Fixation of Tissues

1. Where the best possible morphology is required, animals should be anesthesized and subjected to cardiac perfusion with saline, followed by a 10% formalin flush. If biochemical studies need to be performed on the tissue, a 10% formalin flush should not be used as it may interfere with subsequent analysis.
2. For routine stains where perfusion is not required, tissue is sectioned and  drop-fixed in a 10% formalin solution.  Fixative volume should be 20 times that of tissue on a weight per volume; use 2 ml of formalin per 100 mg of tissue.
3. Due to the slow rate of diffusion of formalin (0.5 mm hr),  tissue should  be sectioned into 3 mm slices on cooled brain  before transfer into formalin. This will ensure the best possible preservation of tissue and offers rapid uniform penetration and fixation of tissue within 3 hours.
4. Tissue should be fixed for a minimum 48 hours at room temperature.
5. After 48 hours of fixation, move tissue into 70% ethanol for long term storage.
6. Keep fixation conditions standard for a particular study in order to minimize variability. (Although set times are best, tissue may be fixed for substantially longer periods without apparent harm.

A few notes on fixation

The usual fixative for paraffin embedded tissues is neutral buffered formalin (NBF). This is equivalent to 4% paraformaldehyde in a buffered solution plus a preservative (methanol) which prevents the conversion of formaldehyde to formic acid.  Because of the preservative, NBF has a shelf life of months, whereas 4% PF must be made fresh.  Optimal histology requires adequate fixation, about 48 hrs at room temperature for thinly sliced tissues.  Inadequately fixed tissues will become dehydrated during tissue processing, resulting in hard and brittle specimens.  Alcohol based fixatives generally do not give good morphology but may be useful in special cases (such as BrdU staining).  A particular challenge for the histopathology is immunostaining fixed specimens.  In many cases formaldehyde fixation will prevent recognition of epitopes by the primary antibody.  Occasionally, “antigen retrieval” procedures will improve results but usually frozen sections are a better bet.  An alternative approach, suitable for thin or porous tissues, is to perform immunohistochemistry on fresh tissues and then post-fix and embed the tissues in paraffin.

Decalcification of bone (optional)

After fixation, bone,must be decalcified, or else it won’t cut on the microtome:

• Immerse tissue cassette in 11% formic acid with a stir bar overnight in a fume hood.
• Rinse in running water for 30- 60 minutes (the smell should be gone).

Storage in 70% Ethanol
After adequate fixation tissues are transferred to 70% ethanol and may be stored at 4°C.

Paraffin infiltration

In this procedure, tissue is dehydrated through a series of graded ethanol baths to displace the water, and then infiltrated with wax. The infiltrated tissues are then embedded into wax blocks. Once the tissue is embedded, it is stable for many years.

The most commonly used waxes for infiltration are the commercial paraffin waxes. A paraffin max is usually a mixture of straight chain or n-alkanes with a carbon chain length of between 20 and 40; the wax is a  solid at room temperature but melts at temperatures up to about 65°C or 70°C. Paraffin wax can be purchased with melting points at different temperatures, the most common for histological use being about 56°C–58°C, At its melting point it tends to be slightly viscous, but this decreases as the temperature is increased. The traditional advice with paraffin wax is to use this about  2°C above its melting point. To decrease viscosity and improve infiltration of the tissue,  technologists often increase the temperature to above 60°C or 65°C in practice to decrease viscosity.

In the schedule below, it is presumed that the working day is from 8:00 a.m. to 5:00 p.m. If other than that, appropriate adjustments should be made.

Trim fixed tissues and keep in neutral buffered formalin (NBF) until ready to proceed. Put tissues in a labeled (usually with pencil, as solvents dissolve the ink) cassette.

Once fixed, tissue is processed as follows, using gentle agitation, usually on a tissue processor, as follows:

1. 70% ethanol for 1 hour.
2. 95% ethanol  (95% ethanol/5% methanol)  for 1 hour.
3. First absolute ethanol for 1 hour .
4. Second absolute ethanol 1½ hours .
5. Third absolute ethanol 1½ hours.
6. Fourth absolute ethanol 2 hour.
7. First clearing agent ( Xylene or  substitute) 1 hour.
8. Second First clearing agent (Xylene or  substitute) 1 hour.
9. First wax (Paraplast X-tra) at 58°C  for 1 hour.
10. Second wax (Paraplast X-tra) at 58°C 1 hour.

