This reference was compiled from the information of several texts. It borrows heavily from L. L. Vacca, Laboratory Manual of Histochemistry, (New York: Raven Press), 1985, a useful but not particularly well put-together book.
10% Neutral-Buffered FormalinTo prepare a 1.0 liter solution:
Citrate SolutionUsed or recommended for antigen unmasking.
- Weigh out 9.60 g of citric acid (or 10.51 g citric acid monohydrate, if using that) into a beaker between 250-600 ml capacity. The beaker should contain an appropriately sized (teflon-coated) stir bar, and rest on a magnetic stirrer. A pH meter, the combination electrode appropriately calibrated, should be nearby.
- Add type I (or highest quality) water to at least 200-250 ml but no more than 400 ml of the mark (depends on beaker size being used).
- Begin stirring until the solid is dissolved, or close to it.
- Place the pH electrode in the solution, and begin reading on the meter.
- Begin dropwise addition of an approximately 1 N NaOH solution until the pH meter reads 6.0.
Tyrode's SalineA commonly used alternative to PBS for histological work.
- 8.00 g/l sodium chloride
- g potassium chloride
- 2.71 g calcium chloride dihydrate
- 0.5 g monosodium phosphate dihydrate
- 2 g magnesium chloride hexahydrate
- 10 g glucose
Tissue Block Processing
Organization and ControlsBoth positive and negative controls should be used whenever possible. This guards against changes in the quality of the reagents, and provides the necessary reference. Controls are recommended for the following staining protocols:
|acid fast (Kinyoun's)||ferric ferrocyanide for ferric ion and hemosiderin (Prussian Blue)||Picro-Sirius acid|
|aldehyde fuchsin||Gridley Fungus Stain||phosphotungstic acid-hematoxylin (PTAH)|
|Alizarin Red S, pH 7 and 9||Levanol Fast Cyanine||Wilder's Reticulum|
|Bielschowsky||MacCallum-Goodpasture||Resorcin Fuchsin-Van Gieson|
|Brown-Benn||Mayer's Mucicarmine||Silver Axon stain|
|Congo Red||Methamine silver||Sirius Supra Scarlet|
|Cresyl Fast Violet||Periodic Acid-Schiff||Tannic Acid-Phosphomolybdic acid-Buffalo Black|
Slide LabelingThe slide label should contain the following information, given in four rows:
- Tissue identification (accession-block number)
- Fixative used
- Staining protocol identifier
- Date and initials of histotechnologist
Slide CleaningDirty slides can be placed in 95-100% alcohol, then wiped with gauze or soft lint-free towel. They can then be air-dried or placed in acetone before drying.
Slide CoatingSeveral methods can be used to make slides more adhesive.
Sectioning of Frozen Tissues
- label the base mold and partially fill mold with frozen tissue matrix
- Place tissues from animal in the pre-labeled base molds, in a way that the tissue is near the bottom and so that tissue is exposed with cut
- Plunge base mold into liquid nitrogen until it almost solidifies (about 30 s). Blocks left too long in nitrogen may crack
- Put block on dry ice (or keep tissue in base mold or transfer to plastic bag), and store at −70° until time for sectioning
- To section, mount block to the cryostat chuck by adhering it with small amount of frozen tissue matrix and allowing to freeze
- Typical depth is 5 μm, with sections collected on to glass slide
- Cover cut surface of block with frozen tissue matrix and freeze and store at −70° if block is to be re-cut later. Do not store tissues on cryostat; many have defrost cycle
- Fix sections in cold (−20°) acetone for 2 min
- Let fixed sections dry completely, label them, and store −70° until they are to be worked up further
FixationPart of the content here from Kiernan JA (1999) Histological and Histochemical Methods: Theory and Practice, 3rd Ed., (London: Arnold).
HeatingThe simplest physical method, but typically used in microbiology.
FreezingFreezing must be done to minimize formation of ice crystals which usually leave cell-size holes. Usually seen in large specimens which are frozen slowly. Freezing must be done rapidly or, if to be done slowly, use of a cryoprotectant is necessary. If the rapid freezing approach is taken, the possibilities include:
- immersion in isopentane cooled to its freezing point (−170° C) using liquid nitrogen
- placing the specimen on a metal block already cooled using liquid nitrogen or dry ice-acetone mixture or even liquid helium (−268° C).
