Staining

The embedding process must be reversed in order to get the paraffin wax out of the tissue and allow water soluble dyes to penetrate the sections. Therefore, before any staining can be done, the slides are "deparaffinized" by running them through xylenes (or substitutes) to alcohols to water. There are no stains that can be done on tissues containing paraffin.

The staining process makes use of a variety of dyes that have been chosen for their ability to stain various cellular components of tissue. The routine stain is that of hematoxylin and eosion (H and E). Other stains are referred to as "special stains" because they are employed in specific situations according to the diagnostic need.

Slides being stained on automated stainer.

Slides being stained on automated stainer.

Frozen sections are stained by hand, because this is faster for one or a few individual sections. The stain is a "progressive" stain in which the section is left in contact with the stain until the desired tint is achieved.

Staining a frozen section.

H and E staining

Hematoxylin is the oxidized product of the logwood tree known as hematein. Since this tree is very rare nowadays, most hematein is of the synthetic variety. In order to use it as a stain it must be "ripened" or oxidized. This can be done naturally by putting the hematein solution on the shelf and waiting several months, or by buying commercially ripened hematoxylin or by putting ripening agents in the hematein solution.

Hematoxylin will not directly stain tissues, but needs a "mordant" or link to the tissues. This is provided by a metal cation such as iron, aluminum, or tungsten. The variety of hematoxylins available for use is based partially on choice of metal ion used. They vary in intensity or hue. Hematoxylin, being a basic dye, has an affinity for the nucleic acids of the cell nucleus.

Hematoxylin stains are either "regressive" or "progressive". With a regressive stain, the slides are left in the solution for a set period of time and then taken back through a solution such as acid-alcohol that removes part of the stain. This method works best for large batches of slides to be stained and is more predictable on a day to day basis. With a progressive stain the slide is dipped in the hematoxylin until the desired intensity of staining is achieved, such as with a frozen section. This is simple for a single slide, but lends itself poorly to batch processing.

Eosin is an acidic dye with an affinity for cytoplasmic components of the cell. There are a variety of eosins that can be synthesized for use, varying in their hue, but they all work about the same. Eosin is much more forgiving than hematoxylin and is less of a problem in the lab. About the only problem you will see is overstaining, especially with decalcified tissues.

Coverslipping

The stained section on the slide must be covered with a thin piece plastic or glass to protect the tissue from being scratched, to provide better optical quality for viewing under the microscope, and to preserve the tissue section for years to come. The stained slide must go through the reverse process that it went through from paraffin section to water. The stained slide is taken through a series of alcohol solutions to remove the water, then through clearing agents to a point at which a permanent resinous substance beneath the glass coverslip, or a plastic film, can be placed over the section.

Coverslipping.

Coverslipping.

Decalcification

Some tissues contain calcium deposits which are extremely firm and which will not section properly with paraffin embedding owing to the difference in densities between calcium and parffin. Bone specimens are the most likely type here, but other tissues may contain calcified areas as well. This calcium must be removed prior to embedding to allow sectioning. A variety of agents or techniques have been used to decalcify tissue and none of them work perfectly. Mineral acids, organic acids, EDTA, and electrolysis have all been used.

Strong mineral acids such as nitric and hydrochloric acids are used with dense cortical bone because they will remove large quantities of calcium at a rapid rate. Unfortunately, these strong acids also damage cellular morphology, so are not recommended for delicate tissues such as bone marrow.

Organic acids such as acetic and formic acid are better suited to bone marrow, since they are not as harsh. However, they act more slowly on dense cortical bone. Formic acid in a 10% concentration is the best all-around decalcifier. Some commercial solutions are available that combine formic acid with formalin to fix and decalcify tissues at the same time.

EDTA can remove calcium and is not harsh (it is not an acid) but it penetrates tissue poorly and works slowly and is expensive in large amounts.

Electrolysis has been tried in experimental situations where calcium had to be removed with the least tissue damage. It is slow and not suited for routine daily use.

Artefacts in Histologic Sections

A number of artefacts that appear in stained slides may result from improper fixation, from the type of fixative, from poor dehydration and paraffin infiltration, improper reagents, and poor microtome sectioning.

The presence of a fine black precipitate on the slides, often with no relationship to the tissue (i.e., the precipitate appears adjacent to tissues or within interstices or vessels) suggests formalin-heme pigment has formed. This can be confirmed by polarized light microscopy, because this pigment will polarize a bright white (and the slide will look like many stars in the sky). Formalin-heme pigment is most often seen in very cellular or bloody tissues, or in autopsy tissues, because this pigment forms when the formalin buffer is exhausted and the tissue becomes acidic, promoting the formation of a complex of heme (from red blood cells) and formalin. Tissues such as spleen and lymph node are particularly prone to this artefact. Making thin sections and using enough neutral-buffered formalin (10 to 1 ratio of fixative to tissue) will help. If the fixative solution in which the tissues are sitting is grossly murky brown to red, then place the tissues in new fixative.

