There’s nothing worse than toiling away all week to repeat an experiment only to have your staining turn up blank. You’re left scratching your head trying to figure out why you’ve got nothing. Recently, Vector Laboratories’ very own Sherlock Holmes, Dr. Craig Pow, joined The Scientist University to discuss Improving IHC and IF Staining Results. You can watch the full on-demand webinar or keep reading for some tips on how to solve your staining mystery.
Absent staining can be particularly frustrating when it’s an assay you’ve done many times before. How can an experiment—that has become so routine you feel as if you could do it in your sleep—suddenly stop working altogether? “I would say in most cases of absence of staining, it’s probably due to something such as an omission of a reagent, possibly the wrong primary antibody, or maybe a misstep in the workflow,” reports Dr. Pow. For most researchers, repeating the experiment will resolve the issue. If that doesn’t work, your first impulse may be to simply rush out and buy new reagents, but Dr. Pow cautions that you may end up with lost time, lost money, and, confoundingly, the same result.
At this point, it’s time to take a deep breath, slow down, and remember that an absence of staining doesn’t mean you have to contemplate alternate careers. In this situation, your most powerful set of tools—the Watson to your Sherlock—is a complementary suite of both positive controls to work out the kinks in your assay and negative controls to check for nonspecific background staining. Although it can be tempting to skip right to the control you think you most need, keep in mind that multiple types of controls are meant to work together. A detective doesn’t set off with their case-cracking bag half-empty, and neither should you!
Positive controls allow you to examine your assay’s specificity and sensitivity, allowing you to test whether everything is working as expected. A good way to start is by choosing a control sample that expresses a moderate to high level of your target antigen. If you’re investigating multiple markers and are concerned about the expense of all those positive control specimens, it’s perfectly fine to use the same tissue for a variety of antigens. For example, Dr. Pow gives the example of choosing a tonsil specimen as a control for both CD20 and Ki67. You also have some flexibility in terms of the organ, tissue type and species: as long as the primary antibody that you plan to use recognizes the material in the positive control sample, it can differ from your experimental sample in these regards.
What is absolutely crucial is to treat the positive control in the same way as you treat your experimental sample—no playing favorites! If you have formalin fixed and paraffin embedded your experimental specimen, you must process your positive control that way too. Patience is key for troubleshooting a multiplex assay in which you are interested in staining for more than one antigen. Good detectives don’t take shortcuts; you’ll need to break down your assay and run positive controls for each of the target antigens.
So, you’ve done your positive control, you know you should be getting a stain and voilà! Now it’s time to run the positive control in parallel with your experimental stain, to determine whether the issue was indeed in your workflow. If even your faithful positive control stain hasn’t worked, however, you may need to troubleshoot your experimental protocol more deeply, by confirming that none of your reagents have expired or trying a different antibody diluent—this could mean changing the pH or composition. If all else fails, you might also consider a new primary antibody.
Villains are notorious for adopting disguises to lead the detective astray, but never fear—negative controls will help you unmask any imposter. These controls allow you to identify and eliminate false positive staining so that you can be certain your results aren’t leading you to an inaccurate conclusion. Inexperienced sleuths have been known to approach this set of controls by simply omitting the primary antibody, but Dr. Pow cautions that this is insufficient. “It doesn’t really address whether or not the primary antibody is contributing to any of the staining,” he explains. “Maybe it’s being bound inadvertently by something in the tissue prep, and this is where issues about reproducibility come into question.”
A better option would be to substitute a non-immune IgG under identical concentrations and conditions to determine whether your primary antibody may be binding nonspecifically to anything in your experimental specimen. For example, if your primary antibody is a mouse IgG, you would use a non-immune mouse IgG. If you do see non-specific staining with this negative control, don’t despair—since non-immune IgG is typically a pool of subclasses, choosing a non-immune IgG of the specific subclass of your primary antibody (for example IgG2a) may give you a cleaner result.
Are you able to do another negative control? Truly, you are following Sherlock Holmes’ directives: “A good detective knows that every task, every interaction, no matter how seemingly banal, has the potential to contain multitudes.” An excellent next step would be using an irrelevant antibody (i.e., one that you know won’t bind anything in your experimental preparation) raised in the same species, of the same isotype, and at the same concentration as your experimental primary antibody. Then, follow with the same detection reagents to determine whether the primary antibody isotype from that particular species is binding nonspecifically to your sample and contributing to your staining signal. Dr. Pow illustrates this with the case of an investigator working on CNS tissue who chooses a primary antibody against stratified, keratinized epithelium, which wouldn’t be found anywhere in their experimental specimen. Other possible options, depending on the study, could include staining in a knockout model or removing the antigen through enzyme activity or digestion. If these negative controls reveal an absence of staining, you can feel confident that your experimental results are real. Elementary, my dear Watson!
Inconsistent staining in your prep—in which you see a patchy or uneven intensity of the staining of your target antigen within the same assay—can strain any gumshoe’s powers of deduction. You may want to throw up your hands and blame Professor Moriarty for sabotaging your detection reagents. Instead, the most common culprits are issues with antigen retrieval methodology, improper fixation, incomplete paraffin removal, partial drying out of the sections during application, and uneven dispersion of reagents across the specimen. Antigen retrieval can be a common culprit. According to Dr. Pow, protocols for high temperature antigen retrieval differ widely between labs and can include a variety of high-tech instruments such as microwaves, vegetable steamers, and pressure cookers, making consistency a challenge. No matter how you go about it, take a careful look at your pH, temperature, and duration of sample immersion to ensure that each aspect of this process is optimized.
As Sherlock Holmes says, “to a great mind, nothing is little”. Zoom in on the details of your workflow to clear up your staining inconsistencies. If the trouble is improper fixation, which usually happens with immersion-fixed specimens, pay attention to the size and thickness of your tissue blocks and take care that specimens are fixed for an appropriate length of time. When incomplete paraffin removal is the offender, take an inventory of your reagents (toluenes, xylenes, etc.) to establish that each one is fresh. Partial drying out of sections can be resolved by processing the samples in smaller batches, and bubbles are the usual suspect in cases of uneven reagent dispersion across the sample.
As Holmes might say: “You know my methods. Apply them”. For more helpful tips and tricks to help you crack the case, be sure to check out our IHC Resource Guide and IF Resource Guide.