Troubleshooting Immunohistochemical staining

When tissue staining has not given the expected results, the experiment should be examined in a systematic way, wherein only single experimental variables are altered at one time. Proper immunohistochemical troubleshooting requires one to determine whether difficulties are related to specimen, antibodies, technique, environment, or slide interpretation? The following checklist may assist in troubleshooting efforts.

No Staining of Either Controls or Specimen

• Confirm that no reagents were omitted (primary antibody, secondary antibody, substrate components)
• Confirm that reagents were added in the correct order, and for sufficient incubation times.
• Re-read labels to confirm that correct antibodies were used. This is especially important when using primary/ secondary antibody combinations. For example, when using a mouse IgM primary antibody, the secondary antibody should be a goat or rabbit anti-mouse IgM (not IgG).
• Check antibody titrations and dilutions. This is particularly important for the primary antibody.
• Check that antibody blocking solutions or diluents do not contain substances that may interfere with or absorb the primary antibody. For instance, serum containing diluents can absorb out or reduce primary reactivity to small molecules such as GABA or amino acids.
• Check reagent expiration dates and storage. Enzymes and fluorochromes are especially prone to breakdown after prolonged storage. Antibodies should be stored in nondefrosting freezers, as self-defrosting models will expose antibodies to repeated freeze/thaw, resulting in antibody breakdown.
• Check specimen storage. Where possible, compare staining of the unknown specimen versus a known positive tissue in a “side-by-side” experiment.
• Check the chromogen/substrate solution. This can be checked by adding a drop of the labeling reagent to a small sample of prepared chromogen. If the chromogen is working, the mixture should change color. Chromogen solutions can deteriorate quickly, so do not use beyond the times recommended by the manufacturer. Lack of color change may be due to inactive enzyme or improperly prepared chromogen.
• The rinse buffer may be incompatible with the reaction reagents. The pH must be appropriate, and buffers to be used with peroxidase enzyme should not contain NaN3.
• The counterstain and mounting media may not be compatible with the chromogen. Check the manufacturer’s recommendations.
• Check that the microscope is adjusted correctly and that the fluorescence lamp is not burned out.

Weak Staining

Points to consider are:

1. Is the intensity of the staining consistent between the positive controls and the test sample(s)?
2. Is the staining specific for the antigen of interest, or is it background staining? This can only be determined by examining the slides.

All the items listed above for No Staining can apply to a lesser degree to the situation of weak staining. However, if the negative controls are devoid of stain and the positive controls and test sample(s) are weakly stained, then possible trouble points include:

• Overfixation, or incorrect fixation for the immunological procedure in use
• Insufficient antigen retrieval
• Antibody concentration may be too dilute. If possible, the concentration should be increased. If this is not feasible, then the incubation time or temperature may need adjusting. When diluted antibody is stored in the refrigerator it sometimes gets absorbed to the walls of the container. Storing the antibody with a protein carrier such as 1%–3% BSA can alleviate this situation.
• Too much buffer rinse has been left on the slide, so that the antibody becomes diluted when added to the sample.

If the negative controls have not reacted, the positive controls are well stained, but the test sample is stained weakly, then either the positive control and the test sample were fixed differently, are of different tissue type, or the outer tissue of the test specimen block has been poorly fixed.

If the negative controls have not reacted, the test sample(s) are well stained, but the controls are weakly stained, then the control material should be replaced.

Background Staining

If the negative control is being stained as well as the positive controls and test sample(s), then the degree and type of background staining must be analyzed. The following are possibilities for investigation:

1. Re-titer antibodies (both primary and secondary) with a dilution series.
2. Incubate with chromogen for a shorter time. Some chromogens, such as DAB, develop very quickly.
3. The chromogen was not totally dissolved, and associated with the tissue. Centrifuge or filter the chromogen solution.
4. Particulates in the antibody solution. These may form upon repeated freeze/thaw and can be eliminated by centrifugation.
5. Insufficient rinsing between steps, or contaminated buffers. Mix new buffers and increase washing steps.
6. Enzyme or biotin in the tissue is reacting with the reagent. This can be prevented by increasing the time or concentration of block, trying different types of block, or using a combination of more than one block, or changing the staining methods. Some tissues (i.e. brain and liver) are known to contain high endogenous levels of biotin or peroxidase activity.
7. The incorrect blocking serum was used, or blocking serum was not used. The blocking serum should be from the species of the secondary antibody. It is possible to use 5% nonfat dry milk or BSA rather than serum.
8. The secondary antibody cross-reacts with endogenous tissue proteins. Secondary antibodies which have been absorbed against immunoglobulin from the species from which the target tissue was obtained will result in significantly lower background, and are indespensible for double-labeling experiments. It is also critical to run a no-primary antibody control to determine if the secondary antibody is the source of the background.
9. Hydrophobic and/or ionic interactions between the reagents and tissue types such as connective, adipose or fatty tissues may give rise to apparent specific reactions. Antigen retrieval procedures can be of great assistance in correcting this predicament. A decreased fixation time in formalin can also help.
10. The embedding media may not be completely removed from the tissue. Review the removal procedure for possible changes.
11. The specimen may have dried out during the procedure, allowing the trapping of reagents under the edges of the specimen. Care should be taken to avoid letting the specimens dry.

If there is background staining in the positive controls and the test sample(s), but not in the negative control, then the issue is most likely associated with the primary antibody. Some possibilities are:

1. The primary antibody was too concentrated, or the incubation period too long. More dilute antibody, or shorter incubation or lower incubation temperature may correct the situation.
2. The tissue may contain Fc receptors, or there may be interfering Ig components (aggregates or oligomers) or there may by naturally occurring, contaminating antibodies. This can be resolved by using Fab fragments rather than whole IgG molecules, filtering out the aggregates, or by diluting the primary antibody and incubating for longer times.
3. The tissue sections may be cut too thick — try thinner sections.
4. The microscope light needs to be adjusted to a higher setting.

If there is background staining in only the test sample(s) - i.e. not in the positive or negative controls, then the most likely cause is that the test sample(s) has been fixed and processed differently from the controls. Use of different tissue type between test sample(s) and controls may also produce this variance. Possibilities include:

• Overfixation of the test sample, resulting in the increased presence of hydrophobic groups, or increased crosslinking. Use of antigen retrieval procedures will amend this.
• A different fixative was used for the test sample(s) than for the control tissue. This difference should be avoided, or the procedures should be adjusted.
• The test sample(s) and the controls are of different tissue type. This should be avoided whenever possible.

If the test sample(s) and positive controls are clean, but the negative control shows background staining, it is likely that the negative control serum is at fault. It may be too concentrated, or contaminated with cross-reacting Ig components, naturally occurring antibodies, or bacterial growth. This can be corrected by using more dilute serum and incubating longer, trying to find a better match for the negative serum, or purifying the serum.