Immunophenotyping by Flow Cytometry

Immunophenotyping is the most common application of flow cytometry. In its simplest form, it uses fluorophore-labeled antibodies for detecting cell surface markers. This editorial looks at the different protocol steps involved in these types of experiments and shares tips for optimization.

Immunophenotyping principles

The types of samples that are used for immunophenotyping include whole blood, tissue biopsy material, and peripheral blood mononuclear cells (PBMCs), as well as induced pluripotent stem cells (iPSCs), organoids, and spheroids. These are initially processed to form a single-cell suspension before being stained with fluorophore-labeled antibodies for cellular markers and analyzed by flow cytometry. By identifying and enumerating specific cell types within different samples, researchers can better understand the cellular response to infection, illness, or vaccination, and can ultimately use this information to help prevent transmission or the development of severe disease.

Types of markers

Immunophenotyping experiments are often based on the expression of cell surface markers, of which the cluster of differentiation (CD) antigens are the most well-known. To date, over 350 different CD markers have been characterized by the Human Leukocyte Differentiation Antigens (HLDA) Workshops—a series of workshops initiated in 1982 to ensure consistency when defining monoclonal antibodies for specific cellular antigens. Reported CD antigens include markers for cell lineage such as the T cell markers (CD3, CD4, CD8), B cell markers (CD19, CD20), monocyte markers (CD14, CD11b), and natural killer (NK) cell markers (CD56, CD161), as well as markers for activation (CD69, CD25, CD62L) and memory (CD45RO, CD27). Immunophenotyping may also involve detecting intracellular markers such as cytokines (e.g., IFN-γ, TNF-α, and IL-2), proliferation markers (e.g., Ki67, CFSE), and antigen specific markers (e.g., major histocompatibility complex molecules), although the exact combination of markers that is used will depend on the experimental questions being asked.¹

Key protocol steps for extracellular staining

When staining for extracellular markers, a typical immunophenotyping protocol involves cell counting and resuspension, followed by blocking, antibody staining, and washing. “For the most part, no fixation is needed when detecting only cell surface markers,” reports Nina Senutovitch, Senior Scientist at Sartorius. “However, there are cases when you might want to fix, such as if you plan to perform analysis on a later day or if you need to inactivate potentially infectious samples. When fixation is required, you should always validate the fixative and antibodies you plan to use. This is because fixation and further wash steps may cause cell loss and affect the final event acquisition of your assay, and some antibodies and fixatives may not be compatible. Generally, 1% PFA tends to better preserve epitope conformation compared to methanol fixation.”

• Cell counting and resuspension

When working with live cells, it is important to be as gentle as possible,” cautions Amber Miller, Ph.D., Flow Cytometry Scientist at Fortis Life Sciences. “This means flick mixing the cells or dragging the tubes along an Eppendorf rack rather than vortexing the samples. Pipetting up and down will also help with resuspension and evenly distributing cells in the system.” A density of 1 million cells/mL is often recommended when resuspending cells, although this should always be optimized. “Using too many cells risks missing events, while using too few cells can extend run times,” explains Miller. She also suggests using resuspension buffers that contain DNase and EDTA and are free of Ca²⁺/Mg²⁺ ions to help prevent clumping.

• Blocking

Blocking is essential to prevent non-specific antibody binding and can also help to stabilize cellular morphology. Common blocking agents include bovine serum albumin (BSA), various commercial protein-free formulations, and normal animal serums. “When performing indirect detection, many researchers block with serum corresponding to the host species of the secondary antibody,” notes Miller. “Often, this is 5% normal goat serum. At Fortis Life Sciences, I traditionally block with 1 µg of IgG Fc fragment per 10⁶ cells, which should bind to any surface proteins that might otherwise bind the Fc region of the primary antibodies being used for immunophenotyping and lead to false positive results.”

• Antibody selection

There are several factors to consider when selecting antibodies for immunophenotyping studies. Paramount among these is identifying antibodies that are validated for flow cytometry, although Janice Yasuda, Reagents and Consumables Product Manager at Sartorius, stresses the importance of performing validation in-house to optimize for your cells and instrument. “Because every immunophenotyping experiment is different, optimization is essential to obtain accurate results,” she says. “Data quality can be further enhanced by using recombinant antibodies to ensure optimal batch-to-batch reproducibility or Fc silent antibodies to reduce background staining from the interaction of antibodies with Fc receptors. And don’t be afraid to perform indirect staining—through antibody engineering, you can now choose to switch the species of primary antibodies for increased diversity in your panel.”

• Washing

Washing serves to eliminate cellular debris, residual media components, and unbound antibody reagents, and must be optimized like any other protocol step. “Optimization should include determining the composition of the wash buffer, as well as the number, duration, and volume of wash steps required,” says Miller. “It is also important to define how the washes will be performed. For example, I like to include BSA and EDTA in my wash buffer to block any non-specific binding and reduce clumping, and I simply dump the supernatant off the samples and tap the plate gently to a paper towel after centrifugation and between washes to avoid damaging the cells.” Depending on the vessel being used and the number of samples being processed, it may be worth using a plate washer instead of manually washing to both save time and minimize cell loss.

• Cell viability

Dead cells can be problematic for immunophenotyping experiments. Not only can they cause cells to clump due to the release of DNA, but they also exhibit higher levels of autofluorescence compared to live cells and can bind non-specifically to antibodies. “Including a viability marker in your panel allows you to remove dead cells from your analysis,” says Yasuda. “One group of viability dyes are the nucleic acid binding dyes, such as propidium iodide, DAPI, and 7-aminoactinomycin D (7-AAD), which are excluded from live cells by the intact plasma membrane. These all have different emission maxima, which means you can be flexible when choosing a suitable dye for your immunophenotyping panel.”

Whatever the aim of your immunophenotyping experiment, companies such as Fortis Life Sciences and Sartorius are happy to help. And don’t forget that Biocompare now offers a Flow Cytometry Panel Builder tool to streamline reagent selection and panel design.

Reference

McKinnon KM. Flow Cytometry: An Overview. Curr Protoc Immunol. 2018 Feb 21;120:5.1.1-5.1.11. doi: 10.1002/cpim.40.