Flow Cytometry Assay（FCM） Detailed Protocol (For Reference Only)

Flow Cytometry (FCM) is a high-throughput analytical technique that uses fluorescent-labeled antibodies to specifically bind to target molecules (e.g., surface antigens, intracellular proteins) in cells. It detects and quantifies fluorescent signals from individual cells as they pass through a laser beam, enabling rapid analysis of cell phenotype, viability, and functional status. The detailed procedures (taking immunophenotyping of surface antigens as an example) are as follows:

I. Pre-Experimental Preparation

Reagent Preparation:

Basic Reagents: Phosphate Buffered Saline (PBS, pH 7.4), Fetal Bovine Serum (FBS, for blocking non-specific binding), cell dissociation buffer (e.g., EDTA-free trypsin, for adherent cells), 7-Aminoactinomycin D (7-AAD, for viability staining), fixation buffer (4% paraformaldehyde, optional for intracellular staining), permeabilization buffer (0.1% Triton X-100 in PBS, optional for intracellular staining).

Core Reagents: Fluorescent-labeled primary antibodies (e.g., FITC/PE/APC-conjugated antibodies targeting surface antigens of interest), isotype control antibodies (same species and fluorochrome as primary antibodies, for excluding non-specific binding), fluorescently labeled secondary antibodies (if using unlabeled primary antibodies).

Equipment Preparation: Flow cytometer (equipped with corresponding lasers for detecting selected fluorochromes), centrifuge, pipettes and tips, flow cytometry tubes (12×75mm polystyrene tubes), biosafety cabinet, incubator (37℃, 5% CO₂).

Sample Pretreatment:

Suspension Cells: Collect cells in logarithmic growth phase, centrifuge at 300×g for 5 minutes at 4℃, discard supernatant. Wash twice with pre-cooled PBS, resuspend in PBS containing 2% FBS, and adjust cell concentration to 1×10⁶ cells/mL.

Adherent Cells: Discard culture medium, wash once with pre-cooled PBS. Add appropriate amount of cell dissociation buffer, incubate at 37℃ for 2-5 minutes until cells detach. Neutralize with complete medium, transfer to flow cytometry tubes, centrifuge at 300×g for 5 minutes at 4℃. Wash twice with pre-cooled PBS, resuspend in PBS containing 2% FBS, and adjust cell concentration to 1×10⁶ cells/mL.

Tissue-Derived Cells: Mince tissue into small pieces, digest with collagenase/trypsin mixture at 37℃ for 30-60 minutes, filter through a 70μm cell strainer to obtain single-cell suspension. Centrifuge and wash as above, adjust cell concentration to 1×10⁶ cells/mL.

II. Blocking Non-Specific Binding

Transfer 100μL of cell suspension (≈1×10⁵ cells) to each flow cytometry tube.

Add 5-10μL of FBS (or blocking buffer specific for Fc receptors) to each tube, gently mix.

Incubate at room temperature for 15-20 minutes (or 4℃ for 30 minutes) to block Fc receptors on cell surfaces and reduce non-specific antibody binding.

III. Antibody Incubation

Surface Antigen Staining:

Add appropriate amount of fluorescent-labeled primary antibody to each tube (according to manufacturer’s instructions, e.g., 1-5μL per 1×10⁵ cells).

For isotype control tubes, add the same volume of isotype control antibody (matching the species and fluorochrome of the primary antibody).

Gently vortex to mix, incubate in the dark at 4℃ for 30-60 minutes (avoid light to prevent fluorochrome quenching).

After incubation, add 2mL of pre-cooled PBS to each tube, centrifuge at 300×g for 5 minutes at 4℃, discard supernatant. Repeat washing twice to remove unbound free antibody.

Intracellular Antigen Staining (Optional):

After surface antigen staining and washing, add 100μL of 4% paraformaldehyde fixation buffer to each tube, incubate at room temperature for 15-20 minutes to fix cells.