Due to the viscosity of molten paraffin wax, some form of gentle agitation is highly desirable. If the processor is to be run overnight it should be programmed to hold on the first ethanol bath and not finish until the next morning so the specimens do not sit in hot paraffin longer than the time indicated.  If specimens are fresh they may incubate in formalin in the first stage on the machine.  It is important to not keep the tissues in hot paraffin too long or else they become hard and brittle. Processed tissues can be stored in the cassettes at room temperature indefinitely.

Embedding tissues in paraffin blocks

Tissues processed into paraffin will have wax in the cassettes; in order to create smooth wax blocks, the wax first needs to be melted away  placing the entire cassette in 58°C paraffin bath for 15 minutes.  Turn the heat block on to melt the paraffin one hour before adding the tissue cassettes.

1. Open cassette to view tissue sample and choose a mold that best corresponds to the size of the tissue.  A margin of at least 2 mm of paraffin surrounding all sides of the tissue gives best cutting support.  Discard cassette lid.
2. Put small amount of molten paraffin in mold, dispensing from paraffin reservoir.
3. Using warm forceps, transfer tissue into mold, placing cut side down, as it was placed in the cassette.
4. Transfer mold to cold plate, and gently press tissue flat.  Paraffin will solidify in a thin layer which holds the tissue in position.
5. When the tissue is in the desired orientation add the labeled tissue cassette on top of the mold as a backing.   Press firmly.
6. Hot paraffin is added to the mold from the paraffin dispenser. Be sure there is enough paraffin to cover the face of the plastic cassette.
7. If necessary, fill cassette with paraffin while cooling, keeping the mold full until solid.
8. Paraffin should solidify in 30 minutes.  When the wax is completely cooled and hardened (30 minutes) the paraffin block can be easily popped out of the mold; the wax blocks should not stick. If the wax cracks or the tissues are not aligned well, simply melt them again and start over.

Lung tissue stained with the H&E technique. Nuclei are darkly stained in this image.

H&E stain, HE stain or hematoxylin and eosin stain, is a popular staining method in histology. It is the most widely used stain in medical diagnosis; for example when a pathologist looks at a biopsy of a suspected cancer, the histological section is likely to be stained with H&E and termed H&E section, H+E section, or HE section.

The staining method involves application of hemalum, which is a complex formed from aluminium ions and oxidized hematoxylin. This colors nuclei of cells (and a few other objects, such as keratohyalin granules) blue. Materials colored blue by hemalum are often said to be basophilic, but this is an incorrect use of the word. The nuclear staining is folowed by counterstaining with an aqueous or alcoholic solution of eosin Y, which colors eosinophilic other structures in various shades of red, pink and orange.

Haematoxylin Solutions

Haematoxylin stains are commonly employed for histologic studies, often employed to color the nuclei of cells (and a few other objects, such as keratohyalin granules) blue. The mordants used to demonstrate nuclear and cytoplasmic structures are alum and iron, forming lakes or colored complexes (dye-mordant-tissue complexes), the color of which will depend on the salt used. Aluminium salt lakes are usually colored blue white while ferric salt lakes are colored blue-black.

The three main alum haematoxylin solutions employed are Ehrlich’s haematoxylin, Harris’s haematoxylin and Mayer’s haematoxylin. The name haemalum is preferable to “haematoxylin” for these solutions because haematein, a product of oxidation of haematoxylin, is the compound that combines with aluminium ions to form the active dye-metal complex. Alum haematoxylin solutions impart to the nuclei of cells a light transparent red stain which rapidly turns blue on exposure to any neutral or alkaline liquid.

Alum or potassium aluminium sulfate used as the mordant usually dissociates in an alkaline solution, combining with OH^? of water to form insoluble aluminium hydroxide. In the presence of excess acid, aluminium hydroxide cannot be formed thus failure of aluminium haematoxylin dye-lake to form, due to lack of OH^? ions. Hence, acid solutions of alum haematoxylin become red. During staining alum haematoxylin stained sections are usually passed on to a neutral or alkaline solution (e.g. hard tap water or 1% ammonium hydroxide) in order to neutralize the acid and form an insoluble blue aluminium haematin complex. This procedure is known as blueing.

