On the mechanism of Verhoeff's elastica stain: a convenient stain for myelin sheaths.

Verhoeff (1908) recommended an iron-hematein formula containing Lugol's solution for demonstration of elastic tissue; sections are differentiated until desired staining patterns are obtained. Verhoeff's stain colored a variety of tissue structures and showed higher substantivity for myelin sheaths than for elastin. Addition of HCL or omission of Lugol's solution decreased or abolished coloration of pseudo-elastica and thus enhanced selectivity for elastin. Substitution of Fe++ for Fe+++ abolished dye binding by elastin. A review of chemical data indicated interaction of components of Lugol's solution with the dye. Hematein and Fe+++ form a variety of cationic, anionic and non-ionic chelates; the ratio of these compounds changes with time. Dye binding apparently occurs mainly via van der Waals forces and hydrogen bonds. Verhoeff's elastica stain is definitely not specific for elastin and is inferior to orcein and resorcin-fuchsin because of the required differentiation with its inherent bias to produce patterns which conform to expectations. However, Verhoeff's elastica stain is far superior to other metal-hematein technics for myelin sheaths. The combined Verhoeff-picro-Sirius Red F3BA stain can be performed in 30 min and does not require differentiation. It is therefore suggested to reclassify Verhoeff's elastica stain as a method for myelin sheaths.     

On the mechanism of Verhoeff's elastica stain: A convenient stain for myelin sheaths

Summary Verhoeff (1908) recommended an iron-hematein formula containing Lugol's solution for demonstration of elastic tissue; sections are differentiated until desired staining patterns are obtained. Verhoeff's stain colored a variety of tissue structures and showed higher substantivity for myelin sheaths than for elastin. Addition of HCL or omission of Lugol's solution decreased or abolished coloration of pseudo-elastica and thus enhanced selectivity for elastin. Substitution of Fe⁺⁺ for Fe⁺⁺⁺ abolished dye binding by elastin.A review of chemical data indicated interaction of components of Lugol's solution with the dye. Hematein and Fe⁺⁺⁺ form a variety of cationic, anionic and non-ionic chelates; the ratio of these compounds changes with time. Dye binding apparently occurs mainly via van der Waals forces and hydrogen bonds.Verhoeff's elastica stain is definitely not specific for elastin and is inferior to orcein and resorcin-fuchsin because of the required differentiation with its inherent bias to produce patterns which conform to expectations. However, Verhoeff's elastica stain is far superior to other metal-hematein technics for myelin sheaths. The combined Verhoeff-picro-Sirius Red F3BA stain can be performed in 30 min and does not require differentiation. It is therefore suggested to reclassify Verhoeff's elastica stain as a method for myelin sheaths.     

Staining with chromoxane cyanine R.

Since Pearse in 1957 introduced chromoxane cyanine R as a dual nuclear and cytoplasmic stain there have appeared numerous procedures for use of this dye. These have differed widely, sharing in common mainly the implication that each is best. A defendable procedure has been developed on an experimental basis and is reported here. Four stock solutions are needed: (1) a 0.2% solution of chromoxane cyanine R in 0.5% aqueous H2SO4 (v/v); boil this solution for 5 min, (2) 10% FeCl3 in 3% HCl, (3) 1% aqueous HN4OH, and (4) 1% HCl in 70% ethanol. The staining solution: 40 ml of dye solution, 2 ml of FeCl3 solution, 8 ml H2O. Dewax and hydrate sections and stain for 10 min. If a myelin sheath stain is desired differentiate for 1 min in solution (3). For a nuclear stain differentiate for 1 min in solution (4). The nuclear stain when counterstained with eosin closely resembles the routine hematoxylin and eosin. Histochemical tests show that the functional group for myelin staining contains nitrogen, and probably hydrogen bonding is involved. The nuclear stain involves a different functional group and possibly neither electrostatic nor hydrogen bonding.     

