Troubleshooting in immunohistochemistry (IHC) can be highly complex and is more than "what's wrong with or why didn't my slides stain?" There are many issues affecting IHC quality and the ability of technical staff to troubleshoot. Some issues related to IHC problems are: deviating from established IHC protocols, no standardization of IHC protocols, introducing multiple variables in the IHC process, antigen retrieval optimization, improper antigen retrieval techniques and use of heat sources.
There are problems associated with having inexperienced staff members. There are huge differences between laboratories and how they are set up to perform IHC. Budget restraints, lack of automation and high volume case loads are another contributing factor with IHC troubleshooting.
Figures/courtesy Jim Burchette
Think about the ever popular buzz word "Lean?" Imagine a highly automated laboratory that has a Lean staff to start with and a lab member is on scheduled time off. Do they have time to perform in depth troubleshooting? Maybe, but the daily process must continue or the system backs up or fails.
One laboratory might be the customer who relies on the vendor to help with troubleshooting, whereas another lab may have experienced technologists on staff with advanced troubleshooting abilities. What about small community hospitals where on-the-job trained histotechnicians must perform the entire histology laboratory functions and IHC?
I am fortunate to work with physicians who have a very strong knowledge of IHC. We are able to brainstorm issues and come up with solutions. However, I have seen many residents over the years go through the program and have no interest in immunohistochemistry. I try to emphasize to pathology residents and pathologist assistants the importance of understanding the technical IHC applications. After all, where will their training take them and what kind of role will they be asked to participate in?
Background or non-specific staining is a common concern in many laboratories. Problems with background in tissue sections can be a result of poor fixation and processing. Not only does this cause problems with tissue adhering to slides, but poor fixation is also a disservice to our patients. I am glad to see fixation guidelines for HER2/neu immunohistochemistry, but what about other tissues, such as sentinel lymph nodes?
I recall a biopsy taken from a past U.S. president. The news media asked the pathologist about the diagnosis, but his response was the specimen needed to be properly fixed before studies could be performed. If that is good enough for our president, why isn't it good enough for the average citizen? Proper fixation only improves the standard of care we all strive for.
Other background issues can come from over retrieving, using the wrong retrieval solution, improper reagent pH, endogenous biotin, concentrated antibodies used too strong and using the wrong non-immune blocking serum when using a more traditional detection system.
Historically, non-immune serum from the host species of the secondary antibody is used as a protein block prior to the application of the primary antibody.
Background issues typically arise when using esoteric antibodies and custom prepared detection systems, such as an avidin-biotin complex (ABC) and biotin-streptavidin. There may be a need to include fragment crystalline (FC) portion if the immunoglobulin molecule as a blocking step to reduce non-specific staining.
What about background staining when a non-biotin, labeled polymer system is used? I rarely see background issues with labeled polymer systems.
Universal blocking solutions, available from various manufactures, can be incorporated into the staining protocol when background is encountered.
What is perceived as background could be diffused antigen in the tissue section, such as the case of some viruses that are degenerating or kappa and lambda antibodies reacting with serum proteins. One should be able to recognize necrosis and the associated background.
On a whole, the newer polymer detection systems are of such high quality, sensitivity and cleanliness, background issues are non-existent. Because of this improved technology, there are voices in the IHC community pushing for revising the College of American Pathologist guidelines regarding the exclusion of running negative controls except in special situations.
Having a laboratory system in place where the IHC process is highly controlled or regulated can lead to standardization if the system is understood by the laboratory staff and protocols are adhered to. Automated IHC is the perfect example where modifications of programs can be blocked by unauthorized personnel. This type of system allows a direct approach to troubleshooting when issues arise.
An IHC problem in this system could be the result of a mechanical or software issues, or an incorrect reagent used somewhere in the process. Another cause of background or faulty reactions can arise from contaminated buffer or water containers. Reagent bottles should be properly cleaned before refilling and put into use. Bacteria growth in a reagent bottle can affect the solution pH and cause altered reactions. Improper reagent preparation can result in a wrong pH that affects the end results.