Chemical Fixation/FixativesImportant considerations in the use of chemical fixation will include the nature of the specimen (tissue block) and its ability to be penetrated by the fixative to be used. Some fixatives rapidly penetrate all kinds of tissues and fixation is complete within 24 h even when the specimen is a 5 mm cube. When the fixative slowly penetrates blocks should not be more than 2 mm thick.
Organic Solvents and Acids As FixativesAcetone, ethanol (EtOH), and methanol (MeOH) all work by removing water surrounding protein and thus affect the hydrogen bonding intra- and intermolecularly. This results in denaturation. Soluble proteins coagulate, and organellar structure disrupted. Nucleic acids remain soluble in water (not precipitated). Below −5°, EtOH precipitates but does not denature proteins, and these remain soluble in water. All of these solvents extract lipids from tissues, whereas glycans are unaffected. Typical applications of acetone, EtOH, and MeOH are to fix films (cell smears) and unfixed cryostat sections. They are usually not suitable for tissue blocks because of severe shrinkage and hardening. It might be suitable to use if enzyme activitiy within a section is to be retained. Plant tissues for anatomical study might be fixed in 50-70% EtOH since it preserves cell wall structure.
Formaldehyde-Based FixationFormaldehyde is a gas (bp −21°) which can be passed into solution to a maximal 40% by weight. Concentrates from commercial suppliers are usually 37-40% (w/v). A 10-15% methanol stabilizer is present in reagent grade formalin to prevent polymerization. Concentrations are expressed as the dissolved gaseous material as percent by weight: thus 4% formaldehyde is a better reference than 10% formalin.
Other Aldehydes & GlutaraldehydeGlutaraldehyde is a difunctional aldehyde with a linking three methylene groups. It polymerizes by the aldol condensation reaction to form olefinic molecules, which occurs with time and in more alkaline solutions. Usually it is prepared as an acidic (pH 3) 25% solution. If polymerization is excessive, it is no longer usable as a fixative.
Karnovsky's FixativeThis fixative is used when ultrastructural (EM) work is required for cultured cells or even tissues.
- 2.5 vol 16% paraformaldehyde
- 1 vol EM-grade 50% glutaraldehyde
- 3.125 vol 200 mM sodium phosphate pH 7.2
- 3.125 vol distilled water
- 2.6 vol 16% paraformaldehyde
- 1 vol EM-grade 50% glutaraldehyde
- 10 vol 200 mM sodium phosphate pH 7.2
- 6.4 vol distilled water
Acetic AcidAcetic acid does not fix proteins but coagulates nucleic acids. This is believed to result from an action of the undissociated acid and not the acetate ion, and other carboxylic acids both miscible in water and in lipids are thought to have the same effect. When included in fixatives the intent is to preserve chromosomes, precipitate chromatin in interphase nuclei, and to counter shrinkage caused by agents like EtOH and picric acid.
Other FixativesThe table shows preparation of various fixatives. Percentages are on a volume/volume basis:
Bouin's Fixative (Picro-formol)
75% saturated aqueous picric acid
60% absolute ethanol
5% glacial acetic acid
10% glacial acetic acid
Duboscq-Brasil (alcoholic Bouin's)
60% absolute ethanol
10% glacial acetic acid
10% formaldehyde (37%)
Zenker (Zenker acetic)
Zenker-Formol (Helly's Fluid)
25 g mercuric chloride
Prepare the Zenker fixative, but omit acetic acid; just before use, add 5 ml formalin for each 100 ml
12.5 g potassium dichromate
5 g sodium sulfate
500 ml distilled water
25 ml glacial acetic acid
(best to add this just prior to use, since it keeps badly otherwise)
Clarke's FluidMade with 3 volumes of absolute EtOH and 1 volume of glacial acetic acid just before using, it is a general fixative and also used to preserve nucleic acid and carbohydrates. After a month, ethyl acetate may become significant but not necessarily detract from fixation. After fix, move to 95-100% EtOH.