Staining with Chromoxane Cyanine R

Since Pearse in 1957 introduced chromoxane cyanine R as a dual nuclear and cytoplasmic stain there have appeared numerous procedures for use of this dye. These have differed widely, sharing in common mainly the implication that each is best. A defendable procedure has been developed on an experimental basis and is reported here. Four stock solutions are needs. (1) a 0.2% solution of chromoxane cyanine R in 0.5% aqueous H₂SO₄ (v/v); boil this solution for 5 min, (2) 10% FeCl₃ in 3% HCI, (3) 1% aqueous NH₄OH, and (4) 1% HCI in 70% ethanol. The staining solution: 40 ml of dye solution, 2 ml of FeCl₃ solution, 8 ml H₂O. Dewax and hydrate sections and stain for 10 min. If a myelin sheath stain is desired differentiate for 1 min in solution (3). For a nuclear stain differentiate for 1 min in solution (4). The nuclear stain when counterstained with eosin closely resembles the routine hematoxylin and win. Histochemical tee show that the functional pup for myelin staining contains nitrogen, and probably hydrogen bonding is involved. The nuclear stain involves a different functional group and possibly neither electrostatic nor hydrogen bonding.     

Staining Myelin, Elastic Fibers and Nuclei with Iron Hematoxylin

Two iron hematoxylin staining procedures were developed. Both use stable stock solutions and can be prepared volumetrically. The nuclear stain is progressive but differentiation is required for myelin sheath and elastic tissue staining. Histochemical procedures demonstrated that acid, hydroxyl, and aldehyde groups play no role in the staining but amine groups are essential. With both types of stains neither electrostatic bonding nor hydrogen bonding is essential but the nature of the union between tissue and the iron hematoxylin complex was not determined.     

Modified Elastic Tissue-Masson Trichrome Stain

A combined elastic tissue-Massou technique is presented which stains elastic fibers of all sizes, nuclei and connective tissue. The modified elastic tissue stain consists of hematoxylin, ferric chloride and Verhoeffs iodine; nuclei and elastic fibers are stained blue-black in six minutes without differentiation. By contrast, cytoplasmic elements are stained red, (Biebrich scarlet-acid fuchsin) and collagen is stained green (light green) or blue (aniline blue). The entire staining procedure takes approximately one hour.     

Modified elastic tissue-Masson trichrome stain

A combined elastic tissue-Masson technique is presented which stains elastic fibers of all sizes, nuclei and connective tissue. The modified elastic tissue stain consists of hematoxylin, ferric chloride and Verhoeff's iodine; nuclei and elastic fibers are stained blue-black in six minutes without differentiation. By contrast, cytoplasmic elements are stained red, (Biebrich scarlet-acid fuchsin) and collagen is stained green (light green) or blue (aniline blue). The entire staining procedure takes approximately one hour.     

Quantitative histochemistry of myelin using Luxol Fast Blue MBS

Synopsis The amount of Luxol Fast Blue MBS in the band of Genarri was measured with two types of scanning microdensitometer and the optical density determined. The amount of stain measured was proportional to the section thickness employed, thus demonstrating that the dye has stoichiometric properties in tissue sections. Blocks of tissue treated with phospholipid solvents showed an increased uptake of stain, suggesting that phospholipids are not a primary substrate for the dye in myelin staining.The dye may, therefore, be used to quantify myelin in tissue sections.     

Standardized thionin-eosin stain in bronchial cytology. A substitute for hematoxylin-eosin Y staining

A standardized thionin-eosinic acid stain was developed as a quick and highly reproducible staining method for bronchial cytology. Bronchial smears and paraffin-embedded sputum samples were stained with thionin-eosin and with the conventional hematoxylin-eosin Y. Spectral absorption characteristics and staining intensity of thionin-eosin-stained cells were investigated by means of cytophotometry. The staining pattern of thionin-eosin is very close to that of the hematoxylin-eosin stain; the contrast between nucleus and cytoplasm is significantly higher for thionin-eosin. Thionin-eosin can be used for "dye-fixation" of cytologic smears and tissue imprints. Blueing and differentiation (as for hematoxylin-eosin) is not required for thionin-eosin; thus, fixation and staining can be performed within two minutes. The spectral absorption characteristics of thionin-eosin allow reliable automated cytophotometric discrimination of cell nuclei and cytoplasm. The standardized thionin-eosin stain is recommended as a substitute for the hematoxylin and eosin stain in bronchial cytology.     