When tap water is not sufficiently alkaline, or is even acid and is unsatisfactory for blueing haematoxylin, a tap water substitute consisting of 3.5 g NaHCO₃ and 20 g MgSO₄.7H₂O in one liter of water with thymol (to inhibit formation of moulds), is used to accelerate blueing of thin paraffin sections. Addition of a trace of any alkali to tap or distilled water also provides an effective blueing solution; a few drops of strong ammonium hydroxide or of saturated aqueous lithium carbonate, added immediately before use, are sufficient for a 400 ml staining dish full of water. Use of very cold water slows down the blueing process, whereas warming accelerates it. In fact, the use of water below 10°C for blueing sections may even produce pink artifact discolorations in the tissue.

The staining of nuclei by hemalum does not require the presence of DNA and is probably due to binding of the dye-metal complex to arginine-rich basic nucleoproteins such as histones. The mechanism is different from that of nuclear staining by basic (cationic) dyes such as thionine or toluidine blue. Staining by basic dyes is prevented by chemical or enzymatic extraction of nucleic acids. Such extractions do not prevent staining of nuclei by hemalum.

Eosin Solutions

Eosin is a fluorescent red dye resulting from the action of bromine on fluorescein. It can be used to stain cytoplasm, collagen and muscle fibers for examination under the microscope. Structures that stain readily with eosin are termed eosinophilic.Eosin is most often used as a counterstain to haematoxylin in H&E (haematoxylin and eosin) staining.  Eosin stains red blood cells intensely red. Eosin is an acidic dye and shows up in the basic parts of the cell, ie the cytoplasm. For staining, eosin Y is typically used in concentrations of 1 to 5 percent weight by volume, dissolved in water or ethanol. For prevention of mold growth in aqueous solutions, thymol is sometimes added. A small concentration (0.5 percent) of acetic acid usually gives a deeper red stain to the tissue.

Other colors, e.g. yellow and brown, can be present in the sample; they are caused by intrinsic pigments, e.g. melanin.

Some structures do not stain well. Basal laminae need to be stained by PAS stain or some silver stains in order to exhibit appropriate contrast. Reticular fibers also require silver stain. Hydrophobic structures also tend to remain clear; these are usually rich in fats, eg. adipocytes, myelin around neuron axons, and Golgi apparatus membranes.

Protocols

There are a large number of H&E protocols available for the histotechnologist. For most tissues, these approaches can be used interchangably, and selection of a particular protocol will be based upon the particular needs of the investigator. Primary differences are dye composition, staining protocol, and intensity of blue dye. Staining contrast for a particular tissue will differ depending upon the approach that is used.

Mayer’s Hematoxylin Protocol

Solutions

Mayer’s Hematoxylin

1. Dissolve 50 g aluminum potassium sulfate (alum) in 1000 ml distilled water.
2. When alum is completely dissolved, add 1 gm hematoxylin.
3. When hematoxylin is completely dissolved, add 0.2 gm sodium iodate and 20 ml acetic acid.
4. Bring solution to boil and cool, and filter

Staining Method

Staining times will vary based upon depth of stain requiredFor slide-mounted immunohistochemistry, counterstain tissue for 30 seconds. For H&E staining, counterstain tissue for 5 minutes.

In order to blue the stain, put slides through 4 changes of tap water, 5 minutes each.

Results

This recipe should create sharp blue nucleus staining with little background.

Harris’ Hematoxylin and Eosin (H&E) Staining Protocol

Solutions and Reagents

Staining Procedure

1. Deparaffinize sections, 2 changes of xylene, 10 minutes each.
2. Re-hydrate in 2 changes of absolute alcohol, 5 minutes each.
3. 95% alcohol for 2 minutes and 70% alcohol for 2 miuntes.
4. Wash briefly in distilled water.
5. Stain in Harris hematoxylin solution for 8 minutes.
6. Wash in running tap water for 5 minutes.
7. Differentiate in 1% acid alcohol for 30 seconds.
8. Wash running tap water for 1 minute.
9. Bluing in 0.2% ammonia water or saturated lithium carbonate solution for 30 seconds to 1 minute.
10. Wash in running tap water for 5 minutes.
11. Rinse in 95% alcohol, 10 dips.
12. Counterstain in eosin-phloxine solution for 30 seconds to 1 minute.
13. Dehydrate through 95% alcohol, 2 changes of absolute alcohol, 5 minutes each.
14. Clear in 2 changes of xylene, 5 minutes each.
15. Mount with xylene based mounting medium.

