Flow cytometry protocol

This protocol provides a step-by-step guide for performing flow cytometry on both intracellular and extracellular targets. It outlines essential procedures including sample preparation, live/dead cell discrimination, fixation, permeabilization, and antibody staining. Designed for researchers working with cell suspensions, the protocol ensures optimal detection of surface and internal markers using fluorescently labeled antibodies. It also includes tips for minimizing cell damage and maximizing signal specificity. Whether you are analyzing immune cell subsets or intracellular signaling proteins, this protocol supports high-quality, reproducible results. Ideal for both novice and experienced users, it integrates best practices and troubleshooting advice to streamline your flow cytometry workflow.

Introduction

Flow cytometry is a powerful technique for analyzing the physical and chemical characteristics of cells. This protocol focuses on staining both intracellular and extracellular targets, enabling comprehensive cellular profiling. By using fluorescent antibodies and viability dyes, researchers can distinguish live from dead cells and identify specific protein markers, including cell surface markers that are key targets for immunophenotyping. The protocol is compatible with various sample types. Identification and characterization of each cell type are achieved by assessing cell surface antigens and biological functions, which are critical for distinguishing cell populations in flow cytometry. This resource is essential for immunology, oncology, and cell biology applications where multiparametric analysis is often required.

Background and principles

Flow cytometry relies on the principle of hydrodynamic focusing and laser-based detection to analyze individual cells in suspension, and is a form of cytometric analysis. Fluorescently labeled antibodies bind to specific antigens on or within cells, allowing for the identification of distinct populations. Extracellular staining targets cell surface proteins, which are readily accessible to antibodies without the need for permeabilization, while intracellular staining requires fixation and permeabilization to access internal components. Viability dyes help exclude dead cells, which can bind antibodies nonspecifically. Gentle handling is emphasized to preserve cell integrity and optimize staining efficiency. By combining multiple fluorophores, researchers can simultaneously assess several markers, making flow cytometric analysis of suspended cells a cornerstone of modern cellular analysis. Flow cytometry analysis involves interpreting the data obtained from stained cell populations to identify and characterize different cell subsets based on their marker expression.

Stage 1 - Sample preparation

First, harvest your cells or tissue and prepare a single-cell suspension. Then, transfer the single-cell suspension into 96-well plates, test tubes, or polystyrene round-bottom tubes, depending on the number of cells and volumes being used.

Materials required

• Cell suspension
• Polystyrene round bottom 12 x 75 mm² Falcon tubes/96-well plate (or any other container compatible with your centrifuge)
• Suspension/washing buffer (PBS, 5-10% fetal calf serum (FCS))
• Optional - red blood cell lysis buffer (for example ab204733)

Steps

Harvest and wash cells according to the manufacturer's guidance.

• For blood samples, we suggest incubating samples with RBC lysis buffer before proceeding.

Determine the total cell number and check cell viability.

• In general, viability should be 90-95%.

Spin down and resuspend cell samples in an ice-cold suspension buffer.

• Centrifuge at ~200 x g for 5 minutes at 4°C.
• Recommended cell concentration for suspension: 0.5–1 x 10⁶ cells/mL.

Proceed to stain with a viability dye.

Stage 2 - Live/dead staining with a viability dye

As dead cells are prone to bind to antibodies non-specifically, we must exclude those cells from the analysis. Using viability dyes allows us to distinguish between live and dead cells and exclude the dead ones during data acquisition and analysis.

DNA binding dyes, such as 7-AAD, DAPI, and TOPRO3, are often used as viability dyes for live/dead staining, as they cannot penetrate the cell membrane of live cells. The compromised cell membrane found in dead cells allows these dyes access to DNA, to which they bind and emit fluorescence.

However, these dyes cannot be used for live/dead staining with fixed cells, where cell membranes would be compromised in all cells. In this case, we must use amine-reactive fixable cell viability dyes.

Materials required

• Viability dye

  • Examples of non-fixable dyes: 7-AAD (ab228563), DRAQ-5 (ab108410), DRAQ-7 (ab109202) or DAPI (ab228549)
  • Examples of fixable dyes: ab176738, ab176741, ab176739, and ab176744

• Suspension/wash buffer (for example, PBS, 5-10% fetal cell serum (FCS))

Steps

Stain cells with a viability dye according to the manufacturer's protocol.

