Flow Cytometry – 8 Common Sample Preparation and Intracellular Staining Protocols

Flow cytometry (FCM), as a precise technology for multi-parameter quantitative analysis of heterogeneous cell populations at the single-cell level, relies heavily on the quality of sample pre-processing for accurate and reliable data. Preparing highly viable, non-aggregated single-cell suspensions with stable antigen expression is the cornerstone for subsequent fluorescence compensation, population gating, and signal analysis. Maintaining cell viability, removing dead cell debris, and excluding doublets directly affect the signal-to-noise ratio (S/N ratio) and the effectiveness of gating strategies. Physical dissociation tailored to tissue mechanics, such as spleen, bone marrow, or thymus, or density gradient centrifugation and red blood cell lysis for peripheral blood, must strictly follow standardized operating procedures (SOPs) to prevent excessive cell damage or enzymatic/epitope loss. For intracellular targets like cytokines and transcription factors, precise fixation and permeabilization are crucial for crossing biological membranes, ensuring antibody access, and maintaining specific binding. This article systematically outlines eight typical single-cell preparation and intracellular staining protocols, providing standardized references for establishing high-quality flow cytometry workflows.

Mouse Tissue Single-Cell Suspension Preparation

Spleen Single-Cell Suspension Preparation

• Remove the mouse spleen and immerse it in 1× PBS buffer.
• Using tweezers, gently mash the spleen tissue over a 200-mesh filter (70 μm cell strainer), or use the flat end of a syringe plunger to grind, until no visible red clumps remain.
• Collect the homogenate into a 15 mL centrifuge tube, then wash the filter with 1× PBS and combine the wash with the collected homogenate. Centrifuge at 875×g, 4°C for 5 min. Note: Different centrifuge models have varying effective radii; centrifugal force and speed can be converted using the formula: G = 1.11 × 10⁻⁵ × R × (rpm)².
• Discard the supernatant and resuspend cells in 4 mL 1× RBC lysis buffer. Lyse at room temperature for 5 min, then immediately add 10 mL 1× PBS to stop lysis. Centrifuge at 875×g, 4°C for 5 min.
• Discard the supernatant, resuspend in 5 mL 1× flow cytometry staining buffer, and centrifuge at 875×g, 4°C for 5 min.
• Discard the supernatant, resuspend in 1× staining buffer, filter through a 200-mesh strainer, and count cells. Adjust cell concentration to 1×10⁷ cells/mL.
• Aliquot 100 μL of the cell suspension into a 2 mL tube for flow cytometry staining.

Tips:

• Using the mouse's left and right sides as reference, the spleen is located in the upper left abdominal cavity, elongated, soft, and fragile. Handle carefully to avoid breaking and remove surrounding fat thoroughly.
• Grind thoroughly to maximize cell yield.
• Spleen tissue is highly vascular; ensure complete RBC lysis.
• Keep cells on ice (0–4°C) during all steps to maintain viability.

Bone Marrow Single-Cell Suspension Preparation

• Carefully cut open the mouse abdominal skin with scissors and tweezers, then along the hind limbs cut the leg skin and around the ankle to fully separate the leg skin.
• Carefully remove leg muscles, cut at the joints, and isolate the femur and tibia as intact as possible. Place them in 1× PBS buffer.
• Cut the ends of the femur and tibia to expose the marrow cavity. Using a 10 mL sterile syringe filled with 10 mL 1× PBS, replace with a 1 mL syringe needle, carefully insert into the marrow cavity, and slowly flush out the bone marrow. Repeat 2–3 times until the marrow cavity turns white.
• After flushing, gently disperse cell clumps with a pipette, filter through a 200-mesh strainer, collect filtrate into a 15 mL centrifuge tube, and centrifuge at 875×g, 4°C for 5 min.
• Discard supernatant, resuspend in 2 mL 1× RBC lysis buffer, lyse at room temperature for 5 min, then immediately add 10 mL 1× PBS to stop lysis. Centrifuge at 875×g, 4°C for 5 min.
• Discard supernatant, resuspend in 5 mL 1× staining buffer, centrifuge at 875×g, 4°C for 5 min.
• Discard supernatant, resuspend in 1× staining buffer, filter through a 200-mesh strainer, count cells, and adjust concentration to 1×10⁷ cells/mL.
• Aliquot 100 μL of the cell suspension into a 2 mL tube for flow cytometry staining.

