Flow Cytometry

Flow Cytometry (FC) is a laser-based analytical technique used for the rapid identification, multiparametric quantitative analysis, and sorting of individual cells or other biological particles (such as microbeads, organelles, microorganisms) in suspension. This technology serves as a core tool in modern life science fields including immunology, cell biology, oncology, and hematology.
Technical Principle: A cell suspension is hydrodynamically focused by sheath fluid, causing the cells to align in a single file. As each cell passes through the laser beam, it generates two types of scattered light: forward scattered light (FSC), which correlates with cell size/volume, and side scattered light (SSC), which correlates with the internal complexity/granularity of the cell. This allows for the preliminary distinction of different cell populations. Simultaneously, the laser excites fluorophores carried by the cell, and the emitted light at specific wavelengths is collected by the detection system. The intensity of this emitted light reflects the expression level of the target molecule. Through the filter-based optical system that separates distinct fluorescent signals, multiparameter, quantitative analysis of thousands of individual cells can be achieved within seconds^[1][2].
Development and Applications: Flow cytometry enables the multiparametric analysis of individual cells, characterizing the expression of cell surface and intracellular molecules to define cell types within heterogeneous populations, assess the purity of isolated cell subsets, and analyze physical attributes such as cell size and volume. It allows for the simultaneous analysis of multiple proteins, gene expression, and cellular functions (e.g., oxidation, viability, cell cycle, apoptosis, and proliferation) in single cells. Furthermore, Fluorescence-Activated Cell Sorting (FACS), a technology derived from flow cytometry, enables the physical separation of specific cell populations based on their fluorescent signals^[3].

Flow cytometry enables qualitative and quantitative analysis of specific cell surface and intracellular molecules using fluorescently labeled antibodies. It typically involves several steps: sample preparation, cell staining, instrumental acquisition, and data analysis.

1. Sample Preparation

A. Cell Line Samples
• Suspension Cells: Centrifuge at 400 g for 5 min. Discard the supernatant. Resuspend the cell pellet in pre-cooled PBS and adjust to an appropriate final concentration (e.g., 1 × 10⁶ cells/mL).
• Adherent Cells: Digest cells with an appropriate amount of trypsin (preferably without EDTA). Neutralize digestion by adding complete culture medium containing serum. Gently pipette to create a single-cell suspension. Centrifuge at 400 g for 5 min, discard the supernatant. Wash the cells with pre-cooled PBS, then resuspend in pre-cooled PBS to an appropriate final concentration.

B. Blood Samples
a. Density Gradient Centrifugation (for isolation of Peripheral Blood Mononuclear Cells-PBMCs)
1) Dilute anticoagulated whole blood 1:1 with PBS.
2) Carefully underlay with an equal volume of Ficoll separation solution.
3) Centrifuge at 400 g for 20 min at room temperature (ensure the centrifuge's brake is turned off).
4) Carefully aspirate the opaque monolayer at the interface between the PBS and Ficoll layers (containing the PBMCs) and transfer to a new centrifuge tube.
5) Add sufficient PBS buffer to wash the cells.
6) Centrifuge at 400 g for 5 min at 2-8°C. Discard the supernatant.
7) If significant red blood cell contamination remains, add 1-2 mL of Red Blood Cell Lysis Buffer (HY-K3010). Incubate for 3-5 min at room temperature protected from light. Then, add an excess of PBS to stop the lysis reaction.
8) Repeat step 6 to wash the cells.
9) Resuspend the cell pellet in an appropriate volume of flow cytometry staining buffer and adjust the cell concentration.
b. Whole Blood Lysis Method (suitable for direct staining analysis)
1) Place 100 μL of anticoagulated whole blood into a flow cytometry tube.
2) Add pre-titrated, fluorescently labeled antibodies. Mix gently and incubate for 15-30 min at room temperature or 4°C, protected from light.
3) Add 2-3 mL of Red Blood Cell Lysis Buffer (HY-K3010). Vortex to mix, and incubate for 10-15 min at room temperature protected from light, until the solution becomes clear.
4) Centrifuge at 400 g for 5 min. Completely discard the supernatant.
5) Wash the cells 1-2 times with PBS buffer.
6) Resuspend the cells in an appropriate volume of flow cytometry staining buffer, ready for acquisition.

C. Tissue Sample Preparation (e.g., Spleen, Lymph Nodes)
1) Collect the tissue in a culture dish and mince into 2-4 mm³ pieces using scissors or a scalpel.
2) Add an appropriate volume of digestion enzyme working solution (e.g., Collagenase, HY-E70005K) and incubate at 37°C.
3) Gently dissociate the cell suspension by repeated pipetting. Filter the suspension through a 70 μm cell strainer to remove undigested tissue clumps and debris.
4) Collect the filtrate. Centrifuge at 400 g for 5 min at 2-8°C. Discard the supernatant.
5) Resuspend the cell pellet in 2-3 mL of Red Blood Cell Lysis Buffer (HY-K3010). Incubate for 3-5 min at room temperature protected from light to lyse red blood cells.
6) Add an excess of PBS to stop the lysis. Centrifuge at 400 g for 5 min at 2-8°C. Discard the supernatant.
7) Wash the cells 1-2 times with PBS buffer.
8) Resuspend the cell pellet in an appropriate volume of flow cytometry staining buffer and adjust the cell concentration.