Centrifuge at 500×g for 5 minutes, discard supernatant. Wash once with PBS.

Add 100μL of permeabilization buffer, incubate at room temperature for 10-15 minutes to permeabilize cell membranes.

Centrifuge at 500×g for 5 minutes, discard supernatant. Wash once with PBS containing 2% FBS.

Add fluorescent-labeled primary antibody against intracellular antigen, incubate in the dark at 4℃ for 30-60 minutes.

Wash twice with PBS, resuspend cells for detection.

IV. Viability Staining (Optional)

After the final wash, resuspend cells in 100μL of pre-cooled PBS.

Add 1-2μL of 7-AAD staining solution to each tube, gently mix.

Incubate in the dark at room temperature for 5-10 minutes (7-AAD binds to DNA of dead cells, excluding dead cells from analysis).

V. Sample Preparation for Flow Cytometry

Centrifuge the stained cells at 300×g for 5 minutes at 4℃, discard supernatant.

Resuspend cells in 300-500μL of pre-cooled PBS (or sheath fluid) to obtain a single-cell suspension (avoid cell clumping, which may block the flow cytometer nozzle).

Filter the cell suspension through a 40-70μm cell strainer into a new flow cytometry tube (to remove clumps and debris, ensuring accurate detection).

Keep samples on ice and protected from light before loading into the flow cytometer (process within 1 hour to maintain cell viability and fluorochrome stability).

VI. Flow Cytometer Operation and Data Acquisition

Turn on the flow cytometer 30-60 minutes in advance to warm up the laser and stabilize the system.

Calibrate the instrument using standard beads (e.g., fluorescent calibration beads) to ensure accuracy of fluorescence signal detection and compensation settings.

Set up detection parameters: Select appropriate lasers and detectors based on the fluorochromes of the antibodies (e.g., FITC: 488nm laser, 530nm emission; PE: 488nm laser, 575nm emission; APC: 633nm laser, 660nm emission).

Adjust forward scatter (FSC) and side scatter (SSC) parameters to gate the target cell population (e.g., exclude debris and dead cells based on FSC/SSC profiles).

Load the sample tube into the flow cytometer, set the acquisition volume or number of events (usually 10,000-50,000 events per sample) to ensure statistical significance.

Acquire data for all samples (including isotype controls, unstained controls, and single-color controls for compensation) sequentially, saving data in FCS format.

VII. Data Analysis

Open the acquired FCS data using flow cytometry analysis software (e.g., FlowJo, FCS Express).

Gating strategy:

First, use FSC vs. SSC to gate the total cell population, excluding debris and cell clumps.

For viability-stained samples, gate live cells by excluding 7-AAD-positive cells (7-AAD vs. FSC).

Use isotype controls to set the threshold for positive staining (usually 1-2% positive rate for isotype controls).

Quantitative analysis: Calculate the percentage of positive cells for each target antigen and the mean fluorescence intensity (MFI) of positive cells (reflecting the expression level of the target molecule).

Statistical analysis: Compare differences in positive cell percentage or MFI between experimental groups using statistical software (e.g., GraphPad Prism), and generate dot plots, histograms, or contour plots to visualize results.

Key Notes

All reagents and samples should be kept on ice or at 4℃ during operation (except for specific incubation steps) to maintain cell viability and prevent antigen degradation.

Fluorescent antibodies are light-sensitive; all incubation and sample handling steps should be performed in the dark or using opaque tubes to avoid fluorochrome quenching.

Isotype controls, unstained controls, and single-color controls are essential for correcting non-specific binding and setting compensation (critical for multi-color flow cytometry).

Cell concentration and suspension quality are crucial: Avoid excessively high cell concentrations (which may cause clumping) or low concentrations (which reduce data reliability).

For multi-color staining, select antibodies with non-overlapping fluorochromes and perform compensation calibration to eliminate fluorescence spillover between channels.

Strictly follow the flow cytometer’s operating procedures; clean the instrument after use to prevent nozzle blockage and cross-contamination.