Shreffler:Confocal Protocols

From OpenWetWare
Jump to navigationJump to search


Collection of protocols used by Alex for the basophil imaging project.

Preparation of Polylysine Coated Slides


  • 12 well tissue culture plate
  • round cover glasses (i.e. "slides")
  • 70% Ethanol (aq)
  • 1X Poly-L-Lysine (1:10, Poly-L-Lysine:dH2O)


  1. Wash circular cover glasses in a small petri dish filled with 70% ethanol. Transfer cover glasses individually to the wells of culture plate with clean forceps and allow to air dry.
  2. Under the hood apply 100 μL of 1X Poly-L-Lysine solution to center of each slide.
  3. Let slides stand for 20 minutes before aspirating off excess Poly-L-Lysine solution. Let slides air dry under hood. Slides can be stored at room temperature until ready to use.

Stimulation of Basophils with Anti - IgE and fMLP




  • For first dilution (1:100), add 1 μL fMLP to 99 μL RPMI.
  • Second dilution, take 3 μL from this dilution of fMLP and add it to a separate tube containing 297 μL RPMI (1:100).
  • Add 250 μL of the second solution to 250 μL of basophil cell solution and incubate for 30 minutes at 37 °C.

RPMI: Unstimulated Control

  • Add 250 μL RPMI to 250 μL basophil cell solution.
  • Incubate for 30 minutes at 37 °C.


  • Add 0.6 μL Anti-IgE to 300 μL RPMI
  • Add 250 μL of this solution to 250 μL of basophil cell solution and incubate for 30 minutes at 37 °C.

Plate & Fix Basophils


  • 30 ml PBS (Phosphate Buffered Saline)
  • 30 ml 4% Paraformaldehyde in PBS (PFA-PBS); made from 5 ml stock PFA in 25 ml PBS


  1. Once basophil cells are extracted from PBMC and stimulated, spin cells down at 300g for 7 minutes.
  2. Aspirate RPMI and add 50μL PBS to each tube.
  3. At 4 °C, apply ~50 μL basophil-PBS solution to center of Poly-L-Lysine-coated slides.
  4. Allow solution to sit for 15-20 minutes, undisturbed at 4 °C, so that cells can settle onto slide.
  5. Check with microscope to ensure cells adequately settled onto slide.
  6. Aspirate liquid from surface of slide, and wash each slide with 1mL PBS. To minimize cell loss allow PBS to run down the side of the well while washing.
  7. Aspirate and fix cells with 1 mL 4% PFA-PBS. Incubate at 4 °C in the dark for 15 minutes.
  8. Keep Cells on ice and in the dark until subsequent procedures are carried out.

Preparing Slides for Ab Tagging


  • 15 mL PBS
  • 100 mL 0.1% Triton X-100 PBS (TxPBS); made from 1 mL stock Triton X-100 in 100 mL PBS
  • 15 mL Stain Buffer (1% bovine albumin in 2 mM EDTA in PBS)


  1. Wash slides once with 1 mL PBS per well.
  2. Wash slides 3X with TxPBS. Each wash should incubate for 5 minutes between washes.
  3. Block non-specific binding by adding 1mL Staining Buffer. Incubate for 30 minutes at room temperature in the dark.
  4. Prepare Ab solution and apply as described in separate procedure.

mAb Staining



  1. Overnight Staining with goat anti-CD203c (1:100 from 200 μg/mL stock); rabbit anti-CD107a (1:400 from 200 μg/mL) in staining buffer
  2. Next morning, wash 3 times for 5 minutes with TxPBS
  3. Stain with biotinylated donkey anti-goat (1:1000), incubate 1 hour.
  4. Wash 3 times for 5 minutes with TxPBS
    Prepare CD63 Ab
    For every 0.5 mL (1 SLIDE) add 4 μL mouse anti-CD63 (1:125) in 15 uL staining buffer, add 8 μL Zenon Alexa 594. Incubate 10 min at RT, add 8 μL Zenon blocking reagent. Incubate 10 min at RT.
  5. Stain with anti-CD63 cocktail, with 1:125 dilution of strepavidin Alexa 488 (for CD203c), and 1:400 donkey anti-rabbit 647 (for CD107a). Incubate for 1 hour.
  6. Wash 3 times for 5 minutes with TxPBS
  7. Fix with 4% PFA. Incubate for 15 minutes.
  8. Aspirate excess PFA, and mount slides with DAPI Vectashield mounting medium. Store at 4 °C in the dark until acquisition.

Plate Setup



discuss this protocol


  1. []