Results

Nuclei should be blue, cytoplasm pink to red.

References

Kiernan JA (2008) Histological and Histochemical Methods: Theory and Practice. 4th ed. Bloxham, UK: Scion.

Lillie RD, Pizzolato P, Donaldson PT (1976) Nuclear stains with soluble metachrome mordant lake dyes. The effect of chemical endgroup blocking reactions and the artificial introduction of acid groups into tissues. Histochemistry 49: 23-35.

Llewellyn BD (2009) Nuclear staining with alum-hematoxylin. Biotech. Histochem. 84: 159-177.

Puchtler H, Meloan SN, Waldrop FS (1986) Application of current chemical concepts to metal-haematein and -brazilein stains. Histochemistry 85: 353-364.

Yarn drying after being dyed in the early American tradition, at Conner Prairie living history museum.

A mordant is a substance used to set dyes on fabrics or tissue sections by forming a coordination complex with the dye which then attaches to the fabric or tissue.^[1] It may be used for dyeing fabrics, or for intensifying stains in cell or tissue preparations. A mordant is always a polyvalent metal ion.^[2] The resulting coordination complex of dye and ion is colloidal and can be either acidic or alkaline.

Common dye mordants

Mordants include tannic acid, alum, urine, chrome alum, sodium chloride, and certain salts of aluminium, chromium, copper, iron, iodine, potassium, sodium, and tin.

Iodine is often referred to as a mordant in Gram stains but is in fact a trapping agent. ^[3]

Dyeing methods

The three methods used for mordanting are:

• Pre-mordanting (onchrome): The substrate is treated with the mordant and then dyed.
• Meta-mordanting (metachrome): The mordant is added in the dye bath itself.
• Post-mordanting (afterchrome): The dyed material is treated with a mordant.

The type of mordant used changes the shade obtained after dyeing and also affects the fastness property of the dye. The application of mordant, either pre-, meta- or post-mordant methods, is influenced by:

• The action of the mordant on the substrate: if the mordant and dye methods are harsh (e.g. an acidic mordant with an acidic dye), pre- or post- mordanting limits the potential for damage to the substrate.
• The stability of the mordant and/or dye lake: the formation of a stable dye lake means that the mordant can be added in the dye without risk of losing the dye properties (meta-mordanting).

Dye results can also rely on the mordant chosen as the introduction of the mordant into the dye will have a marked effect on the final colour.

The dye lake

The dye lake is formed when the complex of dye and mordant are combined, which then attaches to the substrate.^[2]

The term “lake” is derived from the term lac, the secretions of the Indian wood insect Laccifer lacca (formerly known as the Coccus lacca.^[4] The type of mordant used can change the colour of both the dye-plus-mordant solution and influence the shade of the final product.

Cotton

Since metallic mordants are soluble in water and are loosely held by the cotton fibres, these mordants have to be precipitated on the fabric by converting them into insoluble form, or by first treating the fibres with oil or tannic acid and then impregnating treated fabric with solution of mordant, whereby the metallic mordants are held on to cotton via oil or tannic acid.

Wool

Unlike cotton, wool is highly receptive toward mordants. Due to its amphoteric nature wool can absorb acids and bases equally effectively. When wool is treated with a metallic salt it hydrolyses the salt into an acidic and basic component. The basic component is absorbed at –COOH group and the acidic component is removed during washing. Wool also has a tendency to absorb fine precipitates from solutions; these cling to the surface of fibres and dye particles attached to these contaminants result in poor rubbing fastness.

Silk

Like wool, silk is also amphoteric and can absorb both acids as well as bases. However, wool has thio groups (-SH) from the cystine amino acid, which act as reducing agent and can reduce hexavalent chromium of potassium dichromate to trivalent form. The trivalent chromium forms the complex with the fibre and dye. Therefore potassium dichromate cannot be used as mordant effectively.

Animal and Plant Tissues

In Histology, mordants are indispensable in adhering dyes to tissues for microscopic examination.

Methods for mordant application depend on the desired stain and tissues under study; pre-, meta- and post-mordanting techniques are used as required.

