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Herein we discuss spectral flow cytometry panel controls and sample preparation including labeling protocols, biological and technical controls. This is the next step in the spectral flow cytometry experimental process after panel design (Figure 1).
Explore spectral flow cytometry reagents Spectral flow cytometry experimental setup
The buffer system used for testing, panel preparation, and ultimately experimental sample acquisition should be identical to maintain consistency with any effect of the buffer on the sample and the antibodies under evaluation. The buffer systems and protocols selected to prepare and process samples and perform antibody staining steps is largely dependent on the cellular needs and antigens of interest and is selected to optimize detection and reduce non-specific binding. A common cause of non-specific binding is through the binding of the Fc portion of the antibody to Fc receptors expressed on many immune cell subsets. To circumvent this issue, reagents such as purified Immunoglobulin G (IgG), or serum, can be used prior to staining cells. Researchers can also use an Fc blocking reagent, which will bind to the Fc receptors. Lastly, cell blocking reagents specifically formulated to prevent non-specific binding observed with macrophages and monocytes are commonly used to improve resolution of cell populations.
The specific steps for labeling cells with antibodies are dependent on the experimental question. The high number of applications of flow cytometry necessitates an individualized strategy for labeling and should be a component of the experimental protocol. However, considerations that may be applicable across all types of labeling include optimal labeling conditions, order of labeling of reagents, and fixation. Of note, fixation can impact fluorescence intensity, as well as autofluorescence [1].
Learn more: Flow cytometry protocols handbook
A critical step before a pilot experiment is identifying the optimal staining concentration for each antibody-conjugate for the sample of interest through the process of titration [2]. Titration involves staining a sample with a series of antibody dilutions to determine the concentration that gives the best separation between positive signal and background. Antibody concentrations that are too low will under-label the cells of interest and will impair detection of cells expressing low levels of antigen, whereas antibody concentrations that are too high increase the background on negative cells. Any functional or viability reagent should also be titrated to determine their best concentration for use. It is good practice to titrate viability reagents first, and then titration of antibody-conjugates can be performed on viable cells using the optimal concentration of viability reagent [3]. In some cases, the antibody manufacturer provides a recommended concentration, and this can be used as a starting point for the titration. After titration, the stain index or separation index may be calculated relative to the dilution factor. This will reveal the saturating titer, the antibody concentration where additional antibody does not increase the signal intensity (Figure 2). Importantly, this process should be performed one antibody at a time and under conditions that are as close to the planned experimental system as possible (i.e., tissue and cell type, cell activation level, incubation time, buffer type, reaction volume, and temperature). Once antibodies have been evaluated individually, they will need to be assessed and further optimized for use in a combined panel.
Figure 2. Antibody titration and stain index. Human lymphocytes were stained with CD3-Alexa Fluor 647 at increasing dilutions and concatenated data is shown in (A) with the unstained sample represented first. (B) The stain index was calculated to determine optimal separation of the positive and negative cell populations with the optimal titer range identified where the stain index is highest.
Learn more: BioProbes 79: Best practices for multiparametric flow cytometry
Experimental controls, often referred to as biological controls, are necessary to validate the success of the experiment to provide confidence that any observed changes are due to the variable being tested. Biological controls are distinct from the technical controls, such as single stained samples that are necessary for appropriate instrument set up and interpretation of data. There are two types of biological controls.
Technical controls are used to help adjust detector settings, generate the unmixing matrix, and set boundaries for positive expression. Technical controls consist of unstained cells, single stain controls, secondary antibody controls, isotype controls, and Fluorescence Minus One (FMO) controls. Such technical control samples are essential in flow cytometry as they help ensure the integrity of the instrument and facilitate the accurate interpretation of results. It is recommended that these controls be processed alongside the biological samples.
FMO control | Purpose | |
---|---|---|
Cells stained with all fluorophore except for one | ![]() | Identifies spread into the omitted fluorophore signature |
Gate positioning based on spread | ![]() | Critical for targets with low or variable expression level |
FMO controls assist with gate placement by determining the cut-off between background signal due to spreading and positive populations (Figure 3).
Figure 3. Fluorescence-Minus-One (FMO) control. A dual-parameter plot displays the combination of IFNγ-APC and CD4 PE-Texas red antibody conjugates from a larger immunophenotyping panel show (A) unstained cells, (B) FMO control containing all antibody conjugates in the full panel except IFNγ-APC, and (C) full panel staining. The boundary for positive expression of IFNγ-APC as determined by the FMO control accounts for spread of the negative to allow proper identification of the IFNγ positive population.
FMO controls are superior to unstained cells and single stain controls since they consider the influence of all the other fluorophores used in the experiment that contribute to spreading. Because of the sheer number of controls that might be required for very large panels, it is important to keep in mind that FMO controls are most valuable for antigens in secondary and tertiary groups where expression is low, or when a continuum of expression can make it challenging to ascertain positive from negative.
There is a modified FMO control that can be used to investigate the impact certain additions or combinations of reagents may have on background, spread, and population resolution. It is called an FMx control where a sample of cells is stained with a subgroup of the fluorescence reagents, where x refers to the fluorophores that are omitted from the sample. This could also be set up where a large panel could be split into several smaller panels, to investigate the effects of combining multiple antibodies to reveal potential unwanted interactions, or to determine if a specific staining sequence is affecting signal resolution.