Flow cytometry is a technique for the rapid, multi-parameter, quantitative analysis and sorting of single cells or other biological particles suspended in a fluid.

Its principle is that fluorescently labeled cells form a single-cell stream under the sheath fluid and pass through the laser detection zone sequentially. After laser irradiation, two key signals are generated from the cells: scattered light (reflecting cell size and internal complexity) and fluorescence (revealing specific molecular markers). These optical signals are received by a sophisticated optical system, converted into electrical signals, and processed by a computer to form intuitive graphs, enabling the subdivision and quantitative statistics of heterogeneous cell populations.

In addition, instruments with sorting capabilities can precisely separate specific target cells by charging droplets, achieving high-speed and high-purity cell sorting, which is widely used in research fields such as immunology and oncology.

The setup of control groups is the cornerstone of an experiment, with the fundamental purpose of ensuring the specificity, accuracy, and interpretability of experimental results.

Blank Control: Cells without any antibody staining. Used to determine the physical characteristics (size, granularity) of the negative cell population, serve as an initial reference for voltage adjustment, and observe the autofluorescence background of cells.

Isotype Control: Non-specific antibodies of the same species, isotype, and conjugation as the primary antibody, used to exclude non-specific antibody binding.

Positive/Negative Control: Cell samples known to express/not express the target antigen.

FMO Control (Fluorescence-Minus-One Control): One antibody is omitted from a multicolor staining panel, while all other antibodies are added as usual. After removing one fluorochrome, the spillover of other fluorophores into the channel of the omitted fluorochrome can be visually observed, allowing more accurate gating.

Single-Stained Control: Only one fluorescently labeled antibody is added per tube. This is critical for fluorescence compensation adjustment! It can also assist in voltage adjustment to prevent positive signals from exceeding the detection range.

Sample Preparation → Cell Counting and Viability Detection → Fixation and Permeabilization (Optional) → Blocking → Antibody Incubation → Detection and Analysis

1.Whole Blood/PBMC

Red Blood Cell Lysis: Lyse red blood cells using red blood cell lysis buffer, either before or after staining. Thoroughly wash with PBS after lysis.

Density Gradient Centrifugation (e.g., Ficoll): Used to isolate PBMCs and obtain high-purity lymphocytes.

2.Cultured Cell Lines

Adherent Cell Lines: Digest cells with trypsin (containing EDTA). Observe under a microscope; when cells round up and gaps widen, add complete medium containing serum to terminate digestion. Gently pipette cells to detach them from the culture dish and form a single-cell suspension. Transfer the cell suspension to a centrifuge tube, centrifuge at 300–500×g for 5 min, discard the supernatant, and resuspend and wash with PBS.

Suspension Cell Lines: Directly collect the cell culture medium, centrifuge at 300–500×g for 5 min, discard the supernatant, and resuspend and wash with PBS.

3.Tissue Samples (e.g., Spleen, Tumor)

Mechanical Dissociation: Grind in serum-containing medium or PBS using a syringe plunger, homogenizer, or 70 μm cell strainer.

Enzymatic Digestion: For dense tissues (e.g., solid tumors), digest with a mixed enzyme solution containing collagenase and DNase at 37°C for 15–45 min to dissociate cells. Note that the processed sample must be filtered through a 70 μm strainer to remove cell clumps and debris.

1.Cell Counting

Count using a hemocytometer or automated cell counter.

(1) Mix a small volume of cell suspension with trypan blue or a dye specific for automated cell counters.

(2) Perform counting and calculate cell concentration and viability. Viability should be higher than 90%.

2. Viability Detection

Purpose: Dead cells tend to bind antibodies non-specifically. Viability dyes can distinguish live and dead cells and exclude dead cells during data acquisition and analysis. Live/dead dyes are classified as "fixable" and "non-fixable", with the core difference being whether the dye molecules can penetrate intact cell membranes and whether their nucleic acid binding mode is resistant to disruption by fixatives (e.g., paraformaldehyde).

(1) Non-Fixable Live/Dead Dyes

These dyes are membrane-impermeable. They only enter dead cells with damaged membranes, bind to intracellular DNA or RNA, and emit fluorescence. The intact cell membranes of live cells prevent dye entry. When cells are fixed with aldehyde fixatives (e.g., paraformaldehyde), the membrane permeability of all cells is disrupted. After fixation, even originally live cells become permeable to the dye, resulting in staining of all cells. Therefore, staining and detection with these dyes must be completed before fixation; fixed samples can no longer distinguish live and dead cells.

Common Dyes: Propidium Iodide (PI), 7-Aminoactinomycin D (7-AAD), Ethidium Bromide.

Characteristics: They usually bind to nucleic acids via intercalation or non-covalent interactions. This binding is reversible and non-covalent.

