Sample preparation for analysis when using the Cytek Aurora or Becton Dickinson LSR II benchtop flow cytometers

Cell suspensions must be placed in Falcon brand (352052 or 352054 {with caps}) polystyrene 12x75mm test tubes for use with the BD LSR II. The Cytek Aurora accepts any 12x75 polystyrene or polypropylene tube.  The Cytek ImageStream MkII requires 1.5 mL siliconized microcentrifuge tubes. Biohazardous samples should be placed in tubes with caps. Biochemistry Stores carry the correct tubes.

The final cell concentration should be a minimum of 1 x 106 cells per ml for phenotyping, apoptosis, DNA content, GFP or similar types of experiments.

Cytek ImageStream: At least 1 million cells in 50 μL (2x107 cells/mL) in PBS/2%FBS in a 1.5 mL siliconized microcentrifuge tube.

The minimum volume that can be run on Becton Dickinson/Cytek Aurora bench-top instruments is about 300 µl. The minimum recommended volume is 500 µl at a concentration of 1 x 106 cells per ml.

Cytek ImageStream: At least 1 million cells in 50 μL (2x107 cells/mL) in PBS/2%FBS in a 1.5 mL siliconized microcentrifuge tube.

Spectral overlap between fluorochromes in multi-color experiments requires the use of fluorescence compensation controls. (See http://www.drmr.com/compensation/index.html for an in-depth explanation of compensation.)

The Three Rules of Compensation/Spectral Unmixing

  • Controls need to be as bright or brighter than any sample the compensation will be applied to.
  • Background fluorescence should be the same for the positive and negative control (e.g, positive cells vs negative cells, or positive beads vs negative beads).
  • Compensation controls MUST match the exact experimental fluorochrome.

The proper compensation controls include a negative control (unstained cells are recommended) and one tube each of cells (or beads) stained positively with each of the fluorochromes used in the experiment. The negative control (unstained cells) establishes the background fluorescence of the experimental samples and is used to set the baseline PMT (photomultiplier tube) voltages of the instrument. Each of the compensation tubes is subsequently run to establish the spill-over values of each fluorochrome into the other fluorescent channels. It is important that each compensation tube have a population of brightly stained cells (or beads) in order for the spill-over values to be accurately determined.

Several vendors sell beads specifically for use as compensation controls. The beads are stained as if they were cells using the same antibodies and fluorochromes that are used in the experiment, producing both a negative and bright positive population for each color. For experiments that cannot spare cells for compensation, do not have enough positive events, or have only low antigen expression, compensation beads are recommended. If beads are not used, then cells expressing high levels of antigen (does not have to be an antigen of interest in the experiment) are stained with a fluorochrome-conjugated antibody that yields brightly stained cells. Because of the necessity to have brightly stained cells at a relatively high frequency (i.e, above 10% of the population) for accurate compensation, it may be necessary to use the same antibody that stains the high-density antigen while varying the fluorochrome for each tube (see the following example), except when using tandem fluorochromes (see following section about using tandem fluorochromes).

Example: For mouse splenocytes stained with FITC, PE, PerCP, and APC conjugated antibodies, compensation controls should include:

  • tube 1) unstained splenocytes
  • tube 2) anti-CD8 FITC stained splenocytes (or antibody against some other high-density antigen)
  • tube 3) anti-CD8 PE stained splenocytes (or antibody against some other high-density antigen)
  • tube 3) anti-CD8 PerCP stained splenocytes (or antibody against some other high-density antigen)
  • tube 4) anti-CD8 APC stained splenocytes (or antibody against some other high-density antigen)

Or if using beads

  • tube 1) unstained splenocytes
  • tube 2) anti-* FITC stained beads 
  • tube 3) anti-* PE stained beads
  • tube 3) anti-* PerCP stained beads
  • tube 4) anti-* APC stained beads

There are exceptions to these rules when using tandem fluorochromes. See the next section on this page for details.

            * any antibody that is compatible with the beads

List of Compensation Bead Vendors

Please Note: The facility's mission is to serve investigators in their quest to obtain accurate data. The lack of proper compensation controls may yield misleading, confusing, and inaccurate data. In order to live up to the Facility's mission of assuring quality control and reproducibility, facility staff will not assist with running samples when the necessary compensation controls are not provided by the investigator.

