A Tetrachrome Stain for Fresh, Mineralized Bone Sections, Useful in the Diagnosis of Bone Diseases

Fresh, unprocessed bone is ground to sections 75-100 μ thick, stained in an aqueous solution composed of fast green FCF, 0.1 gm; orange G, 2.0 gm; distilled water, 100.0 ml; and adjusted to pH 6.65, then in a mixture of 1 part alcoholic solution of 0.25% celestine blue B and 9 parts of alcoholic solution of 0.1% basic fuchsin. Surface stain is removed by grinding sections to 50 μ and washing them in 1% invert soap (Zephiran) to remove adherent debris. (Commercial detergents and alkaline soaps may interfere with chromophore groups of the dyes.) Wash in tap water; rinse in distilled water and differentiate in 1% acetic alcohol. Dehydrate in ascending alcohols, clear in xylene and mount permanently in a neutral, synthetic resin. Active osteoid seams stain dark to light green; resting osteoid seams, red to bright orange red; transitional osteoid seams, geenish-yellow, orange red to red; older, partly mineralized matrix, orange; new, partly mineralized matrix, red; osteocyte nuclei, red; osteoblasts and osteoclasts, greenish-blue to dark purple nuclei and green or light green cytoplasm. Hyper-trophic and differentiating cartilage cells are stained light pink and dark red respectively. The staining reactions are consistent; the solutions are stable.     

The Differentiation of Live From Dead Sperms in Fowl Semen

A combined stain solution is made by dissolving 0.1 gm bromphenol blue and 0.2 gm nigrosin in 100 ml of a M/15 buffer solution of KH₂PO₄ and Na₂HPO₄ adjusted to pH 7.5. This staining solution was used to prepare stained fowl semen smears. Such smears give stable differentiation of live from dead sperms. The dead sperms are stained with a dark violet color while the live ones are not stained.     

Oxazine Dyes. 2. Celestine Blue B, Gallocyanin and Gallamin Blue with Mordants Other Than Ferric Alum

Experiments with 3 oxazine dyes with 19 mordants failed to produce a more satisfactory staining solution than that recorded by Gray et al. (1956). The most practical solution developed is prepared by adding 0.4 ml concentrated HNO₃ to 1gm celestine blue B and dissolving the resulting mass in 100 ml 5% cupric nitrate containing 14 ml glycerol. This solution is less acid (pH 1.4) than the celestine blue B-ferric alum solution (pH 0.8) previously recommended. It gives as intense and sharp a stain but is slightly less stable. Formulae are given for four other combinations of possible practical application.     

Oxazine Dyes. 3. Simultaneous Nuclear and Cytoplasmic Staining from Single Solutions

Of 84 dyes tested, 26 were found to give a stable solution with celestine blue B dispersions which simultaneously stained nuclei and cytoplasm. The cytoplasmic dye is dissolved in celestine blue B dispersion prepared by the method of Cray et al. (1956). Croceine scarlet (C.I. 286), in the proportion of 0.38 gm to 114 ml of celestine blue B dispersion, gives results strikingly similar to hematoxylin-eosin when used for 2 min on a wide variety of tissues. No differentiation, other than that which occurs during dehydration, is necessary.     

Oxazine Dyes. I. Celestine Blue B with Iron as a Nuclear Stain

It is suggested that celestine blue B can stain as a colloidal dispersion, the nuclear specificity of which is controlled by the pH. The staining solution is prepared by adding 0.5 ml of concentrated H₂SO₄ to 1 gm of celestine blue B and dissolving the resultant granular mass in 100 ml of 2.5% ferric alum containing 14 ml of glycerol. Sections of amphibian, avian, and mammalian tissue placed for 1 min in this solution and then rinsed in water show as sharp nuclear staining as that usually produced by hematoxylin. A wide variety of fixatives is permissible. Overstaining is not possible within reasonable limits of exposure and no differentiation nor bluing is required. Both the staining solution and stained slides are stable.     

Improving the Staining Qualities of Toluidin Blue

To a 1.0% filtered aqueous solution of toluidin blue add drop by drop 4-5 ml of either a saturated aqueous solution of HgCl₂ or of KI. Collect the resulting dark precipitate on a filter paper and wash it with numerous small quantities of distilled water applied to both inside and outside of the filter paper. Wash until the drippings are distinctly blue (equivalent to about a 0.05% dye solution). Remove the paper and its contents from the funnel and dry either at room temperature or at 37°C. When dry, the treated dye can be brushed off the paper and stored. To prepare a staining solution add a weighed amount (0.12 gm if derived from the HgCl₂ treatment, or 0.3 gm if from KI) to 100 ml of distilled water. This insures a saturated solution in either case and gives a satisfactory stain with most sections in 10-30 min. Thionin and other members of the thiazine dyes also showed improvement in staining qualities after this treatment.     

