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.
| Tissue preparation |
| Thickness | No more than 3 mm thick. |
| Area | 20 mm × 30 mm. |
| Fixed tissue | Cut large organs into 3 mm slices and store in neutral buffered formalin for 48 hours. Select tissue from fixed areas, trim to size and refix until the evening. If the trimmed sample is visibly unfixed, refix for a further 24 hours. |
| Unfixed tissue | Slices of tissue should be thoroughly fixed before processing. |
| Times | All times in processing fluids for this schedule are for tissues 3 mm thick or less. Tissues thicker than that will require longer times. |
| Clearing agent | Xylene or another clearing agent that will clear tissues in similar times should be used. |
| Processing time | This schedule takes 12 hours, and processes overnight. On weekends tissues should be left in fixative until Sunday evening with a 48 hour delay. |
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.
The tissue and paraffin attached to the cassette has formed a block, which is ready for sectioning.Tissue blocks can be stored at room temperature for years.
Sectioning tissues
Tissues are sectioned using a microtome. Turn on the water bath and check that the temp is 35-37ºC. Use fresh deionized water (DEPC treated water must be used if in situ hybridization will be performed on the sections). Blocks to be sectioned are placed face down on an ice block or heat sink for 10 minutes. Place a fresh blade on the microtome; blades may be used to section up to 10 blocks, but replace if sectioning becomes problematic. Insert the block into the microtome chuck so the wax block faces the blade and is aligned in the vertical plane.
Set the dial to cut 10 µM sections to order to plane the block; once it is cutting smoothly, set to 5 µM
sections . The blade should angled at 5º. Face the block by cutting it down to the desired tissue plane and discard the paraffin ribbon. If the block is ribboning well then cut another four sections and pick them up with forceps or a fine paint brush and float them on the surface of the 37ºC water bath. Float the sections onto the surface of clean glass slides. If the block is not ribboning well then place it back on the ice block to cool off firm up the wax. If the specimens fragment when placed on the water bath then it may be too hot.
Place the slides with paraffin sections on the warming block in a 65°C oven for 20 minutes (so the wax just starts to melt) to bond the tissue to the glass. Slides can be stored overnight at room temperature.
Haematoxylin Eosin (H&E) staining
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 ofhemalum, 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.
Solutions:
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.
H-uracil, a specific precursor to RNA. Subsequent to this incubation, extract rRNA from the sample and purify through differential centrifugation, column chromatography or electrophoresis. Dissolve the radioactive RNA probe in 4X Saline-Citrate containing 50% formamide to yield a sample that has 50,000 to 100,000 counts per minute, per 30 microliter sample, as determined with a scintillation counter. Add the formamide is added to prevent the aggregation of RNA. 
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