All of our antibodies undergo strict QC analysis by western blotting. We purify each lot with an affinity column, and we use the same protocol and conditions throughout the analysis. Also, we only release our antibodies after we obtain satisfactory results in comparison with the previous lot. If you do not obtain a signal, there could be a fundamental issue with your antibody or technique.
• You may not have used enough primary or secondary antibody. Please follow the recommended antibody dilutions or test different antibody dilutions to determine the optimal antibody concentrations.
• If you’re using a fluorescent detection system, you may not have kept your conjugated primary or secondary antibody in the dark. Ensure that these antibodies are not exposed to light whenever possible.
• There’s the possibility that your primary and secondary antibodies do not work together. Ensure that the secondary antibody was raised against the animal that the primary antibody was raised in.
• The NaN3, present in the antibody solution, can sometimes cause problems.
• Your fixation method could be damaging the epitope and preventing the primary antibody from recognizing it. Either reduce the fixation time or try an antigen retrieval method.
• Your protein of interest may not be present or may be present at very low levels, in the sample that you’re testing. Alternatively, the primary antibody may not recognize the protein of interest in the species that you’re testing. If you are certain the protein is expressed, you can try an enrichment step to improve the signal. You should also check the antibody datasheet to ensure that it cross-reacts with the species that you’re testing.
High background levels may be due to a sub-optimal primary antibody concentration, an insufficient blocking step, or there could be non-specific binding between the antibodies and the blocking reagent.
• Titrate the primary antibody to obtain the optimal concentration. Check the datasheet for the optimal dilution, but we recommend 1:100 as a starting point for most primary antibodies. For antibodies directly conjugated to a fluorophore, we suggest a 1:50-1:60 dilution.
• Try adding 0.1%–0.5% of Tween-20 to the antibody solution and washing buffer since this strengthens the signal and reduces non-specific staining. Tween-20 is more gentle than other detergents such as Triton X-100, which can damage the membrane during permeabilization.
• In some cases, the biotin-avidin amplification can cause high background and there is endogenous avidin binding ability in several tissues. Thus, when we use a biotin-conjugated secondary antibody and then add extravidin or streptavidin conjugated to peroxidase or a fluorophore, it may be bound not only to the biotin from the secondary antibody but also to binding sites in the tissue. To overcome this problem, saturate endogenous binding sites before exposure to the primary antibody by incubation of the tissue sections in a solution containing non-conjugated streptavidin. We use a kit from Vector Laboratories (US): kit SP-2002.
o These steps are for section after antigen retrieval treatments (if required) and after 2 x 5-minute rinses in IHC-PBS.
o Pre-saturate endogenous avidin binding ability: Add 4 drops from the “streptavidin” bottle to 5 ml of IHC-PBS.
o Incubate sections in this solution 30 minutes at room temperature.
o Rinse 2 x 5 minutes in IHC-PBS.
o Saturate residual biotin binding potential: Add 4 drops from the “biotin” bottle to 5 ml of PBS. Incubate sections in this solution 30 min at room temperature.
o Rinse 2 x 5 minutes in IHC-PBS.
o Start IHC and transfer sections to the primary antibody.
• Your sample incubation temperature may be too high. Incubate the tissue sections at 4°C.
• Excessive fixation times can affect the epitope and cause high background levels. Try a fixation protocol with a lower exposure time, lower temperature, and/or reduced concentration of fixative. In general, we incubate the sample for 24–72 hours in 4% paraformaldehyde (PFA). When the samples consist of bloody organs, such as spleen and liver, we recommend you replace the PFA after 24 hours.
• If you grew your cells on a coated chamber slide, the antibody may stick to the surface. In this case, try using another coating solution (e.g., polyethylene glycol, polyethyleneimine, etc.) since it may affect the background levels.
This is due to endogenous peroxidase activity. Try endogenous peroxidase quenching with hydrogen peroxide. For more details, check out our IHC protocol for paraffin embedded sections https://www.alomone.com/info/immunohistochemistry-ihc-protocols-for-paraffin-embedded-sections
This is likely due to damaged tissue from sectioning or other similar procedures.
• Aggregates may form in the antibody solution following reconstitution and especially after thawing of the reconstituted antibody solution. Thus, we recommend centrifuging all of the antibody preparations before use (10000 x g for 5 min).
• Sectioning with a dull blade can cause folding or air bubbles. Moreover, cutting sections too thick can make them difficult to resolve. Ensure your section equipment is well-maintained, and your protocol is optimized.
We find that floating sections are stained more effectively compared to sections adhered to slides. When sections are adhered to slides, there are several potential problems since the access and penetration of the antibody beyond the surface of the sections is limited. You can overcome this to some degree by increasing the Triton X-100 content in the antibody solution. However, we don’t recommend this in many cases where the antibody targets an extracellular epitope because Triton X-100 is a relatively harsh detergent and too much can deteriorate the antigens.
If you’re looking for a protocol, take a look at our detailed IHC/ICC/IF Protocols.