(2) Fixable Live/Dead Dyes

This is a new generation of live/dead dyes that overcomes the major limitations of non-fixable dyes. Mechanism of Amine-Reactive Dyes: These dyes are membrane-impermeable and carry reactive groups (e.g., NHS esters) that bind covalently to free amine groups (-NH₂) on proteins. The membranes of dead cells are compromised, allowing dyes to enter and bind covalently to abundant intracellular free amines, becoming stably fixed inside cells. Live cells have intact membranes that exclude the dye; however, live cells have a small number of free amines on their surface, to which the dye binds weakly. This weak surface binding can be washed away in subsequent washing steps, while the covalently bound dye inside dead cells is not eluted.

Common Dyes: Amine-reactive live/dead dyes. Characteristics: After staining and washing, cells can be fixed and permeabilized. Since the dye inside dead cells is covalently bound, it is not washed away or destroyed by fixatives and permeabilization reagents. Thus, dead cells remain labeled after fixation, while live cells remain unlabeled.

Purpose: Additional fixation and permeabilization steps are required for staining intracellular targets. Fixation preserves the structure of intracellular proteins. Permeabilization disrupts the cell membrane, allowing antibodies to enter and stain intracellular targets.

(1) Common Fixation Reagents

Procedure

1.Fix cells with 4% paraformaldehyde on ice for 15-20 min.

2. Washing: Centrifuge cells at low speed (200×g, 5 min, 4°C) to obtain a pellet, discard the supernatant, and resuspend the pellet in washing buffer. Wash 3 times to remove residual fixative.

Notes

1.The fixation step needs optimization for different antigens.

2.Some epitopes are highly sensitive to methanol; try acetone if any issues occur during detection.

3.Washing times, centrifugation duration, and speed may require optimization.

(2) Common Permeabilization Reagents

Procedure

1.Slowly add pre-chilled methanol to the cells while gently vortexing to permeabilize the cells.

2.Incubate on ice for 10–20 min.

Notes

1.If an organic solvent (methanol, acetone) is used as the fixative, this step is unnecessary, as acetone also permeabilizes cells.

2.The optimal detergent depends on the protein and its localization. Strong detergents such as Triton or NP-40 also dissolve part of the nuclear membrane, making them suitable for nuclear antigen staining. Mild detergents such as Tween 20 or saponin allow antibodies to pass through pores without dissolving the plasma membrane, making them suitable for cytoplasmic antigens, the cytoplasmic face of the plasma membrane, and soluble nuclear antigens.

Purpose: Mainly to reduce non-specific binding, thereby lowering background signals and improving the signal-to-noise ratio and data quality. It is mainly divided into the following two types:

(1) Fc Receptor Blocking Agent: Pre-saturates Fc receptors on the cell surface to prevent non-specific antibody binding via the Fc segment. This step is optional but highly recommended, especially for immune cells (e.g., monocytes, macrophages, B cells with high Fc receptor expression) and experiments using indirect staining methods. Anti-CD16/32 monoclonal antibody (HY-P99125) is the most effective blocker for mouse cells. Purified Homologous Species IgG: For mouse experiments, use purified mouse IgG; for human blood experiments, use human IgG. They occupy Fc receptors and prevent non-specific binding of subsequent antibodies. Fc Blocker: e.g., anti-CD16/32 antibody (for mouse cells), human Fc receptor blocker.

(2) Non-Specific Protein Blocking Agent: Uses inert proteins to occupy potential hydrophobic/charge interaction sites on the cell surface or sample containers. Usually added directly to the staining buffer (e.g., PBS containing 1–5% BSA or 2–10% serum) to provide continuous blocking throughout the staining process and reduce non-specific adsorption of antibodies to cells and tube walls.

1.Bovine Serum Albumin (BSA): Most commonly used, typically 1–5% BSA.

2.Fetal Bovine Serum/Goat Serum.

Procedure

1.Centrifuge cells at low speed (200×g, 5 min, 4°C) to obtain a pellet, discard the supernatant, and resuspend the pellet in blocking buffer.

2. Incubate cells at 4°C in the dark for 30–60 min.

Purpose: Allows sufficient binding of antibodies to the target protein antigen, which determines the localization and specificity of fluorescent signals and further affects the accuracy and reliability of experimental results. Note: This step can choose (i) direct detection, where the primary antibody is directly conjugated to a fluorophore; (ii) indirect detection, where a suitable fluorescently labeled secondary antibody is used to detect the primary antibody. These two methods have their own advantages and disadvantages.

Take direct detection as an example:

Procedure

1.Incubation: Incubate on ice or at 4°C in the dark for 20–30 min.

2. Washing: Add excess washing buffer, centrifuge (300–400×g, 5 min), discard the supernatant. Repeat 1–2 times to remove unbound antibody.

After antibody incubation, fully resuspend cells in an appropriate volume of PBS or sheath fluid for on-machine detection.

Related Product Recommendations: Currently, MCE provides a large number of primary antibodies and secondary antibodies with different fluorescent tags for flow cytometry. Come to the MCE official website to select your preferred antibodies and start your flow cytometry journey!