The most commonly used stain used in diagnostic histology of animal tissues is Harris’ haematoxylin as part of a haematoxylin and eosin (H&E) stain.

References

1. International Union of Pure and Applied Chemistry (1993). “mordant”. Compendium of Chemical Terminology Internet edition.
2. ^{a b} Llewellyn, Bryan D. (May, 2005). “Stain Theory – How stains work”. http://stainsfile.info/stainsfile/theory/mordant.htm/. Retrieved 2009-09-20.
3. Llewellyn, Bryan D. (May, 2005). “Stain Theory – Trapping agents”. http://stainsfile.info/StainsFile/theory/trapping.htm/. Retrieved 2009-09-20.
4. Llewellyn, Bryan D. (May, 2005). “Stain Theory – Lac”. http://stainsfile.info/StainsFile/dyes/75450.htm/. Retrieved 2009-09-20

A semi-permanent mounting media for immunofluorescence microscopy.

Method

1. Add the following reagents to a 250 ml flask or beaker:

• 24 g analytical grade glycerol (Sigma #G-6279)
• 9.6 g Mowiol 4-88 (Fluka, #81381, can be purchased through Sigma-Aldrich)
• 24 ml distilled water
• 48 ml 0.2M Tris buffer, pH 8.5

2. Stir with a clean stir bar on a hot plate on warm (not boiling)at least 4-5 hours until the majority of the Mowiol powder goes into solution.
3. Aliquot into 50 ml centrifuge tubes, weigh and balance
4. Centrifuge at 5000g for 15 minutes. Carefully remove the supernatant without disturbing the pellet at the bottom of the flask.
5. Aliquot into 15 ml conical tubes – add only 10 mls to each tube to allow for expansion with freezing.
6. Aliquots may be stored at -20C for 12 months. Store at room temperature no more than one month.
7. To use, warm solution to room temperature to eliminate bubble formation. use approximately 10 ul of mounting media for an 18 mm coverslip.
8. Allow slides to dry overnight at room temperature in a light-tight box.

Weil’s stain is a modification for paraffin sections of the Weigert-Pal-Kulschitsky technique. The underlying principle of these methods involves the reduction of chrome salt to chromium dioxide by myelin. The chromium subsequently acts as a mordant for the haematoxylin, intensifying the stain.

Procedure

This procedure is generally conducted on sections from formalin-fixed, paraffin-embedded tissue that are cut between 8-15 µm. Spinal cord tissue is rich in myelinated axons and can be used as a positive control.

1. Dewax and hydrate sections to distilled water.
2. Put slides in freshly prepared Staining Solution at 56-60C for 30 minutes.
3. Wash slides well in water.
4. Partially differentiate in iron alum differentiating solution until myelin sheaths stand out blueish-black on a pale grey background, approximately 5 minutes. If you are unsure, check your sections under the microscope at1 minute intervals).
5. Wash slides in tap water for 10 minutes.
6. Complete differentiation in Weigert’s differentiator, 1 to 2 minutes. Control this differentiation step carefully checking under the microscope, until the myelin is an intense deep blue  against a creamy or clear background.
7. Wash well in tap water.
8. Dehydrate through a series of graded ethanol baths, clear in xylene, and mount.

RESULTS

Myelin-containing structures will be stained black, red blood cells will be black, nuclei will be blue, and the background should be clear or yellow.

Solutions

Haematoxylin Solution, working strength

• 10g Haematoxylin
• 100 ml absolute ethanol.

Allow solution to “ripen” naturally for six (6) weeks, forming the basis of a stock solution. Just prior to staining, create a working strength solution by diluting the stock 1:4 with distilled water

Iron Alum Solution

• 4% (w/v) aqueous ferric ammonium sulphate.

Staining Solution

Mix equal volumes of preheated (56-60C) working strength Haematoxylin and Iron Alum solutions just prior to use.

