Optimizing flow cytometric staining
Now that we learned how to set up the instrument in the previous post, it is important to further optimize the staining in order to produce the best results. Using the correct antibodies and fluorophores is important, as well as controlling for background staining.
In order for the vertebrate immune system to effectively fight the wide range of pathogens encountered, lymphocytes play a vital role in producing receptors that can recognize these foreign antigens. The antigen-recognizing molecules produced by B cells are known as immunoglobulins (Ig). Membrane-bound immunoglobulins serve as the B-Cell receptor (BCR), while secreted immunoglobulins are known as antibodies. These antibodies are able to bind to foreign pathogens, eliciting a specific immune response.
Immunoglobulins are divided into 5 major classes: immunoglobulin G (IgG), IgA, IgM, IgD, and IgE. For the purposes of flow cytometry we are going to focus on IgG only. Each IgG molecule is composed of two identical heavy chains (H) and two identical light chains (L) (Figure 1a). The H chains determine the class and subclass, while the L chains are either kappa (κ) of lambda (λ). The IgG molecule can be further divided into the variable (V) and constant (C) domains (Figure 1b). The variable domains of the light and heavy chains together for the antigen-binding site. These variable domains are what enable diverse sites to be created to bind different antigens.
Figure 1. The structure of an immunoglobulin IgG molecule. (a) IgG molecule indicating light and heavy chains (b) The light chains can be either kappa or lambda chains
For the purposes of immunochemistry assays such as flow cytometry, ELISA, and immunohistochemistry, antibodies are produced by injecting immunogen (antigen) into laboratory animals (e.g. rabbits or mice) which then produce IgM and then IgG antibodies specific to that immunogen. The antibodies produced are monoclonal or polyclonal. Monoclonal antibodies are targeted to a single epitope, while polyclonal antibodies are able to recognize and bind to different epitopes or parts of the antigen. For flow cytometry experiments, these antibodies are conjugated to fluorophores, which when excited by the laser emit light at certain wavelengths as discussed in the previous post.
An isotype control is an antibody that cannot bind specifically to the antigen of interest, but matches the other properties of the primary antibody of choice. Fluorochrome-conjugated isotype controls are used to account for the non-specific binding of the fluorochrome-conjugated antibody.
Non-specific binding or background staining can be caused by a number of factors. Fc receptors are found on the surface of many cells, and some antibodies, such as mouse IgG2a isotype, can bind to these Fc receptors non-specifically. Cellular autofluorescence caused by mitochondria, may lead to background staining. Non-specific antibody interactions can also occur when antibodies bind to non-target proteins, lipids and other molecules. Agilent has a range of both mouse and rabbit isotype controls that can be used to eliminate non-specific staining (See: https://www.agilent.com/en/product/clinical-flow-cytometry/reagents-for-clinical-flow-cytometry/clinical-isotype-controls).
Remove dead cells
Cell death in the sample could be caused by factors including time taken to process the sample, processing conditions (e.g. harsh handling or enzymatic treatment), or as a desired experimental outcome (e.g. cell death analysis/apoptosis assay). When cells dye, their cell membranes become damaged allowing for exposure of intracellular antigens that are not normally expressed on the cell surface. This can cause non-specific antibody binding. Dead cells can be detected using a fluorescent DNA viability dye such as propidium iodide, which is only able to enter cells with disrupted membranes, and binds to the exposed DNA. This can then be detected on the flow cytometer. These dead cells can then be gated out of the analysis, or it can help in decision making in whether or not to proceed further with the experiment (e.g. if a lot of cells are dead).
Antibody titration allows you to further optimize your flow cytometry experiment. This is essential to ensuring that the appropriate amount of antibody is used for each experiment. Too much antibody can increase non-specific binding, while too little antibody could mean that you miss detecting the target antigen. Antibody titration allows you to see which the amount of antibody that allows you to distinguish between different cell populations adequately and determine the optimal level of expression.
Antibodies usually come with a recommended concentration or volume range to use, and this should be the starting point of any titration. For example, if the range given for a cell count of 1 million cells/ml is 5-30μl, you should set up a series of tubes (same cell count in each) as follows:
Acquire the data and plot the volume versus the mean fluorescence intensity (MFI) as shown in Figure 2. As the volume of antibody increases, the MFI increases until it reaches a saturation point and the curve flattens out (Figure 2). This is the ideal volume to use for staining, as it ensures that all the molecules that can be stained are. Using a volume of antibody greater than this will have no added value, and will be a waste of antibody and money.
- Dako – Flow Cytometry Educational Guide, 2nd Edition
- Murphy, K., Travers, P., Walport, M. (2008). Janeway’s Immunobiology – 7th edition. Garland Science. p111-113
- https://www.enzolifesciences.com/science-center/technotes/2015/february/10-tips-for-successful-flow-cytometry/ (accessed 3 August 2020)
- https://www.nlsinsight.com/post/beginners-guide-to-choosing-isotype-controls (accessed 6 October 2020)
- https://cancer.wisc.edu/research/wp-content/uploads/2017/03/Flow_TechNotes_Antibody-Titrations_20170918.pdf (Accessed 6 October 2020)