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

Figure 1. Flow cytometry experimental process overview.

Determine labeling protocols for targets of interest

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

Reagent titration

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

Biological controls

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.

  • Positive biological controls: A positive biological control consists of a sample that produces the desired experimental outcome or is known to express the antigen of interest. In some cases, this may involve treating the sample with an agent that is known to induce a desired response, or it may be an alternative tissue or cell type that is known to express the protein. Such biological positive controls help establish what “positive” looks like in a flow cytometry experiment.
  • Negative biological controls: A negative biological control may be cells that are left untreated or treated with a vehicle solution, may be tissues or cells known not to express the target of interest, or samples from known healthy subjects. Note that a positive and negative control suggest there are only two possible outcomes. However, biological systems are complex and therefore a gradation of outcomes is likely. In some cases, a control that sets a benchmark may be more appropriate and treated samples are simply compared to the benchmark. This is particularly relevant when the goal of the study is to determine whether there is a change in protein expression such as up or down regulation [4].

Technical 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.

  • Unstained cells: An unstained cell control is required for determining cellular autofluorescence, and for setting up forward and side scatter parameters. Autofluorescence differs in spectral range and intensity between cell types and is altered when cells age, are treated, activated or fixed. It is important to use an unstained cell control that reflects the condition of the cells and matches the experimental sample. Using an unstained cell control, unmixing algorithms can isolate and extract the spectral contribution of autofluorescence from the rest of the spectral signatures and improve signal resolution. It is recommended that an unstained cellular control be used for every cell type and treatment in the experiment.
  • Single stain controls: A set of quality single stain controls is required for all successful multiparameter flow cytometry experiments. In conventional flow cytometry, single stain controls provide the requisite information to calculate the amount of spillover of each fluorophore into non-primary detectors and correct for this spillover through the process of compensation. In spectral flow cytometry, the process of unmixing utilizes an algorithm to determine the contribution of each fluorophore using the spectral signature for each single stained sample [1]. These single stain controls can be generated from cells or antibody-capture beads by staining with only one antibody-conjugate at a time. Both cells and beads should be tested to determine which control provides the most accurate fluorophore spectral signature for unmixing. Having high quality single-stained controls is a prerequisite for the successful unmixing of any experiment. There are four rules that apply for single-stained controls, whether using cells or beads [4].
    • 1. Within one single stain control the positive and negative must have the same autofluorescence. For example, both populations should be beads of the same type or both populations should be cells of the same type. If the autofluorescence of the positive and negative do not match, errors will be introduced that will be revealed after unmixing.
    • 2. The positive signal from the single stain control must be as bright or brighter than the experimental sample.
    • 3. The single stain control must use the exact same fluorophore as the full experimental sample, and for tandem dyes it is recommended that the very same lot of tandem dye be used to avoid degradation and lot-to-lot variability.
    • 4. All single stained controls should be treated in the same manner as the experimental sample, including fixations, washings, and storage. As cells differ in the spectral range and intensity between cell types and is altered when cells age, are treated, activated, or fixed, use the same cell type and sample preparation for single stain controls as the experimental sample.
  • Secondary antibody controls: A secondary antibody control is needed with indirect staining procedures when the primary antibody is unconjugated. It helps determine if any non-specific binding is occurring due to the secondary antibody alone. Any signal observed above background would be attributed to non-specific staining.
  • Isotype controls: An isotype control is an antibody generated against an antigen not expressed on the cell type or sample being analyzed. For an isotype control to provide value, it needs to be matched to the species, immunoglobulin class, subclass, and light chain of the comparator antibody of interest. Further, it needs to be conjugated to the same fluorophore as the experimental antibody. If these strict parameters are followed, then isotype controls may be capable of demonstrating non-specific background contribution from several sources including Fc receptor binding and endogenous enzymes. However, even if most of the inherent properties of the isotype control and antibody of interest are aligned, variability in the concentration, degree of aggregation and fluorophore to antibody ratio can impact the true negative value represented by the isotype control, and isotype controls should not be the only control used to assess background or non-specific staining [5].
  • Fluorescence Minus One (FMO) controls (multicolor panel controls): FMO controls are a sample of cells stained with all the fluorescent reagents used in a multiparameter panel except one. With increasing numbers of fluorophores being used in flow cytometry, the correction of fluorescence spillover and the inherent error in measurement of the signals contribute to spreading. To distinguish true signal from spreading, especially for proteins with a continuous expression pattern, Fluorescence-Minus-One (FMO) controls can be used (Table 1).

Table 1. FMO controls.

FMO control Purpose
Cells stained with all fluorophore except for oneIdentifies spread into the omitted fluorophore signature
Gate positioning based on spreadCritical 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.

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References and suggested reading
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