Carnoy's FluidMade just before use according to recipe above, it rapidly penetrates and coagulates protein and nucleic acids, extracts lipids. Tissue blocks up to 5 mm thick are fixed in 6-8 h then moved to 95 or 100% EtOH. Hydrolysis of nucleic acids (loss of RNA) can occur if left too long in this fix (18 h); lower (5%) acetic acid can be used instead, called modified Carnoy's.
Picric Acid-Based Fixatives, Bouin'sPicric acid (2,4,6-trinitrophenol) is a strong acid fixative which should be present in water. Dry substance is explosive. Nearly saturated solutions have a pH 1.5-2.0 will precipitate proteins by complexing with basic groups. This does not happen in neutral solutions of the substance, and neutrlization also allows resolubilization of proteins. After fixing in the acid, tissues are transferred to 70% EtOH to coagulate the precipitated protein.
Chromium Compounds: Dichromate FixativesChromium trioxide (CrO3) and potassium dichromate (K2Cr2O7) are the substances of interest. Dissolved in water CrO3 will undergo the following reactions:
2 HCrO4− ⇔ Cr2O72− + H2O
Mercuric SaltsHg2+ typically complexes electrostatically with carboxyl groups on proteins. It will also coordinate with hydroxyl groups, and with phosphate in nucleoproteins. Chromium has the ability to bind proteins together however, causing them to coagulate; mercuric ion does not.
Mercuric ChlorideThe molecule Cl-Hg-Cl in solution contains hardly in free Hg2+ ions. A hydro complex is thought to occur: HO-Hg-Cl + H++ Cl−. When acid is added, this will lead to the reverse reaction forming H2O and HgCl2.
HgCl2 + R-NH3+ ⇔ R-N+H2-Hg-Cl + H+ + Cl−
Osmium Tetroxide (OsO4)Osmium tetroxide is volatile at room temperature. The vapor can cloud corneas and is irritating. It is soluble in water (does not ionize or complex) but more so in organic solvents. Traces of organic matter can react to produce OsO2 • 2 H2O. It is a vigorous oxidizer and for this reason is not toxic since it produces a non-toxic product. It should not be discarded down the sink however: it is more economical to recover the expensive osmium for recycling to the tetroxide.
Zenker'sZenker's base is sufficient as a fixative, but metallic salts penetrate tissues slowly. The use of formaldehyde or acetic acid helps to get better penetration of the metal salts.
Non-Aqueous FixativesAlcohol preserves certain specimens well, such as smears, glycogen, pigments, and films of blood/tissue. A brief denaturation in 60-80% at −20° to 4° can keep proteins and enzymes in an undenatured state (relatively). Alcohol hardens and dehydrates simultaneously. It will not leach many useful components that would be soluble in aqueous fixes. It will not fix chromatin, and nucleic acids converted to soluble forms lost in solution. Negatives include that it can overharden and shrink tissue; it can so rapidly harden that cells deep within tissues can become encased, making penetration of substances difficult. It also coagulates cytoplasm.
DehydrationTissues should be dehydrated prior to any paraffin embedding, or else the wax will not infiltrate the tissues. Alkyl alcohols provide the best results. Ethanol is the most commonly used, being relatively non-toxic. Methanol is used for blood/tissue smears and for tissues fixed in methanol-Carnoy (Methacarn). Butanol is sometimes used. While not an alcohol, acetone also provides dehydration and is rapid.
ClearingAlkyl aromatics provide the best result. Xylene is the clearing agent of choice (there are ortho-, meta-, and para isomers). Benzene is sometimes used because it hardens tissue less than does xylene, but it evaporates quickly and is a carcinogen. Toluene evaporates slightly less rapidly than benzene, and is less hardening than xylene, and is sometimes used as well. Methyl benzoate is used when tissues have been fixed in Methacarn.
