recommended: Fix ces-todes and trematodes (while held flat between glass slides) 0.5–2.0 hr. in the following mixture: formalin, 15; acetic acid (gl.), 5; glycerol, 10; 95% ethyl alcohol, 24; distilled H₂O, 46; all proportions by volume. After freeing them from the slides, wash thoroughly in running water and stain immediately thereafter. Stock staining solution: ferric ammonium alum (violet cryst.), 2 g.; distilled H₂O (cold) 100 ml.; after solution, add 2 ml. concentrated H₂SO₄, bring to a boil; add 1 g. coelestin blue B (Nat. Aniline), boil 3–5 min.; cool and add 10 ml. absolute methyl alcohol and 10 ml. glycerol. Dilute 1 vol. with 3 vol. distilled H20 for use. Stain 5–30 min., depending on size of specimens. Wash with 2 changes 0.5 hr. each of distilled H₂O, then 50% isopropyl alcohol 12–16 hr., 50% isopropyl alcohol 2 hr., followed by graded isopropyl alcohol for dehydration. Ether: ethyl alcohol (equal parts), 1 hr., is followed by embedding in celloidin in a sheet just thick enough to cover the specimens. Trim embedded specimens and dehydrate with isopropyl alcohol, 80%, 90% and absolute. Clear in beechwood creosote. Mount in balsam with cover glasses that overlap the edges of the celloidin 1–2 mm. While drying at 37°C, refill edges of mount with fresh balsam as needed. When dry, remove excess balsam and ring the edges with ordinary gloss enamel paint.     

New Fuchsin-Hematoxylin-Eosin; A Stain for Acid-Fast Bacilli and Surrounding Tissue

This is a staining technique for histopathologic evaluation of tissue reaction in the environs of acid-fast tubercle bacilli (avian and bovine) in sections. Fresh tissue is fixed in 10% neutral formalin and processed in the usual manner for embedding in paraffin. Sections are cut approximately 6 μ. thick, dewaxed, hydrated, and stained with Harris' hematoxylin. They are rinsed in tap water, differentiated in add alcohol, washed in tap water, given a distilled water rinse and stained at 20-30° C in a 1% solution of new fuchsin in 5% phenol. Each slide is then handled individually by placing it directly into a saturated aqueous solution of Li₂CO₃ and agitated gently for a few seconds. This is followed by differentiation with 5% glacial acetic acid in absolute or 95% ethyl alcohol until the color stops running. Two rinses in absolute or 95% ethyl alcohol follow. The sections are then counterstained in the color add of eosin Y prepared according to the method of Schleicher (Stain Techn., 28, 119-23, 1953) and used as an 0.025% solution in absolute alcohol. Following passage through 2 changes of absolute alcohol, the sections are cleared in xylene, then mounted in Permount or similar synthetic resin. The add-fast barilli are emphasized by their bright retractile red color within a contrasting background of hematoxylin and eosin.     

The Surface Staining of Embedded Tissues

The staining procedure is based on the theory that the freshly cut surface of embedded material will absorb stain only in the exposed tissue elements, provided that the embedding compound itself will not absorb the staining fluid. Concentrated stains are used for short intervals to insure minimum penetration. For paraffin embedded materials: (1) Cut block, preferably on microtome, to the desired tissue surface. (2) Rinse in absolute alcohol. (3) Float face down in stain. (Ripe, concentrated alum hematoxylin—Galigher's formula recommended—will stain in 10 to IS minutes. Heidenhain's iron hematoxylin works exceptionally well in some cases.) Mordant 20% alum 5 to 10 minutes, briefly rinse, and stain comparable 5 to 10 minutes in 1 to 1.5% hematoxylin. (4) Allow to become blue in tap water (for hematoxylin stains). (5) Counter-stain if desired. (6) Dehydrate in absolute alcohol for not more than 10 minutes. (7) Dry for 15 to 20 minutes. (8) Trim block to 2-3 mm. and mount between two cover glasses by use of microflame. Attach mount to slide with balsam. For celloidin embedded materials: (1) Dehydrate block with 90% alcohol, phenol-toluene, finally pure toluene. (2) Rinse cut surface with 90% alcohol, then apply stain. (3) Wash, after hematoxylin stains, counterstain if desired. (4) Dehydrate surface, 90% alcohol, phenol toluene, pure toluene, and mount in medium dissolved in toluene.

Possible applications of surface staining technic are suggested and illustrated.     

Plastic Embedding of thick Celloidin Sections of nerve Tissue

A method is described for mounting Golgi-impregnated and Weigert-stained thick celloidin sections of brain and spinal cord in transparent plastic. Finished mounts have good optical properties and are suitable for macroscopic and microscopic observation. The durability of such preparations makes them superior to similar material prepared by the more conventional methods. Holes of suitable size were cut in matrices of 2.5 × 5 × 3/16 inches Plexiglas. Ward's Bio-plastic was used to form a base for the holes and also as the embedding medium for the sections. Plate glass formed a working substrate and gave a polished surface to the plastic base and later to the top of the preparation. For Golgi material (200μ) the celloidin was removed by dioxane. A dioxane-plastic bath preceded plastic embedding. For Weigert material (30-40μ) celloidin was not removed due to fragility of sections. Prior to plastic embedding, they were subjected first to benzol and then to a benzol-plastic bath.     