Weigert’s Differentiator, 200 ml

• Borax (sodium tetraborate), 2g
• Potassium ferricyanide, 2.5g
• Distilled water, 200ml

1. Take up bulk quantities of diaminobenzidine tetrahydrochloride dehydrate in water and bulk quantities of the free base in 0.1 M hydrochloric acid so that the concentration of DAB does not exceed 0.9 mg/ml. Dilute solutions with the same buffer, if necessary, so that the concentration does not exceed 0.9 mg/ml.
2. For each 10 ml of solution, add 5 ml of 0.2 M potassium permanganate solution (31.6 g KMnO4 per liter of solution with water) and 5 ml of 2 M sulfuric acid solution (112 ml concentrated H2S04 per liter of solution with water).
3. Allow the mixture to stand for at least 10 hours.
4. Test to ensure pH of solution is between 6-9. Dispose solution down the drain with copious amounts of water.

In molecular biology, c-Fos is a cellular proto-oncogene belonging to the immediate early gene family of transcription factors. c-Fos has a leucine-zipper DNA binding domain, and a transactivation domain at the C-terminus. Transcription of c-Fos is upregulated in response to many extracellular signals, e.g. growth factors. Additionally, phosphorylation by MAPK, PKA, PKC or cdc2 alters the activity and stability of c-Fos. Members of the Fos family dimerise with Jun to form the AP-1 transcription factor, which upregulates transcription of a diverse range of genes involved in everything from proliferation and differentiation to defense against invasion and cell damage.

The AP-1 complex has been implicated in transformation and progression of cancer, and both Fos and Jun were first discovered in rat fibroblasts.

The viral homologue of c-Fos, v-Fos, is found in the retrovirus Finkel-Biskis-Jinkins murine osteogenic sarcoma virus. In neuroscience research, neuroscientists measure expression of c-fos as an indirect marker of neuronal activity because c-fos is often expressed when neurons fire action potentials.

Staining procedure

1. This is a free-floating staining procedure for formalin-fixed brain tissue. Sections should be cut between 15-30 µm.
2. Transfer sections in 6-well plates loaded with PBS 0.1 M (one brain per well).
3. Rinse sections twice, 10 minutes each rinse, with PBS 0.1 M on a shaker.
4. Incubate sections with fresh 0.3% H₂O₂ in PBS 0.1 M for 30 minutes at room temperature on a shaker.
5. Rinse sections 3 x 10 minutes with PBS 0.1 M on a shaker.
6. Incubate sections with blocking solution  for 60 min at room temperature on a shaker.
7. Incubate sections with primary antibody diluted in blocking solution overnight at room temperature on a shaker.  With certain antibodies, to reduce background staining, consider an incubation for 2-3 days at 4°C.
8. Rinse sections 4 x 10 minutes with PBS 0.1 M on a shaker.
9. Incubate sections with biotinylated secondary antibody, diluted in blocking solution for 2 hours at room temperature on a shaker.
10. Rinse sections 4 x 10 minutes in PBS 0.1 M on a shaker.
11. Prepare ABC solution  at least 30 minutes prior to incubation to allow for ABC complex to form. Add 2 drops of solution A and 2 drops of solution B per 10 ml of blocking solution. Solutions A and B can also be added to plain PBS 0.1 M.
12. Incubate sections in ABC solution for 1-2 hours at room temperature on a shaker.
13. Rinse sections 4 x 10 minutes with PBS 0.1 M on a shaker.
14. Incubate sections in DAB solution for 8 minutes at room temperature on a shaker. DAB solution is highly toxic and carcinogen. Wear gloves and handle with care.
15. Add three drops of 0.3%  H₂O₂ (~125 ul) to each well to reveal staining. When background is satisfactory (after 1 to 5 min), halt the reaction by adding PBS 0.1 M.
16. Rinse sections 4 x 10 minutes with PBS 0.1 M on a shaker.
17. Transfer sections to slides using a brush, allow to air dry. It is best to transfer sections as soon as possible but well plates can be stored for a few days in the fridge at 4°C.
18. Dehydrate slides twice in ethanol 100% for 5 minutes each.
19. Incubate slides twice in toluene or xylene for 5 minutes each.
20. Add mounting medium to slides while still wet. Place coverslips to slides and allow to dry. Examine staining by microscopy.