Niagara Blue 4B As A Fast Simple Stain for the Adenohypophysis

For differentiation of cells of the adenohypophysis, the Niagara blue 4B method requires no special preliminary fixative nor very fresh tissue, and requires no more time than routine hematoxylin-eosin (H-E) staining. The method requires fixation in 10% formalin. After processing to paraffin wax, deparaffinise and hydrate the sections and stain in 1% aqueous Niagara blue 4B solution for 2 min. Stain afterwards with hematoxylin for 1 min then differentiate, wash, dehydrate, clear and mount. This method can be used also for staining old HE slides by removing the covers, applying the Niagara blue 4B and restaining with eosin. The Niagara blue 4B combined with H-E gives the best and most colorful result. This method allows special staining of the adenohypophysis from human post-mortem material to become routine.     

Detection of myelination using a novel histological probe.

Current methods for myelin staining in tissue sections include both histological and immunohistochemical techniques. Fluorescence immunohistochemistry, which uses antibodies against myelin components such as myelin basic protein, is often used because of the convenience for multiple labeling. To facilitate studies on myelin, this paper describes a quick and easy method for direct myelin staining in rodent and human tissues using novel near-infrared myelin (NIM) dyes that are comparable to other well-characterized histochemical reagents. The near-infrared fluorescence spectra of these probes allow fluorescent staining of tissue sections in multiple channels using visible light fluorophores commonly used in immunocytochemistry. These dyes have been used successfully to detect normal myelin structure and myelin loss in a mouse model of demyelination disease.     

Immunocytochemical localization of carbonic anhydrase in the spinal cords of normal and mutant (shiverer) adult mice with comparisons among fixation methods

The peroxidase-antiperoxidase technique was used for immunocytochemical localization of carbonic anhydrase in the mouse spinal cord to detect whether this antigen was normally present in myelinated fibers, in oligodendrocytes in both white and gray matter, and in astrocytes, and to determine where the carbonic anhydrase might be localized in the spinal cords of dysmyelinating mutant (shiverer) mice. The most favorable methods for treating tissue were: 1) immersion in formalin-ethanol-acetic acid followed by paraffin embedding, or 2) light fixation with paraformaldehyde and preparation of vibratome sections. Carnoy's solution, followed by paraffin embedding, extracted myelin from the tissue, while aqueous aldehydes, when used before paraffin embedding, reduced staining everywhere except at sites of compact myelin. The latter conclusion was based, in part, on the almost complete loss of this antigen from the shiverer cord, where compact myelin is known to be virtually absent but where membrane-bound carbonic anhydrase was demonstrated enzymatically. When the optimal methods were used with normal mouse cords, carbonic anhydrase was found throughout the white matter columns and in the oligodendrocytes in gray and white matter. The staining of the white matter was attributed to myelinated fibers because of the similarity in distribution to both a histological myelin stain and the immunocytochemical staining for myelin basic protein. In the mutant mice the oligodendrocyte cell bodies and processes, which were stained in all areas of the spinal cord, were particularly numerous at the periphery of the sections. In contrast to the oligodendrocytes, the fibrous astrocytes appeared to lack carbonic anhydrase, or to have lower than detectable levels, since the astrocyte marker, glial fibrillary acidic protein, had a very different distribution from that of carbonic anhydrase. Even finer localization was obtained in vibratome sections, where the antibody against carbonic anhydrase permitted visualization of the processes connecting oligodendrocytes to myelinated fibers in the normal adult spinal cord.     

Luxol Fast Blue Arn: A New Solvent Azo Dye with Improved Staining Qualities for Myelin and Phospholipids

Luxol fast blue ARN (Du Pont, C.I. solvent blue 37) is a diarylguanidine salt of a sulfonated azo dye. This dye was compared with other Luxol blue and Luxol black dyes. Luxol fast blue ARN has improved staining qualities for phospholipids and myelin, and can advantageously be substituted for Luxol fast blue MBS (MBSN). Appropriate staining times for a 0.1% dye solution in 95% ethanol (containing 0.02% acetic add) at 35°-40° C range from 2-3 hr. After staining, the sections should be rinsed in 95% ethanol, rinsed in distilled water, and differentiated for 2 sec in 0.005% Li₂CO₃, rinsed in 70% ethanol, washed in water, and counterstained as required. Phospholipids and myelin selectively stain deep blue. A fixative containing CaCl₂, 1%; cetyltrimethylammonium bromide, 0.5%; and formaldehyde, 10%, in water gave excellent results with brain. However, 10% formalin can be used. The staining of the phospholipids is probably due to the formation of dye-phospholipid complexes.     