Infiltration/EmbeddingThe table below shows the different types of paraffin and their melting points. The most commonly used is the 56-58° paraffin.
|soft paraffin||50-52°||thick sections — cut cold|
|hard paraffin||56-58°||thin sections — cut hot|
Embedding ProtocolSeveral days are required to prepared fixed tissues for paraffin sectioning. Most of this time is involved in dehydrating the tissues in graded steps.
- Day 1: incubate tissue block in 4% paraformaldehyde at 4°
- Day 2: incubate in PBS at 4°
- Day 3: incubate in 30% ethanol
- Day 4: incubate in 50% ethanol
- Day 5: incubate in 70% ethanol
- Place in 95% ethanol for at least 30-60 min
- Repeat 95% ethanol in second (fresh) batch for 60 min
- Place in absolute ethanol for 30-60 min
- Repeat absolute ethanol for 60 min
- Incubate in Hemo-De (Fisherbrand xylene) for 30-60 min
- Repeat in Hemo-De for 60 min
- Place tissue in Paraffin #1 at 60° for 60 min. Stir occasionally.
- Place tissue in Paraffin #2 at 60° for 60 min. Stir occasionally.
- Place tissue in Paraffin #3 at 60° for 60 min. Stir occasionally.
Standard Fast ProcedureThe times given below are for tissue specimens not greater than 3 mm thick. Use the slower method below if cutting sections presents some difficulties. No viscous clearing agents are used as in the standard slow method.
|70% dehydrating agent||30 min (longer if convenient)|
|100% dehydrating agent (container 1)||30 min|
|100% dehydrating agent (container 2)||30 min|
|100% dehydrating agent (container 3)||30 min|
|Xylene or toluene (container 1)||15 min|
|Xylene or toluene (container 2)||15 min|
|Wax (container 1)||30 min|
|Wax (container 2)||30 min|
|Wax (container 3)||30 min|
Standard Slow ProcedureThe time periods for each step given below are typical for tissue blocks with a thickness of 3-5 mm. Longer times for all steps should be done for thicker specimens.
|70% dehydrating agent||2 h or overnight|
|95% dehydrating agent||2 h|
|100% dehydrating agent (container 1)||2 h|
|100% dehydrating agent (container 2)||1 h|
|Oily clearing agent (container 1)||2-24 h|
|Oily clearing agent (container 2)||2-24 h|
|Benzene or toluene||15-30 min|
|Wax (container 1)||1 h|
|Wax (container 2)||1 h|
|Wax (container 3)||1 h|
|Wax (container 4)||1 h|
- De-paraffinize if necessary using xylol for 5 min. Repeat two times.
- Rinse in absolute alcohol with three changes of alcohol
- Rinse in 95% alcohol with 2-3 changes
- Rinse in 80% alcohol once only
- Remove the pigment according to the type of fixation
Removing PigmentFor formalin-fixed tissues, perform the rehydration to the 95% alcohol step, then place in saturated picric acid/alcohol solution for 5 min, then rinse in tap water until all acid is removed and the tissue clear (ca. 10 min).
