This month's question from the Molecular Biology Forums (online at molecularbiology.forums.biotechniques.com) comes from the “Immunology and Immunochemistry Methods” section. Entries have been edited for concision and clarity. Mentions of specific products and manufacturers have been retained from the original posts, but do not represent endorsements by, or the opinions of, BioTechniques.Molecular Biology Techniques Q&A
How can I improve the sensitivity and specificity of my antibody stain? (Thread 29608)
Q I am using infected rabbit lung sections embedded in paraffin for immunohistochemistry (IHC). Staining results using several different markers show high background or low intensity of the specific signal.
I have varied the concentrations of Tween-20, tried different blocking reagents and concentrations, attempted antigen retrieval, and even used an Invitrogen kit for IHC, but nothing has resolved the problem. What else can I do to improve the signal?
A One detail you may have overlooked is how your slides were de-waxed and re-hydrated. For some techniques, clearing in xylene or Histoclear followed by re-hydration through a series of decreasing percentages of ethanol is sufficient to remove wax from the slides and re-hydrate the tissue. But some tissues require more aggressive de-waxing steps, since residual wax will affect access to the antigen as well as background staining.
Prior to de-waxing, try heating your slides in an oven held at a temperature above the melting point of the wax for 3-5 min; then place them in the clearing agent. You may also heat your xylene to help remove wax. After a couple of washes, re-hydrate as normal.
A You might also want to try heated Histoclear. In the past, I found that cold Histoclear did not fully remove even low melting point wax. We put a 250 mL Duran bottle with Histoclear in the same oven used to keep the wax molten. (Ours is 55°C, but if you are using wax with a 58°C melting point, heat the oven to 62°C.) It will take 20-30 min to reach the correct temperature. When it does, place your slide in it for 5-10 min. After that, proceed with the usual room temperature Histoclear washes prior to rehydration. This will remove the wax.
A Since IHC with multiple primary antibodies shows high background or no signal, I think your secondary antibody might be the problem. You might try a control slide without the primary antibody (secondary antibody only) and look at the background.
A I suggest an extensive titration to determine the best dilution for each antibody. Antibody solutions that are too concentrated or too diluted can lead to high background or poor signal.
A Did you include an endogenous horseradish peroxidase (HRP) or biotin-blocking step in your protocol? If these steps are omitted, you will see high background staining due to endogenous HRP and biotin present in the tissues.
Q What is the best way to block for biotin?
A You can buy biotin-blocking kits, but these tend to be expensive. I use egg whites or skim milk as described in “Technical Immunohistochemistry” by Rodney T. Miller. To block endogenous peroxidase, try H2O2 with and without methanol.
Are you using a biotinylated antibody or an avidin/biotin amplification kit? If you don't use either of these reagents, a biotin block won't be necessary. Depending on your antibody, you may need to use an antigen retrieval technique. Just remember that heating the slides can expose more endogenous peroxidase and biotin, so blocking will become even more important.
Watch your diaminobenzadine (DAB) closely during development. As soon as the signal is slightly lower than your desired intensity, wash and soak the slides for some time -- up to overnight -- in water. The stain will darken during this period. You want to minimize the time spent with the DAB solution since extra conversion of the substrate will cause deposits on the slide that will show up as background.
Also, remember that the antibody dilution listed on the fact sheet is only a suggestion; it must be optimized in your lab. I have used antibodies at the suggested concentration, only to see a very dirty slide. When I diluted the antibody considerably, I got a specific and clean signal.
A Rabbit lung does not contain high levels of endogenous biotin, so the avidin/biotin blocking solutions recommended may not be necessary. (It should be noted that liver, kidney, or tissues from birds, however, do contain high biotin levels.)
You can improve the amplification of the signal by using an unlabeled primary antibody instead of a biotinylated one. Pair that with a biotinylated secondary antibody. For example, try a biotinylated donkey anti-goat antibody and use donkey serum instead of goat serum for blocking. In that case, since the primary antibody was raised in a goat, the use of goat serum in your protocol would be problematic if you used a biotinylated donkey anti-goat secondary antibody.
Using a biotinylated primary antibody invites non-specific signal problems too. If the primary antibody binds non-specifically, the S-HRP reagent will do the same. If you must use a biotinylated primary antibody, use swine serum to block.
Also DAB can cause non-specific nuclear staining within only 5 min of incubation. (Nuclear enzymes develop the DAB; the details are included in the small print of the product literature.) NovaRed from Vector is a much better choice for HRP IHC protocols. It lasts longer on slides than DAB staining and it does not develop non-specific signals in the presence of nuclear enzymes.
Q I tried again, incorporating all of these suggestions. Unfortunately, it still is not working well. Is it time to invest in a monoclonal antibody?
Also, what is the reason for using porcine serum in the previously mentioned example?
A Sometimes the blocking reagents that work well in eliminating the background signal can't be easily explained. In my experience, using porcine serum as a blocker cleans up the slides very nicely for unknown reasons. This is true for electron microscopy applications as well.