Tips:

• Fully isolate femur and tibia to avoid breaking bones and wasting marrow.
• Flush marrow completely until the cavity appears white and semi-transparent.
• Bone marrow samples have distinct cell populations and fewer RBCs than spleen; 1–2 mL RBC lysis buffer per mouse is sufficient, and lysis should not exceed 5 min.
• Keep cells on ice (0–4°C) throughout the procedure to maintain viability.

Thymus Single-Cell Suspension Preparation

• Using scissors and tweezers, cut a small opening at the sternum, then tear the skin to expose the sternum.
• Cut the anterior sternum and lateral ribs in front of the heart, and carefully remove the thymus from the anterior mediastinum above the heart. Immerse in 1× PBS buffer.
• Place the thymus on a 200-mesh strainer and grind thoroughly with tweezers until no visible clumps remain.
• Collect the homogenate into a 15 mL centrifuge tube, wash the strainer with 1× PBS, and combine the wash. Centrifuge at 875×g, 4°C for 5 min.
• Discard supernatant, resuspend in 1× staining buffer, filter through a 200-mesh strainer, and count cells. Adjust cell concentration to 1×10⁷ cells/mL.
• Aliquot 100 μL of the cell suspension into a 2 mL tube for flow cytometry staining.

Tips:

• Avoid cutting surrounding blood vessels when removing the thymus. If vessels are accidentally cut, wash the thymus multiple times with 1× PBS to remove RBC interference.
• Keep cells on ice (0–4°C) during all steps to maintain viability.

Mouse Platelet Isolation

• Hold the mouse with the left hand to protrude the eyeball, then quickly enucleate using tweezers, allowing peripheral blood (~500–600 μL) to flow into an EDTA-K2 anticoagulant tube. Gently invert the tube to ensure complete mixing of blood with anticoagulant.
• Centrifuge at 150×g, room temperature, for 15 min; the sample separates into three layers: bottom RBC layer, middle buffy coat (WBCs and platelets), and top platelet-rich plasma. Carefully transfer the middle and top layers into a 1.5 mL centrifuge tube.
• Centrifuge at 200×g, room temperature, for 10 min to further remove RBCs and WBCs; transfer the supernatant (platelet-rich plasma) to a 2 mL tube and centrifuge at 861×g, room temperature, for 12 min to pellet platelets.
• Resuspend platelets in 1× flow cytometry staining buffer and keep on standby.
• Use 100 μL of the platelet suspension per 2 mL tube for flow cytometry staining.

Tips:

• Avoid contact with fur during eyeball blood collection to prevent hemolysis, which may affect experimental results.
• After collection, invert the anticoagulant tube to fully mix blood with the anticoagulant and prevent clotting.
• When aspirating the buffy coat and platelet-rich plasma, avoid drawing up the bottom RBC layer to maintain platelet purity.
• Experiments should be conducted at room temperature (21–24°C); temperatures outside this range may activate platelets. For downstream processing, PGE1 and ATP-diphosphatase can be added to PBS to prevent platelet activation.