Gomori's Hematoxylin as a cHromosome Stain

The chromic hematoxylin of Gomori (1941) can be used as an excellent chromosome stain after hydrolysis of the tissue in warm 1-N hydrochloric acid. The hydrolysis must be accurately timed for different material as in the case of the Feulgen reaction. The staining of sections can be performed at room temperature and requires about 15 minutes. For pieces of tissue and whole preparations, it is recommended to stain at 60°C. for 40 minutes. Sections stained at room temperature can be differentiated in 1% hydrochloric acid alcohol for one minute and can be counterstained with phloxine according to Gomori's formula. Whole preparations or sections stained at 60°C. must be differentiated in 45% acetic acid for half an hour or more. Tissue pieces may, after staining, be squashed and examined in the acetic acid, but the preparations can also be made permanent. The blue-black stain is very selective and has the advantage of giving high contrast, and it is nonfading, and insoluble in water and other common reagents. It proved definitely superior to other chromosome stains for difficult material such as planarians, rabbit blastocysts, and cleavage stages of sea urchins. Though both the procedure and the result of this method show some similarity to the Feulgen reaction nothing can be said with certainty about its chemical basis.     

Use of eriochrome cyanine R in routine histology and histopathology: is it time to say goodbye to hematoxylin?

Abstract

Eriochrome cyanine R (ECR) is a synthetic anionic dye that forms complexes with cations such as iron. We found that an iron-ECR (Fe-ECR) mixture provided either nuclear or myelin staining depending on the differentiator used. Selective nuclear staining was obtained by differentiation in an aqueous HCl solution, pH 0.95, followed by a wash in slightly alkaline tap water; the pH difference facilitated control of differentiation. When used with an eosin B counterstain, results were nearly indistinguishable from standard hematoxylin and eosin (H & E) staining. Nuclear staining with Fe-ECR provides tinctorial features similar to regressive aluminum-hemateins as well as resistance to acidic solutions such as those of iron hemateins. Fe-ECR also stained selectively intestinal cells of the diffuse neuroendocrine system (DNES). In addition to its use as an H & E substitute, acid differentiated Fe-ECR produced acid-resistant and selective nuclear counterstaining in combination with Alcian blue, and in the Papanicolaou and van Gieson techniques. With alkali differentiation, Fe-ECR produced selective myelin staining, which was compatible with neutral red counterstaining. Myelin sheaths were stained aqua blue. Fe-ECR could be used for both cytological and histological samples, and was suitable for use in automated tissue stainers. ECR also is less expensive than hematoxylin. Hematoxylin still may be preferred as a nuclear counterstain for some immunostaining methods for which Fe-ECR mixtures probably are too acidic.     

An autometallographic technique for myelin staining in formaldehyde-fixed tissue

A new autometallographic (AMG) technique for staining myelin in formaldehyde- or paraformaldehyde- (PFA) fixed tissue is presented. The tissue sections were exposed to AMG development without prior treatment with silver salts. The method was examined on PFA-fixed tissue from mouse, rat, pig, and formaldehyde-fixed human autopsy material. Samples from brain, spinal cord, cranial, and spinal nerves were either cut on a vibratome, frozen and cryostat sectioned, or embedded and microtome sectioned, before AMG development and counterstaining. The AMG-myelin technique results in a specific black/dark-brown staining of myelin in all parts of the CNS and PNS. It works on all species examined, independent of the histological preparation techniques applied. The AMG staining is stable, stays unchanged through decades, allows counterstaining, and has previously been used with immunohistochemical techniques. On perfusion-fixed tissue the technique works without further fixation, but the intensity of the AMG-myelin staining is increased by increased postfixation time. Additionally, immersion fixation has to last for days depending on the size of the tissue block in order to obtain proper myelin staining. The most feasible explanation of the chemical events underlying the AMG-myelin technique is that nano-sized clusters of metallic silver are formed in the myelin as a result of chemical bounds with reducing capacity, exposed or created by the formaldehyde molecule. The AMG method is simple to perform and as specific as the conventional osmium and luxol fast blue stainings. The present technique is thus an effective, simple, inexpensive, and quick myelin staining method of formaldehyde- or PFA-fixed tissue.     

