Flow Cytometry Sample Prep: Best Practices and Tips for Success

Generating high-quality samples for flow cytometry is critical to produce accurate results. We recently spoke with Richard Cuthbert, Ph.D., Global Product Manager, Flow and Antibody Business, Bio-Rad Laboratories, and Maria Jaimes, Vice President, Scientific Commercialization, Cytek Biosciences, who helped provide answers to some of the most common flow cytometry sample prep questions.

Biocompare: What are the main characteristics of a ‘good’ flow cytometry sample?

Richard Cuthbert: “First and foremost, your sample should be a single-cell suspension, with a low number of doublets and aggregates. Failure here could potentially block your instrument, so this factor has wider consequences than ensuring data quality. Other important sample characteristics include cell health and viability, cell concentration, optimized staining conditions, and the choice of an appropriate staining buffer.”

Maria Jaimes: “The cell concentration should be high enough that the acquisition rate is not too slow, but not so high that it exceeds the acquisition limit of the flow cytometer. You should also aim to minimize the presence of debris or ‘contaminating cells’, such as red blood cells when studying leukocyte populations. Additionally, researchers should note that some sample characteristics are interconnected; for example, low viability will lead to clumping and excess debris.”

Biocompare: How can you check the cell concentration and viability of your flow cytometry samples?

Maria Jaimes: “One option is to perform a manual count using a hemocytometer and the Trypan Blue exclusion method. This is inexpensive and does not require specialized equipment, although it can be slow and might show variation from operator to operator. Another approach is to use an automated cell counter, which offers the advantages of high reproducibility, efficiency, and low cost. Alternatively, you can use flow cytometry, which either requires counting beads or a flow cytometer with volumetric count capabilities, such as the Cytek Aurora™ flow cytometer. Although this strategy is more expensive, it offers a more in-depth characterization of the sample, including the absolute counts of various cell types.”

Biocompare: What is the recommended staining buffer for flow cytometry?

Richard Cuthbert: “Cells are typically stained in PBS supplemented with a low concentration of BSA or serum to maintain cell viability and reduce non‑specific antibody binding. Depending on the sample type, additional components may be included; for example, EDTA is commonly added when working with adherent or sticky cells to minimize cell–cell adhesion and reduce clumping and doublets. When the antibody staining panel includes multiple polymer‑based fluorophores, such as Brilliant Violet™ dyes, a specialized staining buffer is recommended to prevent dye–dye interactions and ensure accurate fluorescence detection.”

Maria Jaimes: “When detecting surface markers, the addition of 0.01–0.1% sodium azide to the staining buffer is advised as it stops cells from actively internalizing (endocytosing) the antibody-antigen complex after binding and prevents ‘capping’ (the clustering of surface receptors to one pole of the cell) and the subsequent shedding of these complexes into the media.”

Biocompare: When using flow cytometry to study adherent cells, which detaching agent should you use?

Richard Cuthbert: “When working with adherent cells, your detaching agent should be chosen based on where your markers of interest are located. If you’re measuring cell surface markers, enzyme‑free options like EDTA are recommended, since enzymes like trypsin can damage or cleave epitopes, leading to reduced signal or false negatives. If your markers are intracellular or nuclear, the detachment method is less critical.”

Biocompare: What are some key considerations when using flow cytometry to analyze whole blood?

Maria Jaimes: “The type of anticoagulant used in the collection tubes is an important factor. While EDTA is the preferred anticoagulant for immunophenotyping, heparin is a better choice for functional assays and calcium-dependent processes, since EDTA is a strong calcium chelator and can inhibit cellular function. The time elapsed between blood collection and staining, known as ‘age of blood’, should also be considered.”

Richard Cuthbert: “If you use heparin as an anticoagulant, be aware that it can activate neutrophils, interfere with downstream applications performed on sorted cells (e.g., PCR reactions), and is not as good as EDTA at preserving cells if you intend to store your samples prior to analysis. For flow cytometry experiments in which only white blood cells are of interest, it is beneficial to remove erythrocytes using a red cell lysis step. Erythrocytes make up the majority of cells in blood, so their removal ensures that only data from the cells of interest are acquired, improving identification of leukocyte populations and enabling faster acquisition times.”

