FCF Basic Protocols

Materials
  • Staining medium (SM) with azide
  • 80% Ethanol
  • 2 mg/ml RNAse solution
  • 0.1 mg/ml PI solution
  • Nitex mesh (85- µm pore size)
  • FACS tubes [Falcon 2052]
Prepare in advance
  1. RNASE solution: 2mg/ml RNAse A (Bovine Pancreas Type II, Sigma) in HBSS. Divide into 500ul aliquots and freeze.
  2. Propidium iodide solution: 0.1mg/ml PI (Sigma) in HBSS with 0.6% NP-40.
  3. HBSS Staining Medium (SM) with 2% calf serum
Reagents

Staining medium (SM) with azide

  • 1X Hank’s balanced salt solution (HBSS) [InVitrogen/Gibco 14025-092]
  • 10 mM HEPES•NaOH, pH 7.2
  • 2% (v/v) calf serum
  • 10 mM NaN3
  • To 480 ml sterile 1X HBSS add 5 ml sterile 1 M HEPES•NaOH, pH 7.2, 10 ml sterile FBS and 5 ml sterile 1 M NaN3. Store at 4°C.

2 mg/ml RNAse solution

Prepare DNAse-free RNAse from bovine pancreatic RNAse, type IIA [Sigma R 5000]. Dissolve RNAse at 10 mg/ml in 10 mM sodium acetate, pH 5.2. Heat at 100°C for 15 minutes. Allow to cool to room temperature. Adjust pH to 7.4 by the addition of 1/10 volume 1 M Tris-HCl, pH 7.4. Aliquot and store at -20°C.

Alternatively, purchase RNAse A solution from Qiagen (100mg/ml stock, 17,500 units, Cat #19101). This stock can be stored at room temperature.

0.1 mg/ml PI solution

  • 0.1 mg/ml propidium iodide [Molecular Probes, P1304]
  • HBSS (no serum)
  • 0.6% (w/v) NP-40
  • Aliquot at 500 µl/aliquot and store, protected from light, at -20°C.
Protocol
  1. Collect 2×106 cells by centrifugation at 400 xg for 5 minutes at 4°C. Aspirate supernatant.
  2. Spin and re-suspend pellet in 50 µl SM/CS and then add it to a conical polypropylene tube containing 1 ml of ice-cold 80% ethanol. Vortex quickly. Fix for 30 minutes to overnight at 4°C.
  3. Collect fixed cells by centrifugation at 400 xg for 5 minutes at 4°C. Aspirate supernatant. Re-suspend cell pellet in 500 µl 2 mg/ml RNase A solution. Incubate for 5 minutes at room temperature.
  4. Add 500 µl 0.1 mg/ml PI solution. Vortex to mix. Incubate for 30 minutes at room temperature, protected from light.
  5. Centrifuge at 400 xg for 5 minutes at 4°C. Re-suspend cells in 500 µl SM by gentle vortexing.
  6. Filter cell suspension through Nitex into FACS tubes.
  7. Collect data on cytometer using CELLFIT software.
Materials
  • Staining medium (SM)
  • Sterile filtered calf serum (CS) [0.45 µm TC sterile filtered stored TC fridge]
  • 1X Gey’s solution
  • 0.1% Trypan Blue
  • 1 mg/ml propidium iodide (PI) [in ddH2O] (store frozen in 50 ul aliquots – can thaw & refreeze many times)
  • Nitex mesh, 85- µm mesh size
  • 4 ml conical tubes [Diamed STK 8550]
  • 5 ml round bottom tubes [Falcon 2052]
  • Antibodies as required for experiment

Sample note
For each FACS sample use 0.5-2×106 cells/tube. Up to 4×106 cells/sample can be used in a 50 µl staining volume, especially if analyzing rare cell populations. If larger cell numbers are stained, see Note (3) below. For peripheral blood lymphocytes (PBLs) use 1-8 X 105 cells/tube ie. whatever you can get.

Buffers

Staining media (SM)
1X HBSS with Ca2+/Mg2+; 2% calf serum; 10mM HEPES, pH 7.2; 10 mM NaN3 (to prevent microbial growth). For some cell types it may be preferable to use HBSS without Ca2+/Mg2+. Store at 4°C.

Gey’s solution

  • Stock A: 35.0 g NH4Cl; 1.85 g KCl; 1.13 g Na2HPO4• 7H2O; 0.12 g KH2PO4; 5.0 g glucose
    Bring to 1 L with ddH2O and autoclave.
  • Stock B: 0.42 g MgCl2•6H2O; 0.14 g MgSO4•7H2O; 0.34 g CaCl2
    Bring to 100 ml with ddH2O and autoclave.
  • Stock C: 2.25 g NaHCO3
    Bring to 100 ml with ddH2O and autoclave.
  • 1X Gey’s Solution: 20 parts Stock A; 5 parts Stock B, 5 parts Stock C; 70 parts sterile ddH2O. If not prepared aseptically, add 10 mM NaN3 (to prevent microbial growth). Store at 4°C.

0.1% Trypan Blue
Dilute 0.4% Trypan blue (Sigma, T-8154) 1:4 in 1X PBS, pH 7.2, with 5mM NaN3. Filter through a 0.2 µm filter.

Protocol
  1. Prepare antibody dilutions in SM based on a total staining volume of 50 µl diluted Ab per sample tube. Calculate total volume of antibody solution needed based on number of samples plus one for each stain.

    Note that before using an antibody in an experiment, the optimal antibody concentration for your application should be determined by staining a test cell sample with serial dilutions of the antibody. See Note (5) below.

