Cox Systems Biology

University of Toronto

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Flow Cytometric Analysis of Murine Decidual Immune Cells (E6.5-12.5)

Author: Genlei Lu

Contents

Materials

Tissue Collection
tissue collection materials printable
  • Tissue Digestion Buffer
    • HBSS supplemented with Ca2+ & Mg2+
      • Divalent cations facilitate enzyme function
    • 10% v/v FBS
    • 1% of each of the following from prepared stock solutions:
      • Collagenase IV
      • DNAse I
      • Dispase II
    • When working with uNK cells, enzymatic digestion of tissues has been reported to mask surface marker expression, leading to poor detection. In this case, omit Collagenase and Dispase from the digestion buffer.
  • Dissection kit
    • #5 splinter forceps (Almedic 7727-A10-704)
      • Ensure blades are sharp
    • Surgical scissors
      • Tips/blades should fit into 1.5mL Eppendorf tube
  • PBS/HBSS
    • Interchangeable but should be consistent
  • 1.5mL Eppendorf tubes
  • 15 mL + 50 mL conical tubes
  • 40μm cell strainer
  • Incubator/agitator
  • Hemacytometer
  • Refrigerated centrifuge
Staining
staining materials printable
  • 1.5mL Eppendorf tubes
  • Zombie-series amine-reactive viability stain
    • Alternatives: DAPI, hoerscht, propidium iodide, etc.
  • Antibodies
  • FcX (anti-CD16/32) (Biolegend)
  • Compensation beads
    • For viability: ArcReactive +/- compensation beads (Invitrogen)
    • For antibodies: UltraComp eBeads (Invitrogen)
  • PBS/HBSS
  • 1% w/v PBS-BSA solution
  • Refrigerated centrifuge
  • 4% paraformaldehyde (PFA)

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Tissue Collection

tissue collection steps printable

All centrifuge steps performed at +4C, 500rcf for 5 minutes unless otherwise stated.

  1. Prepare 1.5mL Eppendorf + 2x 15mL + 1x 50mL conical tubes labeled for each sample that will be collected.
    1. 1.5mL + 1x 15mL tube to be loaded with 500μL and 4.5mL digestion buffer respectively; keep cooled in +4C fridge.
    2. Load remaining 15mL tube with 5mL PBS/HBSS.
  2. Euthanize mice per animal facility protocols. Dissect and remove uterine horns & store in 15mL tube containing 5mL PBS. Keep on ice as much as possible.
    1. For optimal viability, dissection of uterine horns (step 4-6) should be completed ASAP, however it is acceptable to delay around 30 minutes to an hour without severe consequences to cell viability (e.g. if intending to complete preparation at alternative site).
  3. Add PBS to a petri dish. The volume should be sufficient to cover the bottom of the dish but insufficient to allow the implantation sites to float.
    1. When possible, chill the PBS/petri dish before use.
    2. Ensure the 1.5mL + 15mL tubes loaded with digestion buffer is prepared, on ice, and nearby.
    3. Tissues must be kept on ice as much as possible.
  4. Working quickly, transfer 1 set of uterine horns to petri dish. Using #5 splinter forceps, remove the myometrium from the decidua. Transfer collected decidua to 1.5mL tube containing digestion buffer ASAP. Complete dissection of remaining implantation sites.
    1. Depending on the population of interest, the myometrium may be discarded at this point.
    2. When possible, a second operator should complete step 7 simultaneously
  5. Discard PBS from petri dish. Rinse and clean dish and tools w/ 70% alcohol, water, and distilled water.
  6. Repeat steps 3-5 for all uterine horns.
  7. Using dissection scissors, cut/mince collected decidua into chunks of approximately 1mm3 in 1.5mL tubes containing digestion buffer.
    1. Enlist a second operator whenever possible as the increased surface area improves O2 access for cells within tissues and therefore viability.
  8. Using a P1000 pipette, transfer the minced decidua in digestion buffer to the corresponding 15mL tube.
    1. Volume within tube should now be approximately 5mL.
    2. Consider cutting off the end of the P1000 tip to improve uptake of tissue chunks.
  9. Invert 15mL tubes to mix tissues. Ensure all tissue chunks are submerged/suspended in buffer.
  10. Place tube in tissue incubator/agitator. Set incubator to +39C & rotate at 120rpm. Incubate 1-1.5hr. Invert all tubes every 15 minutes.
    1. Skip this step if working w/ uNK’s as enzymatic digestion is omitted.
  11. Transfer contents of 15mL tubes to their corresponding 50mL tube through a 40μm cell strainer. Add PBS through strainer to ensure all released cells go through the strainer.
  12. Centrifuge and discard supernatant. Wash with 10mL cold PBS. Repeat for 2-3 times total.
  13. Optional: magnetic cell sorting (MACS) can be performed at this time to enrich for immune cells or other populations of interest. Refer to the protocol for the specific MACS kit.
  14. Count collected cells using hemacytometer.
  15. Aliquot cells into labeled 1.5mL tubes containing 1mL PBS, aiming for 106 cells per tube. Keep on ice.

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Staining

staining steps printable

All centrifuge steps performed at +4C, 500rcf for 5 minutes unless otherwise stated.

Following addition of viability dye, samples should be kept in the dark as much as possible. Work on ice at all times unless otherwise stated.

  1. Prepare 1.5mL Eppendorf tubes for controls as necessary. Transfer cell populations/compensation beads to tubes. For cells, transfer 106 cells in 1mL volume. Follow supplier-provided protocol for compensation beads.
    1. Unstained controls
    2. Fluorescence-minus-one (FMO) controls
    3. Compensation controls (CC)
      1. For abundant and clearly defined markers, collected cells may be used for CC. For low abundance/low signal markers, compensation beads are preferred.
  2. Add 1μL of viability stain to appropriate samples. Incubate 15-30 minutes on ice in the dark.
    1. For cells, this should be 1μL in 1mL of cell suspension.
    2. Ensure that cells are suspended in PBS only. Amine-reactive viability stains can bind to proteins in FBS/BSA/other supplements, leading to drastic increases in background signal.
  3. Equilibrate volumes in all tubes stained for viability. Centrifuge, discard supernatant, and wash with PBS for a total of 2 washes.
  4. Discard PBS supernant. For cells, replace with 0.1mL of 1% BSA in PBS.
    1. For stained compensation beads (i.e. viability), repace with 200-500μL PBS and set aside.
    2. For unstained compensation beads, replace with 0.1mL PBS.
  5. Add 0.5μL FcX to every sample containing cells. Incubate 5 minutes.
    1. FcX = anti-CD16/32 (FcR) to block nonspecific binding.
  6. Add necessary concentrations of fluorophore-conjugated antibodies per supplier recommendations/optimization data to tubes containing 0.1mL cell suspensions + FcX. Incubate 15-30 minutes on ice in the dark.
  7. Top up tubes with PBS-BSA (cells) or PBS (beads) to 1mL volume. Centrifuge & wash in appropriate buffers 2x times.
    1. If using primary-secondary antibody system, repeat step 6-7 using secondary antibodies.
  8. Optional: resuspend cells in 0.5mL 4% PFA and incubate at room temperature for 10 minutes. Centrifuge & wash twice using PBS-BSA
    1. Do so if not analyzing samples immediately.
    2. If stored appropriately (+4C & in the dark), fixed samples are generally stable for 2-3 days and reported to be stable up to 7 days.
  9. Resuspend samples in 200μL PBS-BSA (PBS for beads) into 5mL Falcon flow cytometry tubes with 25μm strainer caps (caps not needed for beads). Analyze immediately unless fixed.

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