一种简便经济的凝胶电泳同工酶特异染色方法

介绍一种简单、经济的同工酶染色方法：用熔化的0．4％琼脂糖处理滤纸备用,染色前将滤纸浸于同工酶染色液中,染色时将滤纸盖在聚丙烯酸胺胶上,然后将胶放在有盖塑料盒中保温染色,染色时间要比普通方法略长。染色后将胶和滤纸移入固定液中用镊子除去滤纸．     

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.     

The Use of Buffered Solutions in Staining: Theory and Practice

The importance of pH in staining tissue is emphasized. The effect of pH upon the selectivity and intensity of staining with iron hematoxylin, malachite green, and eosin Y is considered. Many difficulties may be avoided by staining in the higher alcohols and directions are given for the preparation of buffer solutions from pH 1.2-8 in alcohol. The concentration of stains, time of staining, and order of staining are discussed for progressive and regressive staining. At pH 8 in 95% alcohol very few tissues stain with malachite green at a concentration of 1/1000 saturated. At pH 6 most cytoplasmic elements stain with malachite green at a concentration of 1/1000 saturated or with eosin Y at 1/250 saturated. As the pH is lowered more tissue elements stain until the nucleus is completely stained. This behavior is in accord with the theory of chemical combination of dyes with proteins, which states that proteins combine with basic dyes on the basic side of their isoelectric points and with acid dyes on the acid side of their isoelectric points. With hematoxylin stain the pH range is much shorter. A satisfactory hematoxylin stain is composed of 0.1% hematoxylin, 0.1% FeCl₃, and HCl to bring the pH to 1.2-1.6 in 80% alcohol. With this stain, which may be used immediately, the nuclei of most tissues begin to stain at pH 1.2 and much of the cytoplasm will be stained if the pH is raised to 1.4. The shortness of this effective pH range is thought to be due to the dissociation of the hematoxylin-iron-protein complex. The use of different dyes successively at different pH values, such as hematoxylin at 1.3, malachite green at 8, and eosin at 6, permits better differentiation of the tissue elements, and intelligent variations in the staining technic.     

The Modified Steiner Stain: a New Use for an Old Stain? Staining Cytomegalovirus-infected Cells in Gastrointestinal Biopsies

The modified Steiner stain is a non-specific silver stain for identifying bacteria in formalin-fixed, paraffin-embedded tissues. The principle behind its use is that bacteria are first sensitized using uranyl nitrate solution, making them able to precipitate silver from a silver nitrate solution. It is used routinely for staining gastric biopsies to identify Helicobacter pylori. Upon staining a gastric biopsy from a patient with acquired immunodeficiency syndrome (AIDS) and cytomegalovirus gastritis, we recognized that this technique also stains the viral inclusions of cytomegalovirus-infected cells. We then proceeded to stain 43 consecutive cytomegalovirus-positive gastrointestinal biopsies from 33 immunocompromised patients based on positive cytomegalovirus immunohistochemistry (DAKO-cytomegalovirus monoclonal antibody, clones DDG9 and CCH2). We also stained eight cytomegalovirus-infected, non-gastrointestinal tissue s, including lung, adrenal gland, ovary, skin and neural tissue, to ensure that the stain was staining the cytomegalovirus-infected cells and not argyrophilic or argentaffin neuroendocrine cells of the gastrointestinal tract. In 40 of the 43 cytomegalovirus-infected gastrointestinal biopsies, we saw positive staining with the modified Steiner stain (93% sensitivity). The cytomegalovirus-infected, non-gastrointestinal tissues all stained positively with the modified Steiner stain. Because the modified Steiner stain is frequently used to identify Helicobacter pylori in gastric biopsies, we propose that it be studied further for possible use either as a screen or as a confirmatory tool, or both, for cytomegalovirus inclusions in gastrointestinal biopsies.