Staining and Mounting Helminths

The following procedure is recommended: Fix ces-todes and trematodes (while held flat between glass slides) 0.5-2.0 hr. in the following mixture: formalin, 15; acetic acid (gl.), 5; glycerol, 10; 95% ethyl alcohol, 24; distilled H₂O, 46; all proportions by volume. After freeing them from the slides, wash thoroughly in running water and stain immediately thereafter. Stock staining solution: ferric ammonium alum (violet cryst.), 2 g.; distilled H₂O (cold) 100 ml.; after solution, add 2 ml. concentrated H₂SO₄, bring to a boil; add 1 g. coelestin blue B (Nat. Aniline), boil 3-5 min.; cool and add 10 ml. absolute methyl alcohol and 10 ml. glycerol. Dilute 1 vol. with 3 vol. distilled H20 for use. Stain 5-30 min., depending on size of specimens. Wash with 2 changes 0.5 hr. each of distilled H₂O, then 50% isopropyl alcohol 12-16 hr., 50% isopropyl alcohol 2 hr., followed by graded isopropyl alcohol for dehydration. Ether: ethyl alcohol (equal parts), 1 hr., is followed by embedding in celloidin in a sheet just thick enough to cover the specimens. Trim embedded specimens and dehydrate with isopropyl alcohol, 80%, 90% and absolute. Clear in beechwood creosote. Mount in balsam with cover glasses that overlap the edges of the celloidin 1-2 mm. While drying at 37°C, refill edges of mount with fresh balsam as needed. When dry, remove excess balsam and ring the edges with ordinary gloss enamel paint.     

A Modified Polyester Embedding Medium for Sectioning

A modified polyester resin designated as C. M. E. Tissue Support Resin can be cut on a rotary microtome and can yield sections from 5 to 50μ from tissue blocks that range from 5 to 16 mm in diameter. It is firm enough to support hard structures that lie adjacent to soft ones and retain all in their normal position. The resin-catalyst-promoter system cures oi hardens at a low temperature so that blocks are ready for cutting 6 hr aftei tissue has been routinely dehydrated. The resin is compounded from a plasticized rigid polyester and adjusted to a proper viscosity. Several grades of hardness can be attained by changing the formula. It has been tested with both soft and hard tissues, including limbs and tails of 7-mo old mice and mature whole grains of wheat, and provides a more substantial and more readily prepared embedding medium than celloidin. Sections can be stained before mounting, without removing the embedding material, with aqueous safranin O followed by fast green FCF in absolute alcohol. The plastic remains clear. Other staining processes require modifications to get good results.     

The Histological Examination of Mummified Material

Tissue from Egyptian mummy material is extremely brittle; hence it was handled in perforated glass tubes during processing. The first (softening) fluid consisted of 96% ethyl alcohol, 30 vol; 1% aqueous formalin, 50 vol; 5% aqueous Na₂CO₃, 20 vol. It was used in a fluid to tissue volume ratio of 100:1 and allowed to act overnight. A special dehydrating sequence: 80% alcohol, 3-6 hr; 8% phenol in 96% alcohol, and absolute alcohol followed by 3 changes of amyl acetate, 6-18 hr each; 3 changes of 1 % celloidin in methyl benzoate, 24 hr each; then through benzene and embedding in paraffin completed the special technic. This allowed regular sectioning and staining to be done successfully.     

Embedding and Sectioning Undecalcified Bone, and its Application to Radioautography

A method is described for embedding and sectioning hard, undecalcified bone, which is designed for use by technical personnel. Bone fixed in a variety of ways is progressed through alcohols to ether-alcohol and then infiltrated with ether-alcohol solvented plastic (plasticized nitrocellulose) by a combination of centrifugation and high pressure embedding technics. The ether-alcohol is evaporated in a partially closed container in a manner similar to that employed in celloidin embedding, but differs from the latter by the removal of all of the solvent. Celloidin is the source of nitrocellulose and Amoil-S, the added plasticizer. Undecalcified adult bone of all types is readily cut at a thickness of 5-8μ on a heavy duty sliding microtome. The sections are then mounted on gelatinized slides. The procedures for preparing strip film radio-autograms of bone sections and subsequent staining of the preparation are given. The results obtained are illustrated.     

A Staining Combination for Phloem and Contiguous Tissues

A technic is described for producing critically stained preparations of phloem tissue. The preparations promise to be relatively stable. Sections of fixed unembedded or of embedded (paraffin or celloidin) phloem, cambium, and xylem are (1) stained in Foster's tannic acid-ferric chloride combination; (2) treated with 1% NaHCOg in 25% or 50% ethyl alcohol for 30 minutes; (3) stained in a saturated solution of lacmoid (made alkaline by adding a few ml. of 1% NaHCO₃ in 25% alcohol) for 12 to 18 hours; (4) dehydrated and cleared in a series composed of 1% solution of NaHCO_s in 50% ethyl alcohol, 80%, 95%, and absolute alcohol, equal proportions of absolute alcohol, clove oil, and xylene, and finally pure xylene; and (5) mounted in a neutral resin. Callose and lignified secondary walls are blue or blue-green in color, cellulose walls and stainable protoplasmic contents are generally light brown. The technic has been successful with sections from 5 to 40μ in thickness, and the staining has been satisfactory for both color and black and white photomicrography.     

Affixing Polyester Wax Sections to Glass Slides by Celloidin and Cellulose Acetate Applied in Sequence

Sections cut from material embedded in polyester wax can be firmly attached as follows: One drop of a 2% solution of celloidin in amyl acetate is smeared on clean slides, and sections taken from the floatation water onto these slides are dried at room temperature. After drying the slides are immersed in a 2% solution of cellulose acetate in acetone for 1 min, transferred directly to absolute ethanol, through 50% ethanol, and into water. Sections affixed by this method and stained by either hematoxylin-eosin or toluidine blue schedules do not loosen and have negligible background staining.