The University of Iowa Carver College of Medicine

Flow Cytometry Facility

Sample Preparation

Cell Sample Preparation for Sorting with the Becton Dickinson FACS Fusion

Cell Sample Preparation for Sorting with the Becton Dickinson FACS Aria

Excyte Expert Cytometry recommendations for cell sorting suspension media

Sample Tubes

  • Becton Dickinson FACS Aria and FACS Fusion
    Cell suspensions can be placed in any brand of 12 x 75mm test tube or 15ml centrifuge tube for use with the FACS Aria. Biohazardous samples should be placed in tubes with caps.
  • Becton Dickinson FACScan, FACS Calibur, LSR
    Cell suspensions should only be placed in Falcon brand (352052 or 352054 {with caps}) polystyrene 12 x 75mm test tubes for use with the FACS DiVa. Biohazardous samples should be placed in tubes with caps. Biochemisty Stores carry the correct tubes.

Cell Concentration

  • Analyzing
    The final cell concentrations should be 1 x 106 cells per ml for phenotyping, apoptosis, DNA content, GFP or similar types of experiments.
  • Sorting with FACS Aria or FACS Fusion
    The final cell concentration for cell sorting should be between 5 x 106 and 30 x 106 cells per ml depending on the concentration that the cells tend to aggregate. For lymphocytes, the suggested concentration is 20-30 x 106 per ml. Cell line concentration should be 5-10 x 106 per ml. As a rough estimate, for every 10 million cells per ml, the instrument can be run at 10,000 cells per second. For example, if lymphocytes were concentrated to 20 million per ml, the flow rate at the instrument could be run as high as 20,000 cells per second. Cells larger than lymphocytes require the instrument to be configured with a larger nozzle resulting in a lower flow rate.

Sample Volume

  • Becton Dickinson FACS Aria or FACS Fusion
    For initial setup of the instrument a minimum of 250µl is needed at a concentration of 1 x 106 cells per ml. After initial setup, the minimum volume can be reduced if needed to 100µl at the previous cell concentration for analyzing cells. The minimum recommended volume is 250µl for analysis. For cell sorting, the sample volume can be up to 4ml for 12 x 75mm tubes or 14ml for 15ml centrifuge tubes.
  • Becton Dickinson FACScan, FACS Calibur, LSR
    The minimum volume that can be run is about 300µl. The minimum recommended volume is 500µl at a concentration of 1 x 106 cells per ml.

Sort Collection Media

Improved post-sort cell viability can be acomplished by keeping the sort collection media pH constant and providing a source of protein to the sorted cells. Normal cell culture media uses a CO2 buffering system normally supplied by an incubator. Exposing this media to air during cell sorting allows the pH to drift. It is therefore recommended that PBS or Hepes buffered culture media be placed in the sort collection vessels plus enough serum to replicate culture conditions. The final fluid volume in the sort collection vessel will be a mix of collection media plus cell sorter sheath fluid deposited as result of sorting. The amount of serum should reflect the final expected volume.

Sort Collection Vessels

The FACS Aria and FACS Fusion can sort into 1 ml microtubes, 1.5 ml Eppendorf tubes, 12 x 75 mm test tubes, 15 ml conical centrifuge tubes, 96-well or 384-well plates. Where the sorted population consitutes from 10% to 99% of the original population, 15 ml conical centrifuge tubes should be used. They should be filled with 5 ml of sort collection media (see information in previous section of this page). If the sorted population is less than 10% of the original, then the 15 ml collection tubes should be filled with 10-13 ml of media or 12 x 75 tubes used with several milliliters of media. If sorting into 96-well plates, 100-200 ul (200 ul recommended) of media should be placed in each well prior to sorting.

Spinning the plates post sorting for 30-60 seconds at 300xG will help cells adhere to the plate and increase the number of colonies that will grow.

Cell Sorter Sheath Fluid

Sorted cells ride in droplets composed of sheath fluid on their way to the sort collection tube. Once the cells have arrived in the collection vessel, they are mixed with the sheath fluid from the droplets and culture media that has been placed in the collection tube. There are 3 choices of sheath fluid that can be used in the FACS Aria or FACS Fusion:

  • Facility supplied with antifungal/antibacterial agent
  • Facility supplied without antifungal/antibacterial agent
  • 1X PBS (supplied by investigator)

Both Facility supplied sheath fluids are essentially PBS with or without an antifungal/antibacterial preservative agent (Proclin 300). Most cell types tolerate exposure to the sheath fluid preservative and thrive after sorting. Some cells, such as human stem cells and human dendritic cells, do not tolerate exposure and tend to die quickly. In experiments where cells may not tolerate exposure to the sheath fluid preservative, we recommend substituting either Facility supplied preservative-free sheath fluid or 1X PBS. To allow enough set up time to prepare the instrument using 1X PBS, the lab requesting the sort should bring 4-10L for the FACS Aria or FACS Fusion to the Facility the day before the sort. The amount needed for the sort will depend on the length of time scheduled. Please ask one of the Facility personel for advice on the amount of PBS needed.

Cell Sorting Rates

Nozzle Size Sheath Pressure Sort Rate
     70 um             70 psi 22,000/sec
     85 um             45 psi 11,000/sec
    100 um             20 psi  9,000/sec
    130 um             10 psi  2,500/sec

Compensation Controls for Multi-fluorochrome Experiments

Spectral overlap between fluorochromes in multi-color experiments requires the use of fluorescence compensation controls. (See for an in depth explanation of compensation.) 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 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 fluorchrome for each tube (see the following example).

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 antigent)
  • tube 3) anti-CD8 PE stained splenocytes (or antibody against some other high density antigent)
  • tube 3) anti-CD8 PerCP stained splenocytes (or antibody against some other high density antigent)
  • tube 4) anti-CD8 APC stained splenocytes (or antibody against some other high density antigent)

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

* any antibody that is compatible with the beads

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.

Tandem Fluorochrome Conjugated Antibody Best Practices

  • Use exactly the same tandem antibody conjugates for compensation and experiment samples.

Commonly used tandem fluorochromes used for flow cytometry such as PerCP-Cy5.5, PerCP-eFluor710, PE-Cy7, APC-Cy7, 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 cellís 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, BDís DiVa software accommodates this by allowing multiple compensation tubes of the same color (e.g., CD3 PE-Cy7, CD19 PE-Cy7).

Fluorescence Minus One (FMO) Controls

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)


Certain cell types like monocytes, granulocytes, and adherent cell lines tend to be sticky and form aggregates. These aggregates will plug the instrument (a 77um aggregate will not go through a 76um jet). If you can "see" anything in your sample tube, it probably means that the cells have aggregated. 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 type IIS to all cell preparation steps, including wash steps, to eliminate free DNA from broken cells that leads to aggregation. Cations must be availible 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 you can "see" in the sample tube is too big to go through the instrument.

Just prior to sorting, cells should be filtered through nylon mesh. 70µm mesh filters (Falcon 352350) available through Biochem Stores are recommended. Filtering cells greatly reduces the probability of plugging the instrument during sorting. As a general rule, we will not sort unfiltered samples. We want to insure a successful sort and once the instrument is plugged, it may take as long as an hour to bring the instrument back to its original configuration. Time used to unplug the nozzle and bring the instrument back to sorting status may use up your scheduled time.

Phenyl Red

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

Fixing Cells with Formaldehyde and Increased Autofluorescence

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.