The presence of large irregular clumps of black precipitate on slides of tissues fixed in a mercurial fixative such as B-5 suggests that the tissues were not "dezenkerized" prior to staining. These black precipitates will also appear white with polarized light microscopy.

Tissues that are insufficiently dehydrated prior to clearing and infiltration with paraffin wax will be hard to section on the microtome, with tearing artefacts and holes in the sections. Tissue processor cycles should allow sufficient time for dehydration, and final ethanol dehydrant solution should be at 100% concentration. In humid climates, this is difficult to achieve. Covering or sealing the solutions from ambient air will help. Air conditioning (with refrigerants, not with evaporative coolers) will also reduce humidity in the laboratory. Toluene as a clearing agent is more forgiving of poorly dehydrated tissues, but it is more expensive and presents more of a health hazard than other non-xylene clearing agents

Though alcohols such as ethanol make excellent fixatives for cytologic smears, they tend to make tissue sections brittle, resulting in microtome sectioning artefacts with chattering and a "venetian blind" appearance.

Bubbles under the coverslip may form when the mounting media is too thin, and as it dries air is sucked in under the coverslip. Contamination of clearing agents or coverslipping media may also produce a bubbled appearance under the microscope.

Artefact with undezenkerized tissue.

Problems in Tissue Processing

"Floaters" are small pieces of tissue that appear on a slide that do not belong there--they have floated in during processing. Floaters may arise from sloppy procedure on the cutting bench-- dirty towels, instruments, or gloves can have tissue that is carried over to the next case. Therefore, it is essential that you do only one specimen at a time and clean thoroughly before opening the container of the next case.

The best way to guard against unrecognized floaters is to always separate like specimens in the numbering sequence. For example, if you have three cases with prostate chips, separate them in accessioning with totally different specimens such as uterus or stomach. That way, if numbers are transposed or labels written wrong or tissue carried over, then you will have an obvious mismatch. Carrying over one prostate to another, or transposing the numbers of identical tissues may never be recognized.

If reusable cassettes are employed, you must be aware that tissue may potentially be carried over and appear as "floaters" even several days later, when the cassette is re-used. The problem arises when, during embedding, not all the tissue is removed from the cassette. Then, in the cleaning process, not all of the wax is removed. Then, the next person using the cassette does not pay attention to the fact that there is tissue already in the cassette and puts his specimen in it. The floater that appears on the slide will look well-preserved--it should, because it was processed to paraffin.

Always be sure that you properly identify the tissue! This means that you make sure that the patient label on the specimen container matches that of the request slip. An accession number is given to the specimen. This number must appear with the tissue at all times. You must never submit a cassette of tissue without a label. You must never submit a cassette of tissue with the wrong label. Mislabelling or unlabelling of tissues is courting disaster.

Safety in the Lab

The lab should be well-ventilated. There are regulations governing formalin and hydrocarbonds such as xylene and toluene. There are limits set by the Occupational Safety and Health Administration (OSHA) that should not be exceeded. These limits have recently been revised to reduced levels.

Every chemical compound used in the laboratory should have a materials safety data sheet on file that specifies the nature, toxicity, and safety precautions to be taken when handling the compound.

The laboratory must have a method for disposal of hazardous wastes. Health care facilities processing tissues often contract this to a waste management company. Tissues that are collected should be stored in formalin and may be disposed by incineration or by putting them through a "tissue grinder" attached to a large sink (similar to a large garbage disposal unit).

Every instrument used in the laboratory should meet electrical safety specifications (be U.L. approved) and have written instructions regarding its use.

Flammable materials may only be stored in approved rooms and only in storage cabinets that are designed for this purpose.

Fire safety procedures are to be posted. Safety equipment including fire extinguishers, fire blankets, and fire alarms should be within easy access. A shower and eyewash should be readily available.

Laboratory accidents must be documented and investigated with incident reports and industrial accident reports.

Specific hazards that you should know about include:

• Bouin's solution is made with picric acid. This acid is only sold in the aqueous state. When it dries out, it becomes explosive.

• Many reagent kits have sodium azide as a preservative. You are supposed to flush solutions containing sodium azide down the drain with lots of water, or there is a tendency for the azide to form metal azides in the plumbing. These are also explosive.

• Benzidine, benzene, anthracene, and napthol containing compounds are carcinogens and should not be used.

• Mercury-containing solutions (Zenker's or B-5) should always be discarded into proper containers. Mercury, if poured down a drain, will form amalgams with the metal that build up and cannot be removed.