• Incubate cells with dye in the dark at 4°C, according to the manufacturer's instructions.

Wash cells two times with wash buffer.

• Spin cells down (200 x g, 5 minutes, 4°C), remove the supernatant, and resuspend the pellet after each wash.

Proceed to blocking when detecting extracellular targets or to fixation and permeabilization for intracellular targets.

Stage 3 - Fixation and permeabilization (optional - only for intracellular staining)

When staining intracellular targets, we must proceed with additional fixation and permeabilization steps. Fixation is required to preserve the structure of intracellular proteins. Permeabilization disrupts the cell membrane, allowing antibodies to enter the cell and stain intracellular targets.

When staining extracellular targets, we’ll proceed immediately to the blocking step. When analyzing intra and extracellular targets together, we'll perform cell surface staining (ie, Stage 5) before fixation.

Helpful tips for choosing suitable fixation and permeabilization methods for intracellular staining:

• Antigens close to the plasma membrane and soluble cytoplasmic antigens will require mild cell permeabilization without fixation.
• Cytoskeletal, viral, and some enzyme antigens usually give optimal results when fixed with a high concentration of acetone, alcohol, or formaldehyde.
• Antigens in cytoplasmic organelles and granules will require a fixation and permeabilization method, depending on the antigen.
• The epitope needs to remain accessible.

Materials required

• Cell suspension
• Suspension buffer (PBS, 5-10% FCS)
• Fixative (for example, 1-4% paraformaldehyde, 90% methanol, or acetone)
• Permeabilization solution (for example, Triton X-100, NP-40, or Saponin)
• Alternatively, you can also use our fixation and permeabilization kit for flow cytometry (ab185917), which is suitable for most sample types.

Steps

Fix the cells in your chosen fixative.

• Spin down cells to a pellet (200 x g, 5 minutes, 4°C), discard the supernatant, and resuspend the pellet in fixative.
• Incubate cells with the fixative as indicated below.

Wash the cells twice with suspension buffer.

• Spin down cells to a pellet (200 x g, 5 minutes, 4°C), discard the supernatant, and resuspend in wash buffer.

Permeabilize cells by incubating them with a suitable detergent.

• Spin down cells to a pellet (200 x g, 5 minutes, 4°C), discard the supernatant, and resuspend the pellet in a detergent solution.
• Incubate cells in the detergent for 10-15 minutes at room temperature.

Wash the cells twice with the suspension buffer.

• Spin down cells to a pellet (200 x g, 5 minutes, 4°C), discard the supernatant, and resuspend the pellet in the wash buffer.

Stage 4 - Blocking

Blocking proteins and Fc domains is essential to prevent the non-specific binding of antibodies to cells.

Materials required

• The choice of materials will depend on the type of cells analyzed and, if applicable, the secondary antibody used.
• FcR Blocking buffer (for example, 2-10% goat serum, human IgG, or mouse anti-CD16/CD32)
• Suspension buffer (PBS, 5-10% FCS) 5-10%

Steps

Block Fc receptors with a blocking buffer.

• Spin down cells to a pellet (200 x g, 5 minutes, 4°C), discard the supernatant, and resuspend the pellet in the blocking buffer.
• Incubate cells with one of the buffers below for 30-60 minutes in the dark at 4°C.

Blocking buffers:

• 2-10% goat serum
• Human IgG
• Mouse anti-CD16/CD32

Wash cells twice with the wash buffer.

• Spin cells down (200 x g, 5 minutes, 4°C), remove the supernatant and resuspend the pellet after each wash.

Proceed to antibody incubation.

Stage 5 - Antibody incubation

We’re now ready to stain cells with fluorophore-conjugated antibodies for indirect or direct detection in the flow cytometer.

The following procedures can also be repeated and adapted for multicolor flow cytometry, in which multiple sets of fluorophore-conjugated antibodies are used against different targets. You should minimize any overlap in the fluorophores’ emission spectra when using multiple sets of antibodies.