2. Cell Staining

A. Surface Staining
1) Blocking: Block Fc receptors using an anti-CD16/CD32 antibody (HY-P99125) or serum to reduce non-specific binding.
2) Antibody Incubation: Add directly conjugated, specific surface antibodies (e.g., anti-CD3, CD4, CD8) to the cell suspension. Incubate on ice or at 4°C for 15-30 min, protected from light.
3) Washing: Add 1-2 mL of staining buffer. Centrifuge at 400 g for 5 min. Discard the supernatant. Repeat this wash 1-2 times to remove unbound antibody.
4) Resuspension: Resuspend the cells in an appropriate buffer, ready for acquisition. If fixation is not required, acquire samples immediately.

B. Intracellular/Nuclear Staining
1) Fixation: After completing surface staining and washing, fix the cells using a fixative (e.g., Paraformaldehyde (HY-Y0333)) to stabilize cell structures.
2) Permeabilization: Treat the fixed cells with a permeabilization buffer to make the cell membrane permeable, allowing antibodies to enter the cell interior.
3) Intracellular Antibody Incubation: Add primary antibodies targeting intracellular/nuclear antigens in the permeabilization buffer system. Incubate protected from light. Wash the cells using permeabilization wash buffer. If using an unlabeled primary antibody, perform incubation with a fluorescent secondary antibody followed by washing.
4) Resuspension: Resuspend the cells in flow cytometry staining buffer, ready for acquisition.

3. Instrument Acquisition

Load the prepared single-cell suspensions onto the flow cytometer. Set the total number of cells or events to collect based on experimental requirements.

4. Data Analysis

4.1 Gating Strategy
A typical gating strategy is as follows:
• Exclude Debris: On an FSC-A vs. SSC-A dot plot, gate on the main cell population, excluding debris and noise with very low FSC and SSC signals.
• Exclude Doublets: Using an FSC-H vs. FSC-A plot, gate on the population along the diagonal to exclude doublets or multiple cell aggregates, ensuring the analysis of single cells.
• Identify Live Cells: Based on viability dye staining, gate on the negative population (live cells).
• Identify Target Lymphocyte Population: Within the live cell gate, based on FSC and SSC characteristics, preliminarily gate the lymphocyte population.
• Phenotypic Analysis: Within the lymphocyte gate, use fluorescent antibody scatter plots to further define target subsets. For example, within the CD3⁺ T cell gate, analyze the proportions of CD4⁺ and CD8⁺ cells.
4.2 Fluorescence Compensation
Using single-color control samples collected prior to the experiment, calculate the compensation matrix automatically or manually via the flow cytometry software. This subtracts the spectral overlap between different fluorescent dyes, ensuring accurate detection channel signals.
4.3 Analysis Results
Based on the set gates and applied compensation, the software will calculate parameters such as the percentage of cells in each gate and the Mean Fluorescence Intensity (MFI) for subsequent statistical analysis and graph preparation.

1. Antibody and Panel Design: When designing multicolor panels, prioritize fluorophores with minimal spectral overlap. Titration is essential for any new antibody or new lot to determine the optimal working concentration that provides the highest signal-to-noise ratio, preventing waste and non-specific staining. Different clone numbers may recognize distinct epitopes of the same antigen, potentially resulting in variations in staining efficacy and optimal staining conditions. Record and verify the clone number of all antibodies used to ensure experimental reproducibility.
2. Control Setup: A complete set of controls must be established, including: an unstained control for voltage adjustment, single-stain controls for fluorescence compensation calculation, FMO controls for precise gating, and isotype controls for assessing non-specific binding background. The absence or improper use of any control will compromise data integrity.
3. Sample Preparation: Maintain cell viability >90%. Dead cells cause significant non-specific staining and release DNA, leading to cell clumping. Incorporate a viability dye early in the staining protocol to exclude dead cells and reduce background. For blood or erythrocyte-rich tissues (e.g., spleen), thorough red blood cell lysis and adequate washing are necessary to prevent interference in the analysis.
4. Cell Staining: All staining steps should be performed on ice or at 4°C in the dark to prevent fluorophore quenching and non-specific antibody binding. Intracellular staining must be performed only after surface staining and fixation, followed by permeabilization. If immediate acquisition is not possible, cells can be fixed with 1-4% paraformaldehyde. Fixed samples should be acquired within 24-48 hours.
5. Instrument Acquisition: Prior to sample acquisition, perform instrument calibration using standard beads to ensure optimal performance of lasers, fluidics, and optical detection systems. Use the unstained or FMO controls to adjust photomultiplier tube voltages for each channel, positioning the negative population within the 10⁰-10¹ linear range. Stained samples should be acquired promptly to minimize fluorescence attenuation and changes in cell morphology. Apply an appropriate flow rate during acquisition to reduce shear stress on cells and ensure stable detection.
6. Data Analysis: Before analysis, the compensation matrix calculated from single-stain controls must be applied. Incorrect compensation directly leads to false-positive or false-negative results. For antigens with continuous expression, avoid arbitrary dichotomization into "positive" and "negative" populations. Employ statistical methods (e.g., comparison with FMO controls) or report changes in Mean Fluorescence Intensity.
7. If cells are intended for subsequent culture, the entire procedure must be performed under sterile conditions. Additionally, sodium azide must not be added to any buffers during staining.