Mouse Peritoneal Macrophage Extraction

• Three days after intraperitoneal injection of starch broth, immerse the mouse in 75% ethanol for 5 min.
• Place the mouse in a biosafety cabinet and carefully cut open the abdominal skin using scissors and tweezers.
• Using a 10 mL sterile syringe, withdraw 5 mL 1× PBS containing 0.3% heat-inactivated serum, then replace with a 1 mL sterile syringe needle, carefully insert into the peritoneal cavity, slowly inject, and gently massage the abdomen to distribute the fluid. After 3–5 min, aspirate the fluid into a 15 mL centrifuge tube. Repeat once.
• Centrifuge at 875×g, 4°C for 5 min; discard supernatant and resuspend cells in 10 mL complete DMEM (Basic DMEM + 10% heat-inactivated FBS + 1% dual antibiotics). Transfer to a 10 cm culture dish and incubate at 37°C, 5% CO₂ for 5 hours for cell adhesion.
• Remove culture medium and non-adherent cells; wash residual medium with 1× PBS. Add 3 mL trypsin and digest at 37°C for 1 min. Add complete DMEM to stop digestion and gently pipette to detach adherent cells.
• Collect the cell suspension into a 15 mL centrifuge tube, centrifuge at 875×g, 4°C for 5 min.
• Discard supernatant, resuspend in 5 mL 1× flow cytometry staining buffer, centrifuge at 875×g, 4°C for 5 min.
• Discard supernatant, resuspend in 1× staining buffer, count cells, and adjust concentration to 1×10⁷ cells/mL. Use 100 μL per 2 mL tube for flow cytometry staining.

Tips:

• Avoid cutting through the peritoneum when opening the abdominal skin.
• When injecting PBS into the peritoneal cavity, avoid puncturing nearby organs.
• If peripheral blood accidentally enters the cavity, wash the culture dish several times with 1× PBS after 5 hours of adhesion to remove RBC interference.

Human Peripheral Blood Mononuclear Cell (PBMC) Preparation

Density Gradient Centrifugation

• Collect whole blood in heparin or EDTA anticoagulant tubes; dilute 1:1 with 1× PBS. In a 15 mL centrifuge tube, add 5 mL lymphocyte separation medium (density gradient medium, e.g., Ficoll), then carefully and slowly layer the diluted blood on top.
• Centrifuge at 875×g, room temperature, for 20 min with the lowest acceleration and deceleration settings.
• Carefully collect the PBMC layer, add 10 mL 1× PBS, and centrifuge at 875×g, 4°C for 5 min.

Fig. 1. Schematic of human peripheral blood after centrifugation (BOC Sciences Authorized).

• Discard supernatant, resuspend in 5 mL 1× PBS, and centrifuge at 875×g, 4°C for 5 min to further remove residual platelets, plasma proteins, and separation medium.
• Discard supernatant, resuspend in 1× staining buffer, count, and adjust concentration to 1×10⁷ cells/mL.
• Use 100 μL per 2 mL tube for flow cytometry staining.

Tips:

• The ratio of diluted blood to lymphocyte separation medium should be 1:1; for example, 10 mL diluted blood requires 10 mL separation medium, which should be equilibrated to room temperature before use.
• Use a Pasteur pipette to slowly layer diluted blood along the tube wall to maintain a flat interface and a clear separation layer. This step is critical.
• During density gradient centrifugation, acceleration and deceleration must be set to the lowest to prevent mixing of cell layers.
• When collecting PBMCs, move the pipette tip slowly along the top of the PBMC layer from top to bottom, avoiding upper plasma, lower separation medium, RBCs, and granulocytes. Multiple small-volume aspirations improve PBMC purity.
• Process blood samples within 2 hours of collection for optimal results.

Red Blood Cell (RBC) Lysis Method

Unlike other sample types, peripheral blood requires flow cytometry staining prior to further processing:

• Dilute anticoagulated blood with physiological saline (0.9% NaCl); add 100 μL of diluted blood to a 2 mL centrifuge tube.
• Add Fc block antibody and incubate at room temperature for 10 min.
• Add flow cytometry antibodies and incubate at room temperature, protected from light, for 35 min.
• Add 2 mL 1× RBC lysis buffer per tube, resuspend, and incubate at 37°C for 12 min; centrifuge at 382×g, 4°C for 5 min.
• Add 2 mL 1× flow cytometry staining buffer, resuspend, centrifuge at 382×g, 4°C for 5 min.
• Resuspend each tube in 200 μL 1× staining buffer and proceed to flow cytometry analysis.

