Fixation

Tissue fixation has a significant influence on immunohistochemistry as most antigens are altered during this process (Williams et al. 1997). The purpose of fixation is to preserve tissue and prevent further degradation by the action of tissue enzymes or microorganisms. As discussed in an earlier chapter, good fixation requires tissue to have adequate time in the fixative to allow the solution to penetrate, retaining uniform cellular detail throughout the tissue. However, in the routine laboratory this ideal may be compromised as it is difficult to define a standard tissue size, fixation time, and fixative for each specimen type. Tissues need to be adequately, but not over-fixed, so that antigenicity is preserved without excessive alteration. Prolonged fixation can result in the irretrievable loss of many antigens, particularly membrane-associated antigens such as CD20 and immunoglobulin (Ig) light chains (Miller et al. 1995; Ashton-Key et al. 1996). Lack of adequate fixation, or delay in fixation, may also be equally detrimental to labile antigens (Donhuijsen et al. 1990; von Wasielewski et al. 1998; GEFPICS-FNCLCC 1999).

Any fixative used must be compatible with immunohistochemical staining methods and formalin is still the most universal of fixatives. Formulations differ between laboratories and include 10% neutral buffered formalin (NBF), 10% formalin in tap water, 10% formal saline, and 10% NBF with saline (Angel et al. 1989; Williams 1993; Williams et al. 1997). Even though it may be the pathologist’s choice, it can create a challenge for demonstrating certain antigens. Dabbs (2006) characterizes formalin as:

Williams et al. (1997) investigated the effect of fixation on immunostaining to establish whether a specific preparation schedule would allow for the optimal demonstration of all antigens. Of the fixatives tested, 10% formal saline, 10% NBF (except for CD45RO), and 10% zinc formalin (except for CD3) gave the most consistent results overall and showed excellent antigen preservation. More recently, alcohol-based fixatives have been considered as an alternative to formalin (van Essen et al. 2010) and produced satisfactory immunohistochemistry staining. Other fixatives which may still be used in some laboratories include Bouin’s, B5 (mercury), zinc formalin, 10% formal-acetic and Carson’s, which also have an influence on the reproducibility of staining, each presenting a change in pH, length of required exposure and different artifacts.

Fixatives dictate many factors for immunohistochemical staining, such as dilution, antibody incubation time, retrieval method (if applicable), type of retrieval solution, and special pretreatments (e.g. pigment removal). Depending upon the type of fixative used, the protocols may require slight modifications. With the advent of heat-induced epitope retrieval (HIER) (Shi et al. 1991) many of the problems associated with fixation have been reduced and, in conjunction with automated techniques, good-quality staining is achievable on most tissue sections.

Processing

As with fixation, all tissue must be appropriately processed to produce successful immunohistochemical staining. Tissue that is inadequately processed will potentially produce poor-quality sections, with poor adhesion to the slides – especially fatty tissue such as breast and skin. Modern tissue processors all have the option to include vacuum and temperature variation at each step, allowing for greater optimization of the procedure. However, high temperatures can be detrimental to antigens that are heat labile. It is recommended that paraffin with a low-temperature melting point be used for this reason.

Regarding paraffin processing of tissues for immunohistochemistry, as with fixation, there is no standard protocol for the optimal demonstration of all antigens, as concluded by Williams et al. (1997). In a study of laboratories in the UK, Williams (1993) found nearly as many different schedules as the number of laboratories participating in the survey. Of the nine tissue-processing factors investigated, only two had any significant effect on immunoreactivity. Increasing the temperature of processing from ambient to 45°C, as well as longer processing times for dehydration and wax infiltration, were both found to improve immunostaining. Other factors including type of processor, type and quality of reagents, time in clearing agent, use of vacuum, most of which had been suggested as possible causes of poor processing (Horikawa et al. 1976; Trevisan et al. 1982; Anderson 1988; Slater 1988), were found to have no effect on subsequent immunohistochemistry.

Microwave processing is now being introduced into some laboratories to speed the processing time and reduce turnaround time for diagnostic specimens, and has been used successfully in conjunction with routine antibody staining. Acceptable staining was achieved when compared to tissues processed in a conventional processor (Emerson et al. 2006). As with all processing, if the tissue is not completely fixed then artifacts will be introduced.

One important point is that any control tissues used in the laboratory should be processed using the same protocols established for patient samples.

