When planning an immunofluorescence (IF) experiment on fixed cells, it is important to consider which protocol will best suit your experimental setup. Usually you can check the literature to see what has been done previously (and successfully), but as protein expression and localization can vary depending on cell type and treatment, sometimes one has to optimize to get a clean, clear, and strong signal. There are numerous protocols available online, and it’s likely that more than one will work well for your cells, protein of interest, and reagents. The key is to choose a fixative that is best for both your protein of interest and antibody, to best permeabilize the cells to fit your experimental needs, and to optimize your signal-to-noise ratio using an adequate amount of primary antibody and reducing potential background.
1. Choose your fixative
Essentially, you fix to “end” your experiment and preserve the contents and structure of your cells. Fixatives can cause a change in the conformation of your protein, which may or may not block the antibody’s epitope. In that sense, it is best to consider the recommended fixative from the antibody supplier.
The most common fixative, formaldehyde/paraformaldehyde, fixes by crosslinking and thus stabilizing your protein of interest within the cellular environment. Fixation time can range from 10-15 minutes. Glutaraldehyde is another cross-linking fixative, but used less commonly for IF. After fixation with aldehydes, there is an optional quenching step to prevent potential nonspecific background fluorescence caused by the fixative.
Alcohols can also be used as fixatives. Methanol and ethanol are the most common and they work via dehydration of your sample. It’s generally recommended to use ice-cold alcohol and to fix the cells at -20°C, as lower temperatures are thought to facilitate the fixation process and better preserve structure. You can fix for 10 minutes, 5 minutes, or by dipping the coverslip in for 30 seconds—I’ve tried all, and each has worked well for given proteins or antibodies. Methanol both fixes the cell and permeabilizes the cell membranes, allowing for antibody penetration. Acetone is another common fixative and is similar to ethanol but also permeabilizes the cell, like methanol. Often an acetone:ethanol or an acetone:methanol mixture can be recommended.
Following formaldehyde/paraformaldehyde or ethanol fixation, one must permeabilize the cell membranes with a detergent, allowing for entry of the antibody to “find” the protein of interest. As noted, methanol and acetone both fix and permeabilize in one, and further treatment with detergent may damage the cell. Common detergents are Saponin, Triton and Tween. Saponin is thought to only permeabilize the cell membrane and not the nuclear membrane, thus allowing you to probe only the cytoplasmic/membranous pool of your protein of interest. Tween and Triton are identical, and Triton seems to be more commonly used. Triton and Tween concentrations and permeabilization times can vary and should be chosen based on your experimental needs.
Both shorter incubation times (such as 5 minutes) and less detergent (such as .1%) allow for minimal permeabilization and tend to promote selective recognition of membrane proteins. With higher detergent concentrations and longer incubation time (up to 20 minutes), nuclear and nucleolar protein availability are increased. You either want to maximize the detergent concentration or the permeabilization time, as, for example, using .5% Triton for 20 minutes may damage the cell. For experiments where the target protein is nuclear or nucleolar, subsequent steps will likely include Triton/Tween to ensure the antibody does reach its target.
The goal is to optimize the signal-to-noise ratio by blocking to reduce “background” by the antibody binding nonspecifically. Blocking can be done with either a protein solution, such as Bovine Serum Albumin (BSA), or with normal serum or serum of the animal that your secondary antibody is raised in. In addition, your solution can range from 1-10% solution in either PBS or in PBS including a detergent. Timing also can range from 10 minutes to 1 hour.
Following blocking, it’s time for primary antibody incubation, which is either for 1 hour at room temperature or overnight at 4°C. I recommend overnight if you can spare the time for a nicer signal. You have many options for solutions to dilute the antibody into, including PBS, PBS with BSA/serum, or also with detergent. Some antibodies will work with any of these options and others will be more particular. Antibody dilution may also have to be optimized, and it’s always best to start with the recommended dilution from the company. Sometimes you need a higher antibody concentration, but beware of nonspecific binding. Finally, add the secondary, followed by a nuclear counterstain, mount, and you’re ready to image.
Don’t forget to wash between steps!
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