Laboratories should exercise proper pH meter use. Electrodes should be stored in electrode storage solution and not pH 7.0 buffer standards or deionized water. Electrode storage solution is a specialized solution designed to keep the electrode stable, to have a fast response rate and to prevent fluctuations. I realize many laboratories do not use pH meters and rely on pH indicator strips. Some companies want a questionable reagent returned so it can be checked in house for problems.
Errors may be a result of a user-filled container containing the wrong reagent or a miscalculation of user-diluted antibody preparation. I'm not saying this is true across the board, but I have seen many technicians who are challenged with laboratory math.
After proper instruction, understanding of basic math formulas and daily use, they should become proficient in diluting antibodies and preparing laboratory reagents. I often quiz new employees with math questions, not to intimidate, but to gain an understanding of their level of knowledge and where I need to focus on training.
Antibody dilutions seem to be a troublesome area of math for some techs whom have not had need for this math in previous areas of employment. Here is my approach to a simple rule of thumb for antibody dilutions. First, determine the volume of diluted antibody needed. Divide this volume by the antibody dilution factor. This will give you the amount of antibody needed for this preparation. Subtract the antibody volume from the total volume for the amount of diluent needed to prepare this dilution.
Check your math by adding the antibody volume and the diluent volume. This should equal to total volume of diluted antibody. Not all dilutions pertain to antibodies. We all receive buffer and retrieval reagents in various stock concentrations. The same formula pertains to liter size and greater volumes. A quick reference chart can be prepared once the dilutions and volumes have been worked out.
Pigments can often interfere with interpretation of true signal as in the case of melanin, hemosiderin and anthracotic pigment. If diaminobenzidine (DAB) is used as the chromogen of choice in a section of tissue containing melanin, a giemsa counter stain can be applied to the tissue post DAB. This will color the melanin pigment green, giving better contrast to the brown DAB signal. Alkaline phosphatase (AP) labeled detection systems and fast red chromogens create a vivid red immune complex signal easily differentiated from pigments. AP detection also allows easier visualization of small bacteria, such as spirochetes and Rickettsia rickettsia.
Some Final Issues
IHC signal enhancement can be accomplished in several manners. Incubation with primary antibody can be extended to overnight at 4° C. Sometimes, applying the primary antibody twice will result in a more intense signal. Extending the chromogen reaction time as long as it does not contribute to background is another option.
The use of a metal enhancing, post DAB solution in combination with a lighter hematoxylin or methyl green counter stain will offer better contrast. Catalyzed signal amplification reagents are available, but I find them best suited for select applications and not routine IHC.
A host of technical artifacts can lead to slide and staining problems. Debris artifacts such as graphite deposits, wax pencil debris and squamous cells floaters can be problematic. Staining or background on the edge of the tissue section can be a result of the tissue section drying or a difference in penetration of fixative.
When manually performing endogenous peroxidase activity quenching, it is important to drain the slides prior to placing them in the hydrogen peroxide solution, especially if multiple racks of slides will go through the H2O2 solution. The addition of water or methanol will further dilute the H2O2 and cause it to not properly block the endogenous peroxidase activity in red blood cells and neutrophils.
Blocking endogenous peroxidase activity is traditionally performed after paraffin removal and before primary antibody. The use of high concentrations of H2O2 can lead to a reduced signal with some antibodies such as CD4. A solution of H2O2 less than three percent is preferred and can be applied anytime after the application of primary antibody as an alternative.
Jim Burchette is an analytical specialist, Immunopathology Lab, Molecular Pathology lab group, Duke University Health System, Clinical Laboratories, Durham, NC.
Burchette would like to thank the following people for their contributions to this article: David Howell, MD, PhD; Volker Nickeleit, MD; Sheron Lear, HT, HTL(ASCP)QIHC; and Billie Zimmerman, MT(ASCP)QIHC.