Mayer's Acid Hemalum-Eosin (H & E)
Harris' Acid Hemalum-Eosin (H & E)
Weigert's Acid Iron Chloride Hemalum
Modified Heidenhain's Iron Hematein
Chrome Alum Gallocyanin
Celestin Blue B
Iron Alizarin Blue S
Nuclear Fast Red (Kernechtrot)
Modified Cresyl Fast Violet
Best's Carmine Method
Alcian Blue, pH 2.5, 1.0, 0.4
Toluidine Blue, pH 0.5 or 4-4.5
Hale's Colloidal Ferric Oxide
Oil Red O
Sudan Black B
Lillie's Sulfuric Nile Blue Technic
Baker's Acid Hematein
Phosphotungstic acid Hemaotyxlin (PTAH)
Luxol Fast Blue-Periodic Acid Schiff-Toludine
Luxol Fast Blue-Levafix Red Violet (LFB-Levafix)
Tannic Acid-Phosphomolybdic Acid-Levanol Fast Cyanine 5 RN (TPL)
Tannic Acid-Phosphomolybdic Acid-Thiazine Red R
Fibrin, Muscle Striations
Phosphotungstic Acid Hematoxylin (PTAH)
Gomori's Green Trichrome
Weigert's Hematoxylin-Picro-Sirius Red (PSR)
Puchtler's Alkaline Congo Red
Highman's Crystal Violet
Sirius Supra Scarlet or Sirius Red
Modified Resorcin Fuchsin-Van Giesen (RFVG)
Modified Verhoeff's Elastica Stain
Puchtler's Blue Trichrome
Masson's Trichome Method
Phosphomolybdic Acid-Aniline Blue
Modified Lillie's PAS Allochrome
PAS Phosphomolybdic Acid (PMA)-Sirius Supra Blue FGL-CF
Periodic Acid-Sodium Bisulfite-Resorcin Fuchsin (PBRF)
Resorcin Fuchsin-Tannic Acid-Phosphomolybdic Acid (RFTP)-Orseillin
Alkaline, Neutral, and Acid Alizarin Red S
Von Kossa's Technique
Cresyl Fast Violet
Gridley's Stain, Bauer Variant
Kinyoun's Carbol Fuchsin (Acid Fast Technique)
Brown-Brenn Bacterial Stain
Grocott's Methenamine Silver
Fontanna-Masson Silver Method
Silver Axon Stain
Holmes Silver Stain-Luxol Fast Blue
Gold Sublimate Stain
Silver Carbonate Method
Demonstration of Metals
Gomori's Prussian Blue Reaction
Lillie's Ferric Ferrocyanide and Ferric Ferricyanide Tests
Tissue Affinity for Iron
Uzman's Rubeanic Acid Method
Polarization of Uric Acid Crystals
Gomori's Methenamine Silver
De Galantha's Method
Gomori's Aldehyde Fuchsin
Acid Fuchsin-Aniline Blue Method
Modified Rosindole Reaction/Dimethylaminobenzaldehyde
Dihydroxy-Dinaphthyl-Disulfide Method (DDD)
Adenosine Triphosphatase (ATPase)
DOPA Oxidase (tyrosinase/polyphenol oxidase)
Peroxidase (Lillie-Washburn Technique)
Trypsin, Neuraminidase, Collagenase
Dyes/Stains[many notes in this section from Kiernan JA, Histological & Histochemical Methods: Theory & Practice, 3rd Ed.]
Nomenclature & ClassificationBecause the chemical names of many dyes are too long, trivial or informal names are given to them.
- Basic dyes. Used a lot on proteinaceous and cellulosic fibers mordanted with tannic acid.
- Acid dyes. Have sulfonic and carboyxlic auxochromes, although many are amphoteric (have amine groups). In textiles used to stain proteinaceous fibers (silk, wool). Usually stain cytoplasm and extracellular structures.
- Direct dyes. Nearly all are azo dyes, being anionic with large molecules. Bind to cellulose directly without mordants.
- Mordant dyes. By definition metal ions are used in the staining process. The mordant may be applied prior to the chromophore, with it as a complex (metachrome), or after (after-chrome).
- Reactive dyes. Combine covalently with the substrate.
- Solvent dyes. Chromophore that are not water soluble but hydrophobic and embed in hydrophobic layers. Also calledlysochromes.
- Vat dyes. Important only in textile industry. These dyes are applied in colorless (
leuco) form and then an oxidizing treatment (air, steam) is done to produce color and insolubility (fixation). The classic application is indigo staining of cotton. Has no basis in histochemistry.
- Sulfur dyes. A process involving heating various organics with sulfur or alkali metal polysulfides. This usually causes reduction of a leuco compound to a color form. No purpose in histology.
Dyes Classed By Chromophoric System
Nitroso DyesNitrous acid (HNO2) reacts with phenols to produce nitrosophenols, creating an aromatic substitution ortho and para to the phenol group. The ortho product can form colored chelates with metal ions and be mordant dyes. They are not really used in histology but have properties that might be exploited.
Nitro DyesThese are dyes with an -NO2 group. Picric acid is a yellow anionic dye, forming additional compounds as charge-transfer complexes, with bonding neither ionic or covalent. It is also a fixative, soluble in water (to 1.3%) and more so in alcohol and aromatric hydrocarbons. Xylene does not clear it.