Staining Tissue of the Central Nervous System with Luxol Fast Blue and Neutral Red

The tissue is fixed in 10% neutral saline formalin for 1 day to 3 wk depending on the size of the block, dehydrated and embedded in paraffin. The sections are stained at 57° C for 2 hr, then at 22° C for 30 min, in a 0.0125% solution of Luxol fast blue in 95% alcohol acidified by 0.1% acetic acid. They are differentiated in a solution consisting of: Li₂CO₃, 5.0 gm; LiOH-H₂O, 0.01 gm; and distilled water, 1 liter at 0-1° C, followed by 70% alcohol, and then treated with 0.2% NaHSO₃. They are soaked 1 min in an acetic acid-sodium acetate buffer 0.1 N, pH 5.6, then stained with 0.03% buffered aqueous neutral red. Sections are washed in distilled water, 1 sec, then treated with the following solution: CuSO₄·5H₂O, 0.5 gm; CrK(SO₄)₂·12H₂O, 0.5 gm; 10% acetic acid, 3 ml; and distilled water, 250 ml. Dehydration, clearing and covering complete the process. Myelin sheaths are stained bright blue; meninges and the adventitia of blood vessels are blue; red blood cells are green. Nissl material is stained brilliant red; axon hillocks, axis cylinders, ependyma, nuclei and some cytoplasm of neuroglia, media and endothelium of blood vessels are pink.     

Specific Staining of Glycogen with Haematoxylin and Certain Anthraquinone Dyes

Specific staining of glycogen in rat liver fixed in chilled 80% alcohol, chilled formol alcohol or 10% neutral formalin has been accomplished with acid alizarin blue SWR, alizarin brilliant blue BS, alizarin red S, gallein, haematein, and haematoxylin solutions. TO prepare a staining solution, 1 gm dye, 1 gm K₂CO₃ and 5 gm KCl were dissolved by heating in 60 ml of water. Concentrated NH₄OH (0.880 sp.gr.), 15 ml, followed by 15 ml of dry methanol were added to 20 ml of the cooled solution. Paraffi sections were stained for 5 min, rinsed in dry methanol, cleared in xylene, and mounted in D.P.X. The high specificity obviated the need for counterstaining: nuclei and cytoplasm were unstained. Precipitation of stain onto the slide was rare. As all the dyes carried, like carminic acid, numerous groups capable of forming hydrogen bonds, it is suggested that the staining mechanism involved hydrogen bonding.     

Mercuric Bromphenol Blue Staining of Precipitin Lines in Agar

Lines formed by antibody-organ antigen reactions are stained particularly well by a modification utilizing the mercuric bromphenol blue (MBB) mixture of Mazia et al. (Biol. Bull., 104: 57-67, 1953). The agar covered slides are placed overnight in 0.85% NaCI at 4 C, followed by washing for 2 hr in 0.85% NaCI at 25 C. They are then rinsed for 10 min in distilled water, and dried overnight at 37 C. The precipitin lines are fixed by immersing the slides for 25 min in 95% alcohol, followed by 5 min hydration in distilled water. They are stained for 25 min in MBB mixture (HgCI₂, 10 gm; bromphenol blue, 0.1 gm; 95% ethanol, 100 ml). Excess stain is removed by immersing in acidified alcohol (95% ethanol, 98 ml; glacial acetic acid, 2 ml). Finally, the slides are passed through alcohol and xylene, and resin-mounted under coverslips.     

Acid alcoholic brilliant cresyl blue stain for gastric and other mucins

Summary Some but not all samples of brilliant cresyl blue (6-methyl-7-dimethylamino-2-phenoxazin chloride) under C. I. No. 51010 in Conn's Biological Stains when dissolved at 1% level in 50–70% alcohol containing 1% concentrated (12 N) hydrochloric acid, stain (in 30 min) a wide variety of human and laboratory animal mucins blue black on an almost unstained background. The mucoprotein of the gastric surface epithelium and of the peptic gland neck cells of several species reacts strongly. A 16 hr 60° C methylation in 0.1 M methyl-sulfuric acid in methanol is required to block the staining of these gastric and some intestinal mucins, while 1–2 hr intervals suffice to prevent the staining of mast cells, cartilage and metachromatic sulfomucins generally. Saponification (1% KOH/70% alcohol, 20min) does not restore staining in either location group, indicating that sulfate mucins are probably reacting in both.Most other basic dyes fail to stain mucins from acid alcohol solutions: azure A, toluidine blue, resorcin blue, orcein, resorufin, azoresorufin brown, azolitmin, lacmoid, gallocyanin, Nile blue, methylene green, pararosanilin, crystal violet, Victoria blue R. Some staining occurred with one of three lots of Victoria blue B, with two lots of Victoria blue 4 R and with one lot each of Bernthsen's methylene violet, elastin violet PR and elastin purple PP.The stain may be preceded by the Feulgen reaction to give red nuclei, or followed by a brief collagen stain in an alcoholic acid fuchsin (0.05–0.1%), picric acid (1.5%) solution.Presented before the Symposium of the Histochemische Gesellschaft in Hamburg, 28. September 1968.Supported by National Cancer Institute Grant No. C-4816, National Institutes of Health.     