Tips:

• RBC lysis must be performed at 37°C for complete lysis.
• If detecting RBC surface markers, RBC lysis is not required; dilute peripheral blood approximately 20-fold before antibody incubation.

Comparison of Two Methods

Cell Line Sample Preparation

Compared with tissue single-cell suspensions, preparing cell line samples is simpler and can follow the standard experimental workflow below:

Suspension Cell Line Preparation

• In a biosafety cabinet, gently pipette to fully disperse cells; collect cell suspension into a 15 mL centrifuge tube and centrifuge at 315×g, 4°C for 5 min.
• Discard supernatant, resuspend in 5 mL 1× staining buffer, centrifuge 315×g, 4°C for 5 min.
• Discard supernatant, resuspend in 1× staining buffer, count, and adjust concentration to 1×10^7 cells/mL.
• Use 100 μL per 2 mL centrifuge tube for flow cytometry staining.

Adherent Cell Line Preparation

• In a biosafety cabinet, discard culture medium and wash cells once with sterile 1× PBS.
• Add appropriate amount of trypsin to digest adherent cells; digestion time varies by cell type, generally 2 min at 37°C. (If unsure, observe under microscope: cells turning from spindle-shaped to round indicates digestion is complete.)
• Add complete medium to stop digestion; gently pipette to detach remaining cells and ensure full dispersion.
• Collect cell suspension into a 15 mL tube, centrifuge 315×g, 4°C for 5 min.
• Discard supernatant, resuspend in 5 mL 1× staining buffer, centrifuge 315×g, 4°C for 5 min.
• Discard supernatant, resuspend in 1× staining buffer, count, adjust to 1×10^7 cells/mL, and stain 100 μL per 2 mL tube.

Intracellular Flow Cytometry Staining

After preparing the eight common single-cell suspensions above, surface protein staining can be performed. For intracellular targets such as cytokines and transcription factors, fixation and permeabilization are required. Fixatives preserve cell morphology, while permeabilization reagents create small pores in the membrane, allowing antibodies to access intracellular antigens. The following illustrates human interleukin-2 (IL-2) as an example:

Experimental Workflow

Fig. 2. Workflow for intracellular flow cytometry staining of IL-2–expressing PBMCs cells (BOC Sciences Authorized).

Procedure

• Stimulate human PBMCs to induce IL-2 production; adjust cell concentration to 1×10^7 cells/mL and aliquot 100 μL per 2 mL tube. Add viability dye and incubate at 4°C in the dark for 30 min.
• Add 2 mL 1× staining buffer, centrifuge at 382×g, 4°C for 5 min.
• Discard supernatant; resuspend in 100 μL 1× staining buffer, add Fc block antibody, and incubate at room temperature for 10 min.
• Add appropriate fluorescently labeled surface antibodies to the Fc-blocked cells, mix, and incubate at 4°C in the dark for 35 min.
• Wash with 2 mL 1× staining buffer, centrifuge at 382×g, 4°C for 5 min.
• Discard supernatant; fix each tube with 100 μL 4% PFA at room temperature, protected from light, for 15 min.
• Centrifuge at 861×g, 4°C for 5 min, discard supernatant; resuspend in 1 mL 1× permeabilization buffer to wash away residual fixative.
• Centrifuge 861×g, 4°C for 5 min, discard supernatant; resuspend in 100 μL 1× permeabilization buffer and incubate at 4°C in the dark for 30 min.
• Add fluorescently labeled intracellular antibodies and incubate 35 min at 4°C in the dark.
• Wash with 2 mL 1× permeabilization buffer, centrifuge 861×g, 4°C for 5 min.
• Discard supernatant; resuspend in 200 μL 1× permeabilization buffer and proceed to flow cytometry analysis.

Experimental Results:

Fig 3. Gating strategy and IL-2 expression in CD4+ T cells post-stimulation (BOC Sciences Authorized).

Tips:

• IL-2 is a secreted protein, primarily localized in the cytoplasm and extracellular space.
• For nuclear antigens such as Foxp3/transcription factors, specific fixatives and permeabilization reagents are required.

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