Azo DyesThis is a huge class of dyes useful in industry and biology. The process starts by the oxidation of anilines (primary aromatic amines) with nitrous acid in an acidic solution at near-freezing temperatures:
butter yellow, insoluble in water and not a histological dye. Adding a sulfonic acid group para to the aniline converted to the diazonium makes it water soluble: that product is methyl orange (CI 13025, Acid orange 52), which is an acid-base indicator and not a dye.
leveling dyes.They have sulfonic and phenolic substituents, such asorange G. This is a cytoplasmic stain used in conjunction usually with anionic dyes. Very soluble in water and less so in EtOH. The anhydrous dye should at least be 80% by weight in quality.
Milling dyesare larger molecules with bis- and trisazo functions. These anionic dyes are useful for connective tissue.Biebrich scarlet is soluble in water, only slightly in alcohol, and used as a counterstain for collagen. It does not change color over a wide pH range. Alkaline solutions containing it might be used for staining strongly basic proteins.
Stains-Alland used to stain sections various hues of blue, purple and red. However, the stain fades rapidly so it is of limited utility.
Basic red 9
Basic violet 14
Basic violet 2, Magenta III
Basic violet 3
Basic blue 20
458 (as dichloride)
Acid violet 19
Patent Blue VF
Acid blue I
All of these dyes are cationic or anionic as a net charge, with either chloride or sodium counterions.
methyl greenare always ethyl green. The methyl green dye is used in methyl green-pyronine for nucle acid stain. Older samples of methyl green are likely to contain crystal violet because one of the alkyl groups detaches easily on the quaternary nitrogen.
aniline blue water-soluble.Insoluble in alcohol and soluble in water. Change to a red color in strong alkaline solution and used as anionic dyes of large molecular size, especially for connective tissue.
|Substituent on fluorescein skeleton at carbon number|
(carbon numbering is after Lillie, 1977)
|Eosin (eosin Y, eosin yellowish)|
Acid red 87
|Eosin B (bluish)|
Acid red 91
Acid red 98
Acid red 92
Acid red 95
Acid red 51
Acid red 94
- Rinse any material away that can react with the fixative; it is best to use saline, perhaps one that is phosphate-buffered
- Fix in zinc formalin for the appropriate period (3 min)
- Wash in tap water well, then rinse in distilled water (total 1 min)
- Stain with hematoxylin dye (2-3 min)
- Wash in tap water, rinse in distilled water (total 1 min)
- A clarifying agent can now be used but is optional (30 sec)
wash in tap and rinse in distilled, if used
- Dip 6 times in eosin
- Dip 8 times in one container of absolute ethanol
- Dip another 8 times in another container of absolute ethanol
- Dip in primary container of xylene until the sheets slide off evenly
- Dip 10 times (or until saturated) in a secondary xylene container
- Keep held in a third container of xylene until ready to apply coverslip
- Progressive staining involves adding the right amount of acid (if any) and alum to begin with, and staining for the proper period
- Differentiation selectively washes (de-stains) an overstained specimen, but even with the same technician and technique, the same specimen can have a different appearance. This is sometimes an acceptable outcome
- Wash the culture dish/well with PBS three times.
- Fix cells for 10 min in neutral-buffered 10% formalin.
- Wash with distilled water extensively.
- Prepare the stain as 3 parts of 0.5% Oil Red O in isopropanol and 2 parts water filtered through 0.45 μm. Stain the tissue for 10-15 min.
- Plates are washed 3 × in water.
- The percent adipocytes are determined from counting 50-100 cells in multiple fields.
- Cultures are washed in PBS (maybe a couple of times)
- Fix the cells in ice-cold 70% ethanol for 60 min
- Rinse with water
- Stain with 1 ml of 40 mM (2% w/v, according to Phinney et al) Alizarin Red S, adjusted to pH 4.1 with ammonium hydroxide. The 1 ml volume assumes that 12-well culture plates are being used; adjust volume as necessary. Put flasks on rotary (orbital) shaker during the staining procedure
- Rinse 2-3 times with PBS (water, according to Phinney et al) to remove excess or non-specifically bound stain.