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.     

Oxazine Dyes. Iv. Simultaneous Nuclear and Double-Contrast Cytoplasmic Staining from A Single Solution

A dye mixture, consisting of a celestine blue B dispersion (prepared according to Gray et al. 1956), orange G, and acid fuchsin in one solution, simultaneously stains nuclear elements and gives double contrast staining of cytoplasmic elements. Orange G, 0.16 gm, and acid fuchsin, 0.04 gm, dissolved in 100 ml of celestine blue B dispersion and adjusted to pH 0.8 gives, when applied for 1.5 min, results comparable or superior to other “triple contrast” stains on a wide variety of tissues. No differentiation other than that which occurs during dehydration is necessary.     

A Selective Stain for Elastic Tissue (Orcinol-New Fuchsin)

A selective stain for elastic tissue (designated orcinol-new fuchsin) is described. Two grams of new fuchsin (C.I. No. 678) and 4 gm of orcinol (highest purity) are added to 200 ml of distilled water and the solution boiled for 5 min. Then 25 ml ferric chloride solution (U.S.P. IX) are added and the solution is boiled 5 min longer. The precipitate is collected and dissolved in 100 ml 95% ethanol. This is the staining solution. Sections are deparaffinized and brought to absolute ethanol, stained for 15 min at 37 °C with orcinol-new fuchsin, differentiated for 15 min in 70% ethanol, dehydrated, cleared and covered as usual.     

Chlorazol Fast Pink B And Trypan Blue Tests for Virus and Other Proteinaceous Inclusions

A 1% solution of chlorazol fast pink B in 0.9% NaCl can be used like trypan blue to detect virus inclusions and proteinaceous entities in peelings from leaves or thin sections taken from living plant tissue. Like trypan blue, a solution of the pink dye causes somatic nuclei to swell and thus facilitates observation of their structure. The two dyes combine into a beautiful differential bicolored stain. Mix 5 ml of 0.5% trypan blue stock solution with 35 ml of 1% chlorazol pink B in 0.9% NaCl. Stain fresh tissue 1-2 minutes. The combination stain is superior to either dye alone for differentiating virus entities.     

A Formalin-Wright Staining Technique for Avian Blood Cells

A rather concentrated alcoholic staining solution, an aqueous formalin-containing diluent, and a mixture of ethyl ether and absolute methyl alcohol are required. Formulas: A. Wright's stain (Harleco, Cert. No. LWr-52 was used), 3.3 gm; methyl alcohol, 500 ml. B. Formaldehyde solution 40% USP (Fisher's used), 0.25 ml; distilled water, 500 ml with its pH adjusted to 6.8 by addition of either 0.25% Na₂CO₂ or 0.25% HCl, as needed. C. A I:I mixture of ethyl ether and absolute methyl alcohol. Procedure: Prepare thin smears of normal or pathological avian blood, air dry, place the slides on a drying rack, cover with solution A, and let stand for about 8 min. Dilute the stain by dropping on a volume of B estimated to be equal to the volume of the partially evaporated stain, and let stand for 2-5 min, or until the surface is well covered by a metallic sheen. Wash with distilled water adjusted to pH 6.8 with the 0.25% Na₂CO₂ solution or 0.25% HCl. Dry the preparations quickly by blotting with filter paper. Differentiate and adjust the color intensities by dipping 6-10 times into C. Check the results microscopically and differentiate further if the colors are not properly balanced. Dry, uncovered preparations may be examined under oil; or, a cover glass can be applied with balsam or a synthetic resin for permanent mount. Results are similar to those described in textbooks, but have been more consistent than those obtained with other techniques for blood cells of chicken, pheasants, American and Indian partridge, quail, pigeon, turkey, goose, canary, and the Himalayan snow partridge.     

