Protocols and Guidelines

General Good Practices for Flow Cytometry and Cell Sorting

Basic Instructions and Advice for Your Experiments

  • Sample preparation
    • Flow cytometry requires homogenous single cell suspensions. Check sample quality (and viability) under a light microscope before running your experiment. Samples with visible aggregates should never be put in the instrument.
      • Cell clumps and other aggregates will clog the instrument and interrupt your experiment
      • Debris and dead cells may affect the speed and accuracy of your flow analysis and purity/recovery of your cell sorts
    • Always strain your cells; we recommend using 35 µm filter cap FACS tubes
      • Highly aggregating cell types may even require restraining in the middle of long sample runs.
      • Shapiro's First Law of Flow Cytometry: "A 51 µm Particle CLOGS a 50 µm Orifice!"
    • Serum free conditions, divalent cation chelation by EDTA (1-5 mM) and/or DNAse treatment (200 µg/mL) may reduce cell aggregation.
    • Typical cell concentration is 1-10 million cells per mL and minimum recommended sample volume 200-300 µL
  • Sample buffers

    Typical FACS buffers consist of Ca/Mg free PBS with little protein (and sometimes other additives) to improve cell viability, decrease unspecific staining, and avoid sticking to plastic surfaces. Suspending cells in culture media is usually not recommended due to relatively high autofluorescence, poor pH buffering in typical flow conditions and presence of adhesion promoting components.

    • Start with PBS containing 0.5% BSA (or 1-2% FBS) and optimize as needed
  • Sample tubes
    • Cell analyzers should use rigid (5 mL) polystyrene tubes to ensure good seal with the instrument O-ring; AriaII cell sorter can use either polystyrene or polypropylene tubes (5 mL or 15 mL)
    • The following tubes are known to work with our instruments but other brands and types may also be used:
      • 5 mL FACS tubes with 35 µm filter caps (Corning, cat#352235)
      • 5 mL FACS tubes with snap caps (Corning, cat#352054)
      • 15 mL conical tubes (Corning, cat#352096)
  • Panel design tips

    The goal of panel design is to optimize assay resolution by creating bright positive populations while minimizing spreading of the negatives (see also compensation controls and FMO controls).

    1. Know your instrument’s optical configuration and fluorophore spectra
    2. Divide fluorophores across multiple lasers and distant emission ranges
    3. Put bright fluorophores on rare cells/low expression markers, dim fluorophores on abundant cells/highly expressed markers
    4. Put high spillover fluorophores on mutually exclusive markers
    5. Use a single dump channel to exclude undesired populations
  • Compensation tips
    1. Single stain controls must be as bright or brighter than actual samples
      • But signal must remain in linear range
    2. Background (=autofluorescence) of negatives and positives must match
      • The two populations must be similar type of cells or beads
    3. The fluorophores in controls and samples must be identical
      • Tandem dyes must come from same vial
      • Treat controls like samples (fix/perm, staining process, storage etc.)
    4. Collect enough events in both negative and positive populations
      • Heat kill cells, induce the desired phenotype, use capture beads instead of stained cells etc. to get clear positive population for rare events.
  • Cell sorting tips

    Common sorting rules:

    • The nozzle size should be at least 5 times greater than cell diameter
    • Total event rate (including debris and dead cells) should be no more than 1/4 of drop frequency

    Our default settings for all cell sorts using AriaII are: 100 µm nozzle, 20 psi pressure and 30 kHz drop frequency. You can typically expect ≥80% sorting efficiency and ≥90% purity when running ≤5 million total cells per hour using purity mask setting.

    • Non-homogenous samples, debris and faster flow rate often decrease the sorting efficiency and the purity of the recovered populations.
    • Observed cell recovery (manual count under light microscope) may be significantly lower than instrument reported sort efficiency if e.g. drop delay is poorly calibrated, cells die during sorting process (pH and pressure changes, drying on tube wall) or cells are lost in downstream processing steps(tube and buffer changes)
    • Recovery volume is 3 mL (of PBS) per 1 million cells
    • On special request we can speed up the sorting process using 70 µm nozzle (mostly for lymphocytes) or improve side stream quality using 130 µm nozzle (for unusually large cells).