- Perform steps 1-3 of the Oil Red O staining procedure. This is all about the necessary typical steps of washing, fixing, and rinsing cells.
- Stain the culture for 30 min in 10% Alcian Blue in 0.1 N HCl
- Wash 3 × TBS (50 mM TrisHCl, pH 7.4 + 0.15 M NaCl)
At this point, it is possible to store cells at −20° until the activity quantitation is to be done
- Scrape the cell layer of the well with 0.5 ml of 50 mM TrisHCl, pH 7.4 present
- Sonicate the scrapings (a Fisher Sonic Dismembranator Model 150 set at 30-40% max power is useful)
- Determine activity using 3 mM p-nitrophenylphosphate in 0.7 M 2-amino-2-methyl-1-propanol, pH 10.3 + 6.7 mM MgCl2.
- Prepare a Citrate Working Solution by diluting 2 ml of Citrate Concentrate to 100 ml with deionized water. This should be stored at 2-8° if it is not entirely consumed.
- Fixative solution is prepared as 60% citrate-buffered acetone by adding 2 volumes of warm (18-26°) Citrate Working Solution to 3 volumes of acetone, with constant stirring. This must be discarded after use.
- 48 ml water should be at room temperature (18-26°). Dissolve the diazonium salt (Fast Blue RR or Fast B Violet) in the water; stirring may help.
- Add 2 ml Naphthol-AS-MX Phosphate Alkaline Solution the diluted diazonium salt solution.
- The fixative should be at room temperature. Add the citrate-buffered acetone for 30 seconds. Rinse gently in deonized water for 45 seconds. Don't allow the plates to dry.
- Cells should be washed 2-3 × in Tyrode's solution
- Cell layers are fixed in 10% formalin for 1 h (others have used 1% glutaraldehyde for 15 min)
- Rinse 2-3 × in distilled water
- Treat with 2% silver nitrate for 10 min in the dark
- Wash well with deionized water
- Expose to a bright light for 15 min while immersed in water
- Rinse again in water, then dehydrate with 100% ethanol (progressive dehydration is a possibility)
- pH 0.5: dissolve 0.5 g TBO in 100.0 ml of 0.5 N HCl
- pH 4-4.5: dissolve 0.5 g TBO in 100 ml of 10 mM sodium acetate, pH 4-4.5
Counterstaining with TBO To Minimize Quenching of Fluorescent DyesD. K. Chelvanayagam and L. B. Beazley (J. Neurosci. Methods 72, 49-55, 1997) have reported that TBO can be used carefully with certain fluorescent dyes and not quench the fluorescence of those dyes.
Iodine SolutionsGram's Iodine is 1 g iodine and 2 g potassium iodide dissolved in 300 ml quality water (another source says it is 1% iodine in 2% potassium iodide).
Saturated Picric AcidTake approximately 90 g picric acid crystals and add it to 4 L of 95% ethanol and mix well.
This information was taken from IHC World Online Information Center for Immunohistochemistry and is reproduced here.
- Weigert’s Iron Hematoxylin. Mix in equal volumes a 1% hematoxylin in 95% ethanol solution with acidic ferric chloride (to 4 ml of a 29% ferric chloride, add 95 ml distilled water, then 1 ml concentrated hydrochloric acid). This solution is stable for a month
- 10 ppm Fast Green (FCF). Prepare 100 mg Fast Green (FCF, C.I. 42053) in 1 liter of distilled water.
- 1% acetic acid
- 0.1% Safranin O. Add 100 mg Safranin O (C.I. 50240) to 100 ml distilled wtter.
- If working with paraffin-embedded tissues, de-paraffinize and rehydrate using standard procedures. Make sure the last rehydration step is in pure water.
- Stain 10 min with Weigert's iron hematoxylin.
- For 10 min, wash in running tap water.
- Stain 5 min with the Fast Green (FCF) solution.
- Quickly rinse for no more than 10-15 seconds the slides in 1% acetic acid.
- Let sit in the Safranin O solution for 5 min.