A Mixture of Methyl Green and Pyronin for Cell Fractionation Studies

A staining mixture consisting of 0.57% methyl green and 0.1 1 % pyronin B (calculated from the actual dye content) dis- solved in glycerol, 20 ml.; 2% aqueous phenol, 100 ml.; and 95% ethanol, 25 ml., was found to be optimum for differentiating cell components containing desoxypentose and pentose nucleic acids. The stain can be used for either fresh suspensions or unfixed dried smears of tissue homogenates. Nuclei are stained bright blue, and nucleoli and cytoplasmic particles, bright pink.     

Zinc Chloride Methylene Blue, Ii. Preparation of Giesma and Wright Type Low Azure B Blood Stains from Dichromate Oxidized Commercial Dye

TO determine the amount of K₂Cr₂O₇ required to produce optimal Giemsa type staining, six 1 g amounts (corrected for dye content) of zinc methylene blue were oxidized with graded quantities of K₂Cr₂O₇ to produce 4, 8, 12, 16, 20 and 24% conversion of methylene blue to azure B. These were heated with a blank control 15 minutes at 100 C in 60-65 ml 0.4 N HCI. cooled, and adjusted to 50 ml to give 20 mg original dye/ml. Aliquots were then diluted to 1% and stains were made by the “Wet Giemsa” technic (Lillie and Donaldson 1979) using 6 ml 1% polychrome methylene blue, 4 ml 1% cosin (corrected for dye content), 2 ml 0.1 M pH 6.3 phosphate buffer, 5 ml acetone, and 23 ml distilled water. The main is added last and methanol fixed blood films are stained immediately for 20-40 min.

For methylene blue supplied by MCB 12-H-29, optimal stains were obtained with preparations containing 20 and 24% conversion of methylene blue to azure B. With methylene blue supplied by Aldrich (080787), 16% conversion of methylene blue to azure B was optimal. Eosinates prepared from a low azure B/methylene blue preparation selected in this way give good stains when used as a Wright stain in 0.3% methanol solution. However, when the 600 mg eosinate solution in glycerol methanol is supplemented with 160 mg of the same azure B/methylene blue chloride the mixture fails to perform well. The HCI precipitation of the chloride apparently produces the zinc methylene blue chloride salt which is poorly soluble in alcohol. It appears necessary to have a zinc-free azure B/methylene blue chloride to supplement the probably zinc-free eosinate used in the Giemsa mixture.     

Staining Residual Lipids in Ultrathin Sections of Tissues Embedded in Polyester Resin

Rat suprarenal glands fixed in Palade's 1% OsO₄, buffered at pH 7.7 with veronal-acetate, to which 0.1% MgCl₂ was added, were embedded in Vestopal-W and sectioned at 0.2-1 µ. The sections were attached to slides by floating on water, without adhesive, and drying at 60-80° C, placed in acetone for 1 min and then treated with the following staining procedure: Place the preparation in a filtered solution of oil red O, 1 gm; 70% alcohol, 50 ml; and acetone, C.P., 50 ml; for 0.5-1 hr. Rinse in absolute ethyl alcohol; drain; counterstain with 0.5% aqueous thionin for 5 min; rinse in distilled water; drain; stain in 0.2% azure B in phosphate buffer at pH 9, for 5 min. Dry and apply a drop of immersion oil directly on the section. The preparations are temporary. Ciaccio-positive lipids, rendered insoluble by OsO, fixation, stained red to ochre.     

Luxol Fast Blue Mbs: A Stain for Phase Contrast Microscopy

A staining method to increase the contrast of sectioned material for phase contrast microscopy is described. Two stock solutions of the stain are required. The first is made by dissolving 2 gm of luxol fast blue MBS in 100 ml of 95% ethanol. The second solution is made up of 4 ml of a 29% aqueous solution of FeCl₃, 95 ml of 95% ethanol, and 1 ml of concentrated HCl. The staining solution is made by mixing equal parts of the two solutions. Sections are deparaffinized and taken to 70% alcohol, stained for 1.5 hr, dehydrated, cleared and covered as usual.     

Removal of Ribonucleic Acid from Bacterial Cells by Dilute Salt Solutions

Bacterial cells were impressed upon a clean glass slide, fixed in ethyl alcohol and immersed at 37°C in either of the following two salt solutions: (A) NaCl, 7.8 gm; KCl, 0.7 gm; distilled water, 1000 ml; adjusted to pH 7.0; or (B) 0.1M NaH₂PO₄, 400 ml; 0.1M Na₂HPO₄, 600 ml; KCl, 0.7 gm. After 1-5 hr soaking to remove ribonucleic acid, the slide was stained by Giemsa's method as usual. The staining revealed slender chromatinic bodies with reasonable clarity extending the whole diameter of the moderately swollen cell. The results of this method seemed to be much like those obtained after ribonuclease digestion.