Please Note: When using tandem antibody conjugates in multicolor staining panels, it is important to use exactly the same tandem conjugate (combination of antibody plus fluorochrome) for compensation tubes that are used for staining experiment samples.

Commonly used tandem fluorochromes used for flow cytometry such as PerCP-Cy5.5, PerCP-eFluor710, PE-Cy7, APC-Cy7, BV605, BV650, BV711, BV786, etc. vary in their ability to transfer energy from donor dye to acceptor dye across antibody lots and over time due to fluorescence resonance energy transfer differences. These variations result in leakage of donor dye fluorescence (e.g., PE fluorescence leaks from PE-Cy7) and diminished fluorescence emission strength (brightness) of the acceptor dye over time (e.g., Cy7 has reduced brightness as PE-Cy7 ages due to less energy transfer from PE).

When using tandem antibody conjugates in multicolor staining panels, it is important to use exactly the same tandem conjugate for compensation tubes that are used for staining experiment samples. Otherwise, compensation will not be calculated correctly leading to erroneous measurements and uninterpretable data. Tandem conjugates are also degraded by fixation making it important to run fixed samples stained with tandem conjugates as soon as possible.

Examples where the compensation tube does not equal the experiment tube:

  • PE-Cy7 (BD) ≠ PE-Cy7 (Biolegend) for any antibody conjugate
  • CD4 PE-Cy7 (BD) ≠ CD8 PE-Cy7 (BD) for compensation
  • CD4 PE-Cy7 (BD lot X) ≠ CD4 PE-Cy7 (BD lot Y)
  • CD4 PE-Cy7 (BD lot X) ≠ CD4 PE-Cy7 (BD lot X) formaldehyde fixed
  • CD4 PE-Cy7 (BD lot X newly purchased) ≠ CD4 PE-Cy7 (BD lot X one month old)
  • Substitute compensation beads for cells when the antigen density is low or the positive cells represent a low percentage of the population.

A common problem arises when choosing antibody conjugates for compensation tubes when the cell’s antigen of interest is either low density (i.e., does not yield bright staining) or the positive cells represent a low percentage of the population. This problem is exacerbated when using tandem conjugated antibodies since another antibody with the same tandem fluorochrome cannot be substituted. In these cases, instrument manufacturers and antibody vendors recommend substituting compensation beads for cells. The compensation beads can be stained with the same antibody conjugate used for experiment samples providing a brightly stained sample for compensation. This solves the problem of matching compensation samples with experiment samples when using tandem conjugates. Several vendors sell compensation beads.

  • DiVa software will accommodate multiple, same-tandem compensation tubes.

If it is necessary to use different antibodies conjugated with the same tandem conjugate for different samples in the same experiment, BD’s DiVa software accommodates this by allowing multiple compensation tubes of the same color (e.g., CD3 PE-Cy7, CD19 PE-Cy7).

A Fluorescence Minus One (FMO) control is a tube of cells stained with all fluorochromes used in the experiment except one. A multi-color immunofluorescent experiment has one FMO control for each fluorochrome. FMO controls are used to determine the cut-off point between background fluorescence and positive populations in multi-color immunofluorescent experiments. They are very useful and therefore highly recommended where a positive cell population is presented as a smear instead of being distinctly separate from the negative population. The lack of distinction between positive and negative populations is exacerbated by "spreading" of the negative populations due to the contributions of fluorescence overlap compensation from multiple fluorochromes. In cases where negative population spreading or positive population smearing is present, it is not recommended to use either unstained or isotype controls to determine positive population cut-off points.

Example: For mouse splenocytes stained with FITC, PE, PerCP, and APC conjugated antibodies, FMO controls should include:

  • FITC FMO control) cells stained with PE, PerCP, and APC conjugated antibodies (no FITC)
  • PE FMO control) cells stained with FITC, PerCP, and APC conjugated antibodies (no PE)
  • PerCP FMO control) cells stained with FITC, PE, and APC conjugated antibodies (no PerCP)
  • APC FMO control) cells stained with FITC, PE, and PerCP conjugated antibodies (no APC)

When more than one Brilliant Violet (BD Biosciences) or Super Bright (ThermoFisher / eBioscience) polymer-based fluorochrome is used for staining in the same sample, specialized staining buffer should be used to eliminate non-specific reactivity between the polymers which can result in under-compensation of data.