- Dehydrate with 95% EtOH, then absolute EtOH. Clear with xylene. In all of these steps, do 2 changes each, and let sit 2 min each.
- Use a resin for mounting and then observe.
- Kahveci Z, Minbay FZ, Cavusoglu L (2000) Safranin O staining using a microwave oven. Biotech Histochem. 75:264-268.
- Tran D, Golick M, Rabinovitz H, Rivlin D, Elgart G, Nordlow B (2000) Hematoxylin and safranin O staining of frozen sections. Dermatol Surg. 26:197-199.
- Camplejohn KL, Allard SA (1988).Limitations of safranin 'O' staining in proteoglycan-depleted cartilage demonstrated with monoclonal antibodies. Histochemistry 89:185-8.
Perl's Reagent (for Prussian Blue Staining)This is used for iron (specifically ferric, Fe3+)-containing tissues or cells. Ferrous (Fe2+) will produce no colored product.
- 4% (v/v) HCl stock.
- Potassium ferrocyanide: 2% (w/v) stock.
- Working acidified potassium ferrocyanide: equal volumes of 4% HCl and 2% potassium ferrocyanide, which should be each half the volume of the total volume required. Prepare just before use.
- Neutral red. 1% dye in 1% (v/v) acetic acid (add acid after dissolving first in water).
- After fixing (buffered formalin good), rinse well in distilled water
- Flood with equal parts of ferrocyanide and HCl and leave for 10 min.
- Wash several times with distilled water over a 5 min period
- Counterstain with filtered neutral red for 1 min
- Rinse in water. Dip in absolute EtOH to rapidly dehydrate, let clear, then mount.
Diaminobenzidine (DAB)DAB (diaminobenzide: (3,3'-diaminobenzidine 4HCl) will oxidize in presence of H2O2, although slowly (more quickly with peroxidase). The presence of nickel or cobalt salts can produce a greater darkness (blue-black instead of brown) polymers; nickel is more popular.
Use of DAB in Photoconversion of Fluorescent Dyes/StainsWhen a fluorophore is unstable or will be lost over time, it is useful to exploit the finding that fluorophores can be used in the photo-oxidation of DAB into an insolube form, replacing the fading fluorophore with insoluble DAB product.
- Rinse the affected area in 0.1 M Tris pH 8.2 to start, and blot away excess.
- Place one drop containing 1.5 mg DAB/ml (0.15%) in Tris buffer.
- Use the filter set that achieves the brightest fluorescence and position field on the slide that is to get the photoconversion. A lower objective will give a wider field, so consider this.
- Replace with fresh DAB (on ice) every 10 min. Reaction usually complete in 20-40 min, but monitor the reaction under visible light to make sure the photo conversion is occurring.
- Rinse in Tris buffer at end, then continue processing with other stains or mounting.
Fluorophores and Their Chemistry, Fluorescent Dyes and Stains
Prolong Gold with DAPI & antifadeProlongGold with DAPI + antifade (Invitrogen P36931) is stable for 6 months at −20°.
- Warm to room temperature before using.
- Remove "excess moisture" from slide
- Apply to specimen on slide, or if specimen on coverslip, apply to slide and then drop coverslip (cell side down) on to slide.
- Cure the specimen: place on flat dry surface and leave 24 h at RT in dark
- Coverslip edges should be sealed with epoxy or nail polish and sample stored at 4 or -20 deg for extended length storage
Crystal MountCrystal Mount is originally a product of Biomeda, Inc. (company no longer exists) now sold by Sigma (Cat #C0612). It is a water soluble material that when it dries protects the specimen from dust. 30 ml will be useful for some 50 slides.
- To cover the area about the size of a quarter, apply 3 drops of Crystal Mount.
- Rotate the slide/surface angled so that the precession spreads the solution about the size of a quarter.
- Drying/curing is done in 1-2 h at 20-37°, or it can be done in 30 min in a 40-50° oven. For fluorophores a 60-70° oven can be used for 20 min.
Best Wishes: Dr.Ehab Aboueladab, Tel:01007834123 Email:email@example.com,firstname.lastname@example.org