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Protocol for the Preparation and Fluorescent ICC Staining of Non-adherent Cells

The following protocol has been developed and optimized by R&D Systems IHC/ICC laboratory for fluorescent ICC experiments using cell smears.

This protocol provides a basic guide for the preparation, fixation, and fluorescent staining of cell smear samples. Each investigator must determine the precise experimental conditions required to generate a strong and specific signal for each antigen of interest. If R&D Systems primary antibodies are employed, please refer to the product data sheets to obtain the recommended working dilutions. In the staining protocol, signal visualization is achieved using R&D Systems NorthernLights™ range of fluorescent secondary antibodies and reagents. For all other reagents, please follow the manufacturer’s instructions.

Please read the protocol in its entirety before starting.

Protocol for the Preparation of a Cell Smear for Non-adherent Cell ICC

Smearing non-adherent cells across a gelatin-coated slide forms a monolayer of cells that can be easily visualized by ICC.

Reagents Required

  • Deionized H2O
  • Fixative: 4% formaldehyde in PBS (1x)
  • 1X PBS: 0.137 M NaCl, 0.05 M NaH2PO4, pH 7.4

Materials

  • Microfuge tube (1.5 mL)
  • Gelatin-coated microscope slides
  • Hot plate
  • Hydrophobic barrier pen
  • Microfuge

Procedure

  1. Fix the cells with 4% formaldehyde for 20 minutes at room temperature by adding formaldehyde directly to the culture media and adjust to approximately 1 x 106 cells/mL.
  2. Add 1 mL of the cell solution to a 1.5 mL microfuge tube.
  3. Spin down the cells for 30 seconds in a microfuge.
  4. Pour off the supernatant and resuspend the cell pellet in 1 mL of deionized H2O.
  5. Spin down the cells for 30 seconds in the microfuge.
  6. Pour off the supernatant, and resuspend the cell pellet in 200 µL of deionized H2O.
  7. Add 5 µL of the cell suspension to a gelatin-coated slide (3 spots per slide), and smear with the side of a pipette tip.
  8. Place the slide on a hot plate (low heat setting), and allow the liquid to evaporate.
  9. Check the slide under a microscope, and make sure that there are no salt crystals. If salt crystals are observed, wash the slide with water.
  10. Surround the cell spot with a hydrophobic barrier using a barrier pen and air dry.
  11. The slides can be stored at 2-8 °C for up to 3 months or they may be stained immediately.

Protocol for the Fluorescent ICC Staining of Cell Smears

This protocol has been developed and optimized using R&D Systems NorthernLights fluorescent secondary antibodies but can be modified accordingly.

Reagents Required

  • Primary Antibodies
  • Blocking buffer: 10% normal donkey serum, 0.3% Triton® X-100
  • DAPI (4',6-diamidino-2-phenylindole) solution: Add 1 µL of 14.3 mM stock for every 5 mL of PBS. Store any unused DAPI at 2-8 °C, wrapped in aluminum foil
  • Deionized H2O
  • Dilution buffer: 1X PBS, 1% bovine serum albumin (BSA), 1% normal donkey serum, 0.3% Triton X-100, and 0.01% sodium azide
  • Anti-fade mounting medium
  • NorthernLights-conjugated secondary reagents (or equivalent)
  • 1X PBS: 0.137 M NaCl, 0.05 M NaH2PO4, pH 7.4
  • Wash buffer: 0.1% BSA in 1X PBS

Materials

  • Gelatin-coated microscope slide covered with monolayer cell smear.
  • Fine tweezers

Procedure

  1. Wash the slides containing the fixed cells two times in 400 µL of wash buffer
  2. Block non-specific staining by adding 400 µL of blocking buffer, and incubate for 45 minutes at room temperature.
  3. Remove blocking buffer. No rinsing is necessary.
  4. Dilute the unconjugated primary antibody (or fluorescence-conjugated primary) in dilution buffer according to the manufacturer’s instructions. For fluorescent ICC staining of cell smears using R&D Systems antibodies, it is recommended to incubate at room temperature for 1 hour. Alternatively, incubate overnight at 2-8 °C.
    Note: Appropriate controls are critical for the accurate interpretation of IHC/ICC results. All IHC/ICC experiments should include a negative control using the incubation buffer with no primary antibody to identify non-specific staining of the secondary reagents. Additional controls can be employed to support the specificity of staining generated by the primary antibody. These include absorption controls, isotype matched controls (for monoclonal primary antibodies), and tissue type controls.
  5. Wash two times in 400 µL of wash buffer. If using a primary antibody with a direct fluorescent conjugate, go to step 8.
  6. Dilute the secondary antibody in dilution buffer according to the manufacturer’s instructions. Add 400 µL to the wells, and incubate at room temperature for 1 hour in the dark. From this step forward, samples should be protected from light.
    Note: R&D Systems NorthernLights fluorescent secondary antibodies and streptavidin conjugates are bright, resistant to photobleaching, and are ideal for multi-color fluorescence microscopy.
    Note: If a biotinylated antibody was used in step 4, apply the appropriate NorthernLights Streptavidin conjugate in step 6.
  7. Rinse two times in 400 µL of wash buffer.
  8. Add 300 µL of the diluted DAPI solution to each well, and incubate 2-5 minutes at room temperature. DAPI binds to DNA and is a convenient nuclear counterstain. It has an absorption maximum at 358 nm and fluoresces blue at an emission maximum of 461 nm.
    Note: DAPI counterstain can obscure visualization of targets localized in cell nuclei.
  9. Rinse once with PBS and once with water.
  10. Carefully blot to remove any excess water. Dispense 1 drop of anti-fade mounting medium onto the microscope slide and mount using an appropriately sized coverslip.
  11. Visualize using a fluorescence microscope and filter sets appropriate for the label used. Slides can also be stored in a slide box at < -20 °C for later examination.
    Note: Initial IHC/ICC studies often require further optimization and/or additional troubleshooting steps.

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