  • Super Bright Complete Staining Buffer is available from ThermoFisher (Please note that this product is not compatible with UltraComp compensation beads. Use standard staining buffer with UltraComp beads when staining single-color compensation tubes with Super Bright-conjugated antibody. Super Bright Complete Staining Buffer is compatible with Super Bright and Brilliant Violet fluorochromes when both are used for staining in the same tube of cells.)

  • BD Horizon™ Brilliant Stain Buffer is available from BD Biosciences.

  • BD Horizon™ Brilliant Stain Buffer Plus is available from BD Biosciences (This buffer was developed to have a reduced test volume for applications where total staining volume is a concern).

Dead cells tend to be more autofluorescent than live cells, bind antibody non-specifically, and are difficult to completely eliminate from analysis based solely on forward and side scatter. Therefore it is recommended that a fluorescent viablity marker be added to most cell preparations before performing flow cytometry.

The following dyes stain DNA. They identify dead cells by passing through a dead cell's compromised membrane and staining the nucleus. The Flow Cytometry Facility supplies the following two dyes. They can be added to live cell preparations immediately before running on a flow cytometer.

  • Isotonic Prodidium Iodide (PI)

PI has a broad excitation range and emits maximally at 620 nm. It is excited by 488 nm and 561 nm lasers.

  • Hoechst 33258

Hoechst is optimally excited by a 355 nm UV laser, but will also excite with a 405 nm violet laser for live/dead discrimination (405 nm excitation does not work well for cell cycle analysis). It emits predominanlty in the blue region around 460 nm.

Fixable Viability Dyes

Dead cells allow fixable viability dyes to cross their membranes where they stain intracellular amines that are more abundant in the cytoplasm than the extracellular amines on the cell surface of live cells. Cells can be formaldehyde fixed post staining. Cells stained with these products can also be run unfixed.

Cell types such as monocytes, granulocytes, and adherent cell lines tend to form aggregates that will plug the instrument if they're too large. If clumps or strands of cells are visible in your sample tube, it probably means that the cells have aggregated or not dissociated properly. Those samples must be filtered with nylon mesh to remove the aggregates or dispersed by some other method before running on the flow cytometer. Add 0.02mg/ml DNase I (type IIS) (Sigma-Aldrich Cat. No. D4513-VL) to all cell preparation steps, including wash steps, to eliminate free DNA from broken cells that lead to aggregation. Cations must be available to the DNase in order to work properly (i.e., avoid using EDTA). A commercial product, Accumax , has been developed for the specific purpose of keeping cells from clumping. Other sources of large debris such as solid tissue should also be filtered with nylon mesh. In general, anything that looks like a clump or strand in the sample tube is too big to go through the instrument.

When fixing cells for immunofluorescent experiments with formaldehyde, a common problem is increased autofluorescence. The resultant decrease in separation between the negative and positive populations can render some experiments useless. The most common reason for increased autofluorescence is pH drift of the formaldehyde. It is important that correct pH is established in fresh formaldehyde and that pH is monitored as the fixative solution ages.

The Facility's Cytek Aurora makes direct volumetric measurement during sample acquisition, allowing investigators to determine the total cell count per cell sample. The Becton Dickinson flow cytometers do not calculate absolute cell counts (total number of cells per sample). In order to make that calculation using Becton Dickinson flow cytometers, the total volume of cell sample fluid passing through the instrument during data acquisition must be determined. Using a known concentration of beads mixed into the cell sample, the cell sample fluid volume passing through the instrument during acquisition can be established by counting the beads acquired during data acquisition. The cells per unit volume can then be calculated from the volume that passed through the instrument. Several vendors sell beads for this purpose.

(A/B) x (C/D) = number of cells per total volume in the sample tube (cell concentration as cells/uL)

  • A = number of vender beads added to cell sample
  • B = total volume of cell sample
  • C = cell count from acquired data
  • D = bead count from acquired data

Resuspension of cells to be analyzed in media containing phenol red should be avoided whenever possible. Phenyl red may increase the background fluorescence of cells.