Reversal of fixation/epitope retrieval

The quality and reproducibility of immunohistochemical staining relies upon the reversal of fixation, which results in the targeted epitope being exposed, allowing for the antigen binding site to be available. The revolution of reversing the hydrogen cross-bonds formed by formalin was introduced by Shi et al. (1991). There are now numerous methods for epitope retrieval including protein digestion techniques or, more commonly, heating the slides in a buffered solution. Cattoretti et al. (1993) introduced the solution most commonly used in standardized retrieval methods. They used a citrate buffer at pH 6.0, which is inexpensive, stores easily, and is readily available commercially or easily prepared in the laboratory. Other buffers used include EDTA-based solutions at a higher pH range, which produce more intense staining of some antibodies. These methods have allowed for the successful demonstration of a much greater range of antigens in tumors, including proliferation markers and oncogene expression. The use of automated immunostainers has brought greater standardization of retrieval methods, as these use standard retrieval solutions with defined reproducible protocols. Non-automated laboratories may have a number of variables that require internal standardization in the antigen retrieval technique including the choice of heating method (e.g. pressure cooker, microwave, etc.), retrieval solution, pH, temperature, volume of the fluid, and the temperature and exposure time while heating and cooling slides.

Equipment commonly used to perform epitope retrieval includes the modified pressure cooker, initially reported by Norton et al. (1994), microwaves, waterbath or a pretreatment module. Some automated platforms have on-board retrieval where individual slide bays can be heated with the appropriate solution on the slide.

Other factors required for successful retrieval include the proper drying and complete removal of water from slides. In addition to avoiding wrinkles or tears in the tissue, these factors will all assist the adhesion of tissue to the slide. With respect to enzymatic proteolytic ‘epitope retrieval’, such as trypsin digestion prior to immunostaining, the choice of enzyme usually dictates the temperature and pH of the solution, as different enzymes have different preferential pH and temperatures. For example, the optimal values for a mammalian-derived trypsin are pH 7.8 at 37°C, with 0.1% calcium chloride included as an activator (Huang et al. 1976). The concentration of enzyme required is dependent on the proteolytic qualities of the product being used. A typical concentration used for many commercial trypsins employed in immunohistochemistry protocols is 0.1%. The concentration, pH, and temperature are then usually held constant, while the time of digestion is varied. The time required for optimal digestion will vary, depending on the antigen under investigation, the quality (proteolytic capabilities) of the trypsin and the length of formalin fixation. For antigens that are only present in small amounts, e.g. immunoglobulin light chains on the surface of B cells, the time for optimal digestion may vary from case to case, depending on how long each case has been fixed in formalin.

Buffers and diluents

The buffer and diluent used for wash steps and antibody dilution will affect the results of immunostaining. The pH of these reagents needs to be monitored and must be checked and documented prior to use. If it falls outside of the range proscribed by the established protocol, corrections must be made or the reagent discarded.

Many antibody diluents contain additives such as sodium azide to stabilize and maintain the protein. Although this extends the shelf life of the antibody, the additives may interfere with, or inhibit, staining if present at excessive levels.

Antibodies

The storage temperature of an antibody is critical to its stability. Commercially available antibodies should be accompanied by a specification sheet, containing storage and handling instructions.

Concentrated antibodies tend to have a longer shelf life than pre-diluted ‘ready to use’ antibody preparations. Concentrated antibodies can be mixed with glycerine to prevent ice crystal formation, aliquoted into cryovials, then ‘snap’ frozen and stored in a −80°C freezer. This greatly extends their shelf life; compliance with the expiry date printed on the antibody packaging is important. The storage temperature for antibodies and reagents should also be monitored closely, as any fluctuation in temperature may cause increased deterioration of reagents. Frost-free −20°C freezers should hence be avoided for the storage of antibodies due to damage caused by the freeze-thaw cycles that these types of freezer perform.

Block and slide storage conditions

Processed blocks should be kept in a cool dry place. Resealing paraffin blocks following cutting can help protect the tissue from everyday elements such as air drying, excessive moisture or physical damage. The use of fresh cut controls for each run would be ideal but this is not always feasible in a busy laboratory. Repeated sectioning of a control tissue block can also result in loss of usable tissue, although some laboratories now have a control on the same slide as the test section.

Pre-cutting control slides is more time efficient and serial sections result in minimal tissue loss. The correct storage of pre-cut slides is important and frequently overlooked as a potential source of error in staining. Some studies have found deterioration of antigens in stored sections (Raymond & Leong 1990; Bromley et al. 1994; Prioleau & Schnitt 1995). Others have found no deterioration of some markers investigated, including estrogen receptor (ER), CD3, CD20, CD45RO, vimentin, and Ig light chains, in sections stored for up to four months at room temperature (Williams et al. 1997; Eisen & Goldstein 1999).

The viability of the antigen and speed of antigen deterioration in cut tissue sections is highly dependent upon the antigen under consideration and the temperature used for section adhesion. For example, whilst antigens such as CD30 and PSA seem relatively robust, CD117 (C-kit) deteriorates quickly and should be cut fresh.

The temperature at which the slides are dried can also affect the immunoreactivity of the antigens. It is advisable to dry all slides for immunohistochemical staining at 37°C. Urgent cases can generally be dried at 60°C for up to 4 hours. The exception to this is HER2, where it is recommended that sections should not be dried at 60°C for more than 1 hour.

An appropriate number of control slides and blocks should be kept with these factors in mind. This stock level will vary depending on the workload in each individual laboratory.