Reagents

• Sodium phosphate, monobasic anhydrous NaH₂PO₄ (FW 120.0). Sigma,  S-0751, 1Kg
• Sodium phosphate, dibasic anhydrous, Na₂HPO₄ (FW 142.0). Sigma, S-0876, 1Kg
• Hydrogen peroxide, H2O2  30% (w/w) solution. Sigma, H-1009, 100 ml
• Albumin Bovine fraction V, min 96%, electrophoresis. Sigma, A-9647, 50g
• 3,3′-diaminobenzidine tablets (DAB). Sigma, D-5905, 50 tablets
• Goat serum. BioWest, Cat# S2000, 100ml or similar
• Vectastain ABC Kit, Elite standard. Vector, PK-6100
• Triton X-100 (t-Octylphenoxypolyethoxyethanol). Sigma, T-9284, 100 ml
• Toluene or xylene from VWR or Fisher
• Ethanol 100%

Antibodies

Titrate new batches of antibodies for appropriate concentration before using in experiments as effective concentrations may vary across batches of antibody?.

Primary

Rabbit anti-Fos polyclonal IgG, Oncogene Research Products (Ab-5, Cat.# PC38). Recommended dilution, 1:20 000.

Secondary

Biotin-SP-conjugated affiniPure Goat anti-rabbit IgG (H+L) (minimal cross reaction to Human , Mouse and rat serum proteins). Made in goat. Jackson Immunoresearch, Cat.# 111-065-144. Recommended dilution: 1:2000.

Solutions

Phosphate buffer solution, 0.2 M,  pH 7.4

1. Collect 1000 ml of distilled water in a graduated cylinder. Pour about 400 ml of water in a beaker and stir.
2. Weigh 4.8 g of Sodium Phosphate monobasic NaH₂PO₄ and 22.72 g of sodium phosphate dibasic Na₂HPO₄ .
3. Add to the 400 ml of water. When dissolved, add the rest of the water and continue stirring for 5 min. Take pH which should be around 7.4.

Phosphate buffer solution, 0.1 M,  pH 7.4

Make phosphate buffer 0.2M solution as described above and add 1000 ml of distilled water to bring it to 0.1 M, total volume 2 liters. pH should be around 7.4. Solution can be kept at room temperature or at 4°C.

Blocking solution (PBS 0.1 M; 0.1 % BSA; 0.2% Triton X-100; 2% serum)

Collect about 800 ml of phosphate buffer 0.1 M in a graduated cylinder. Add 20 ml of serum, 2 ml of Triton X-100 and 1 g of BSA. Stir for 10 min. Add more PBS 0.1 M to reach 1000 ml. Stir another 5 min. Store blocking solution in 50-ml aliquots (50-ml Falcon tubes) at -20°C

DAB solution, 0.05% (w/v)

Add 1 tablet (10 mg) of DAB in 20 ml of PBS 0.1 M in a 50-ml Falcon tube. Vortex vigorously until dissolved. Solution should be used fresh, or may be frozen in single-use aliquots and stored at -20C until use. Wear gloves and inactivate solution using a 10% bleach solution (dilute DAB with an equal volume of bleach) when finished and dispose in appropriate biohazard container. DAB is highly toxic and carcinogen; do not dump solution down the drain without treatment.

Neutralization of DAB

Although chlorine bleach is commonly employed in many laboratories as a neutralization procedure, it is not effective in removing the mutagenic properties of DAB. A potassium permanganate-sulfuric acid procedure must be used.

1. Take up bulk quantities of diaminobenzidine tetrahydrochloride dehydrate in water and bulk quantities of the free base in 0.1 M hydrochloric acid so that the concentration of DAB does not exceed 0.9 mg/ml.  Dilute solutions with the same buffer, if necessary, so that the concentration does not exceed 0.9 mg/ml.
2. For each 10 ml of solution, add 5 ml of 0.2 M potassium permanganate solution and 5 ml of 2 M sulfuric acid solution.
3. Allow the mixture to stand overnight, decolorize by the addition of sodium ascorbate, neutralize and dispose solution down the drain with copious amounts of water.

0.3% (v/v) H₂O₂ solution

Add 0.5 ml of H₂O₂ 30% solution to 50 ml of PBS 0.1 M in a 50-ml Falcon tube. Vortex. Use fresh.

Equipment

• Microscope
• 2D Shaker
• 6-well plates
• Gelatin-coated slides or precleaned superfrost plus slides (25 x 75 x 1 mm). VWR, Cat.# 48311-703
• Coverlips (micro cover glasses) 24 x 60 mm, No. 1. VWR, Cat.# 48404 454.
• Mounting medium (Eukit or Cytoseal 280 from Richard-Allan Scientific (8311-4) or similar)