Biocompare: Which dissociation method is best when analyzing solid tissues with flow cytometry?

Maria Jaimes: “The choice of dissociation method depends not only on the tissue of interest, but also on the cell type that is going to be analyzed. For example, while mechanical disaggregation works nicely in spleen to analyze T or B cells, enzymatic digestion is often required when it comes to macrophages. Researchers are advised to search the literature for guidance on their specific sample type.”

Helpful resources include Best Practices for Preparing a Single Cell Suspension from Solid Tissues for Flow Cytometry and the Worthington Tissue Dissociation Guide.

Biocompare: When should you use a cell strainer during flow cytometry sample prep?

Richard Cuthbert: “The most important time to use a cell strainer is immediately before you load your sample into the flow cytometer. However, don’t be afraid to do an additional filtration at any point in the preparation process. If you see your cells starting to aggregate, filter them straight away—don’t centrifuge before you have had a chance to filter them otherwise that small aggregate will turn into a big one very quickly and you will lose a lot of your sample. Selecting the right pore size for your target cells is important, and you should always rinse the filter thoroughly to minimize cell loss.”

Biocompare: Which fixative is recommended for flow cytometry?

Richard Cuthbert: “The choice of fixative depends on the target that is being detected. Alcohol fixatives precipitate and denature proteins, as well as permeabilizing the cell membrane, making them ideal for cell-cycle assays and detection of phosphorylated proteins. However, alcohol fixatives will also denature protein-based fluorophores, such as PE, APC, PerCP, and tandems of these—something to bear in mind if you are doing surface staining followed by intracellular staining. Aldehyde fixatives, commonly 1–4% paraformaldehyde, are gentler solutions that cross-link proteins and preserve surface marker structure, meaning they are often the preferred choice. To detect intracellular proteins in aldehyde fixed cells, a subsequent permeabilization step is necessary.”

Biocompare: How should flow cytometry samples be blocked?

Richard Cuthbert: “Including an Fc block prior to staining is an essential step to avoid non-specific binding of antibodies. Commercially available blocking solutions are easy and convenient, although a cost-effective alternative is to use serum matched to the species the cells originate from. But be careful, mouse serum alone is not an effective block for mouse cells. Instead use anti-mouse CD16/CD32 as your blocking agent as this more effectively blocks Fc binding.”

Maria Jaimes: “In our laboratory, for mouse samples, the standard procedure is to incubate cells with anti-mouse CD16/CD32 for 15–30 minutes at 4°C or on ice, prior to antibody addition. For human samples, we consistently block the binding of certain fluorochromes (e.g., PE-Cy7, APC-Cy7, and PE-Cy5) to monocytes and macrophages by adding True-Stain Monocyte Blocker™ or equivalent reagents at the same time as the antibodies.”

Ten tips for successful immunostaining in flow cytometry

• Know the biology of your sample; for example, some antigens are rapidly shed from the membrane (CD62L) or internalized (CD115) at room temperature, which can be avoided by keeping samples at 4°C.
• Select antibodies that are validated for flow cytometry and make sure to identify clones that are suitable for your experiment; for example, CD3 clones such as OKT3, SK3, and UCHT1 are all validated for flow, but some will activate cells and they all bind different epitopes.
• Titrate antibodies to maximize the signal-to-noise ratio.
• Optimize every step of the staining protocol; incubation time and temperature can have a significant impact on the quality of the staining.
• Choose fluorophores that are compatible with your flow cytometer’s lasers and detectors, and that are spectrally distinct in order to minimize spectral overlap.
• Take steps to minimize autofluorescence, such as considering methanol fixation instead of aldehyde fixation, lysing red blood cells, and selecting fluorophores with longer emission wavelengths (autofluorescence is most prominent in the green channel).
• Always include a viability stain in your panel.
• Consider overnight staining protocols; these have recently been gaining more traction as they present advantages for high-parameter panels, such as lower antibody titers and higher resolution.
• Include appropriate controls to show that the assay is performing as expected; examples include biological controls, unstained controls, and fluorescence-minus-one (FMO) controls.
• Be sure to optimize the protocol for your particular model system; never completely rely on someone else’s protocol, although this is usually a great place to start.