The University of Iowa Carver College of Medicine

Flow Cytometry Facility

Sample Preparation

Cell Sample Preparation for Sorting with the Becton Dickinson FACS Aria II or FACS Fusion

Excyte Expert Cytometry recommendations for suspending cells to be sorted.

Analysis vs Cell Sorting - What's the difference?

  • Analyzing - Becton Dickinson FACScan, FACS Calibur, LSR II
    Analyzing involves using a flow cytometer to interrogate cells stained with fluorescent markers and classifying them into groups. Cells pass through the instrument once and go straight to a waste container. There is no cell recovery.
  • Cell Sorting - Becton Dickinson FACS Aria II, FACS Fusion
    Cell sorting involves using a flow cytometer with cell sorting capability to interrogate cells stained with fluorescent markers, classifying them into groups, and then physically seperate the groups into test tubes or multi-well plates. A fraction of the starting population defined by the gating strategy is recovered, the remainder of cells go to a waste container.

Sample Tubes

  • Analyzing - Becton Dickinson FACScan, FACS Calibur, LSR II
    Cell suspensions must 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 Sorting - Becton Dickinson FACS Aria II, FACS Fusion
    Cell suspensions can be placed in any brand of 1.0ml microtubes, 12 x 75mm test tube or 15ml centrifuge tube for use with the FACS Aria II or FACS Fusion. Biohazardous samples should be placed in tubes with caps.

Cell Concentration

  • Analyzing - Becton Dickinson FACScan, FACS Calibur, LSR II
    The final cell concentrations should be 1 x 106 cells per ml for phenotyping, apoptosis, DNA content, GFP or similar types of experiments.
  • Cell Sorting - Becton Dickinson FACS Aria II, 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.
  • Cell Sorting Rates
Nozzle Size Sheath Pressure Maximum 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

Sample Volume

  • Analyzing - Becton Dickinson FACScan, FACS Calibur, LSR II
    The minimum volume that can be run on these instruments is about 300 µl. The minimum recommended volume is 500 µl at a concentration of 1 x 106 cells per ml.
  • Cell Sorting - Becton Dickinson FACS Aria II, FACS Fusion
    For cell sorting, the maximum sample volume can be up to 4 ml for 12 x 75 mm tubes or 14 ml for 15 ml centrifuge tubes. Initial instrument setup for cell sorting requires a minimum of 250 µl at a concentration of 1 x 106 cells per ml. After initial setup, the minimum volume can be reduced to 100 µl at the previous cell concentration if you are using the cell sorter for analysis only. The minimum recommended volume is 250 µl for analysis.
  • 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.

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 II 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 II 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 II 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.

Compensation Controls for Multi-fluorochrome Experiments

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 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)

Eliminating Dead Cells from Analysis

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.

  • Viability Dyes for Live Cell Preparations
  • 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 preperations 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 optimally excited by a 532 nm laser, but is also 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. 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.

  • BD
  • Biolend
  • BioRad
  • Biotium
  • eBioscience
  • Miltenyi
  • Thermo Fisher

Aggregates

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.