Materials required

• Conjugated primary antibody
• Samples (cell suspension at 0.5 - 1 x 10⁶  cells/mL)
• Suspension buffer (PBS, 5-10% FCS)

Steps

Dilute the conjugated primary antibody in the suspension buffer.

• Suggested dilutions for each antibody will often be provided on the datasheet.

Incubate cells in the pre-diluted primary antibody.

• Spin down cells to a pellet (200 x g, 5 minutes, 4°C), discard the supernatant, and resuspend the cells in the primary antibody solution.
• Incubate in the dark for 20-30 minutes at 4°C.

Wash the cells two times with the suspension buffer.

• Spin cells down (200 x g, 5 minutes, 4°C), remove the supernatant, and resuspend the pellet after each wash.

Proceed to detection in the flow cytometer as soon as possible.

• If the cells are not analyzed in the flow cytometer immediately after antibody staining (within 1 hour) and were not stained earlier, stained cells can be fixed at this step (1-4% PFA, 20 minutes, 4°C). Fixation helps preserve the cells for several days, stabilizing the light scatter and inactivating most biohazardous agents. The controls will require fixation using the same procedure. Note that fixation will kill cells.
• After fixation, wash cells three times and store the cell suspension in the suspension buffer.
• Follow the manufacturer's instructions.

Stage 6 - Detection and data analysis

After antibody incubation, we can run our experiment in the flow cytometer. The procedure depends highly on the equipment used, so always refer to the manufacturer in the first instance. For a more detailed discussion of fluorescence compensation, gating strategies, controls, and visualization methods, please refer to our flow cytometry application guide.

When surface-stained cells are live and not fixed or permeabilized, they can be separated using fluorescence-activated cell sorting (FACS). With FACS, live cells can be sorted into distinct populations based on their properties. We can then perform downstream analyses on the separated cells.

For more information, check out our free online flow cytometry training designed to help you get the best possible data from your cells.

Comparison to other methods

Flow cytometry offers rapid, quantitative, and multiparametric analysis of thousands of cells per second compared to techniques like immunohistochemistry or western blotting. Unlike microscopy, it provides statistical power and population-level insights. While ELISA quantifies protein levels, it lacks single-cell resolution. Flow cytometry also surpasses mass cytometry in accessibility and cost, though the latter allows for higher multiplexing. The ability to analyze both surface and intracellular markers in a single workflow makes it more versatile than methods limited to one compartment. Its compatibility with live or fixed cells further enhances its utility across diverse experimental designs.

Applications

Flow cytometry is widely applicable in immunophenotyping, cell cycle analysis, apoptosis detection, and intracellular signaling studies. It is particularly valuable in immunology for identifying T cell subsets, B cells, and monocytes based on surface markers. Intracellular staining enables the detection of transcription factors and phosphorylated proteins, supporting research in inflammation, cancer, and drug response. The method is also used in clinical diagnostics and vaccine development. The flexibility of flow cytometry allows adaptation to various cell types and experimental conditions, supporting a wide range of experimental design considerations, including panel setup and control selection.

Limitations

Despite its strengths, this protocol has limitations. Intracellular staining requires fixation and permeabilization, which can alter epitope accessibility and affect antibody binding. Over-fixation may reduce signal intensity, while under-permeabilization can hinder intracellular access. Fluorescence overlap between dyes can complicate data interpretation, necessitating careful panel design and compensation. Dead cells may bind antibodies nonspecifically if not excluded properly. Additionally, flow cytometry does not provide spatial context, unlike imaging techniques. Sample preparation must be optimized to avoid clumping and ensure single-cell suspensions. Finally, instrument calibration and operator expertise significantly influence data quality and reproducibility.

Troubleshooting

Common issues in this protocol include weak signal, high background, and poor cell viability. Weak staining may result from low antibody concentration, inadequate fixation, or expired reagents. High background often stems from insufficient washing or non-specific binding; using Fc block and titrating antibodies can help. Poor viability may be due to harsh centrifugation or prolonged handling. Keep cells cold and minimize processing time. Check permeabilization efficiency and antibody compatibility with fixation conditions if intracellular targets are not detected. For clumped samples, filter cells before acquisition. Always include appropriate controls, such as isotype and fluorescence-minus-one (FMO), to validate staining specificity.