  2. (a) For ex vivo cells, dissect desired tissue (usually lymphoid tissue) from animal and place on ice in a tissue culture dish with ice-cold SM.
    (b) Make single cell suspension. Place tissue on pre-wet steel mesh screen that is sitting in a 35-mm tissue culture dish. Mince tissue with scissors and push through steel screen using the plunger from a 3 ml syringe. Wash remaining cells through screen with ice-cold SM. Transfer cell suspension from tissue culture dish to a 15-ml Falcon 2096 tube on ice. Keep cells on ice throughout procedure.
  3. For cultured cells, collect cells by decanting media and cells into a 15-ml Falcon 2096 or 50-ml Falcon 2070 tube. Keep cells on ice throughout procedure.

    Exceptions apply for certain cultured cell lines. eg. 293T cells should be kept at room temperature as they tend to die with prolonged incubation on ice. Centrifugation, counting and staining of these cells should be done at room temperature. Staining times should be kept to a minimum to avoid changes in observed phenotype due to metabolism during staining.

  4. Underlay cell suspension with 0.5 ml CS. Collect cells by centrifugation at 400xg for 5 minutes at 4°C (1500 rpm in Beckmann GS6-KR).
    The calf serum underlay helps make a more compact cell pellet and gets rid of some dead cells and cellular debris.
  5. Aspirate supernatant.
    Optional step
  6. Lyse red blood cells by re-suspending cell pellet in 1X Gey’s solution. Incubate at room temperature 2-5 minutes. Add 10 ml SM, mix well and filter through Nitex. Underlay with 1-2 ml calf serum.

    Incubation longer than 5 minutes should be avoided as this will lead to lysis of nucleated cells. Gey’s solution should be used at a concentration of approximately 2×107 cells/ml. For one mouse spleen, resuspend pellet in 4-5 ml Gey’s solution. For the bone marrow from one mouse, resuspend cell pellet in 2 ml Gey’s solution. Thymocytes and lymph node cells do not need to be lysed, if tissues have been dissected properly. Thymocytes are sensitive cells, avoid lysis step if possible.

  7. Collect all cells by centrifugation at 400 xg for 5 minutes at 4°C. Aspirate supernatant.
  8. Re-suspend cell pellets in appropriate volume of SM (try to approximate a final concentration of 1-5×107/ml).
  9. Count live cells by trypan blue exclusion in 0.1% Trypan blue on a hemacytometer.
  10. Aliquot appropriate number of cells (see Sample Note above) into pre-labeled 4 mL FACS staining conical tubes. Add 0.5 mL of SM to each tube and underlay with 0.3 ml CS. Collect cells by centrifugation at 400xg for 5 minutes at 4°C.

    The calf serum underlay helps make a more compact cell pellet, gets rid of some dead cells and cellular debris, makes the supernatant easier to aspirate without loss of cells, and also leaves less residual fluid behind that can further dilute the antibody solution.

  11. Completely aspirate supernatant. Resuspend cell pellet in 50 µL 1° Ab stain in each tube. Incubate for 20-40 minutes on ice. Protect from light if fluorochrome-labeled 1° antibodies are being used.

    Residual fluid can further dilute the antibody solution and adversely affect your staining results.

  12. Add 0.5 mL SM to each tube (i.e., 10 staining volumes) and underlay with 0.3 mL of CS using 5 ¾” pasteur pipettes.
  13. Collect cells by centrifugation at 400xg for 5 minutes at 4°C.
  14. Completely aspirate supernatant. Resuspend cell pellet in 50 µl 2° antibody. Protect from light. Incubate for 20-25 minutes on ice. If biotinylated 1° antibodies were used, incubate avidin-fluorochrome conjugate 2° stage for 30 minutes on ice.

    Residual fluid can further dilute the antibody solution and adversely affect your staining results.

  15. Repeat wash steps 13 & 14.
  16. Aspirate supernatant. Resuspend cell pellet in 0.5 mL of SM containing 1 µg/mL PI. Filter through Nitex screens (to remove clumps and debris) into labeled 5 mL round-bottom FACS tubes (Falcon 2052).
  17. Collect data on flow cytometer. If this step is to be delayed more than 2-3 hours after staining is complete, they can be fixed in 1.6-4% paraformaldehyde and stored at 4°C overnight or for a few days. However, be sure to use SM without PI.
Notes
  1. If using NaN3 in SM, make sure to keep cells cold as the azide kills cells at room temperature.
  2. If cells are being used in subsequent in vivo or in vitro assays, leave out the azide, filter sterilize all solutions and handle cells aseptically. Nitex can be autoclaved in sealed pouches. Alternatively, sterile cell strainers (Falcon #2350; 70µm mesh) can be used if dealing with limited sample numbers.
  3. To scale up staining, keep cells at 4-10 x 107cells/ml. Wash with 5-10x staining volume, and underlay with 0.5-l mL CS. After last wash, filter through nylon mesh and adjust cell density to 1-10 x 106cells/mL in SM with PI.
  4. If you are troubled by high levels of non-specific background staining, try removing antibody aggregates: airfuge reagents (concentrated unconjugated, FITC, TR, or biotin conjugates) for 10 min. For APC/PE conjugates → microfuge for 10 min in cold room. Always keep Ab stocks on ice and in the dark if using directly conjugated preps.
  5. For each 1° or 2° Ab, the saturating concentration must be determined empirically. A 3-fold diltution series is often most useful (eg. 1/50, 1/150, 1/450, etc.) with appropriate cells. Antibody vendors often recommend a concentration to use, but in our experience these are usually excessive and often not optimal. Depending on cell frequency and antigen expression levels, different tissues may require different antibody concentrations. Staining should always be done with Ab at 2X saturation. For further information, consult our Antibody Titration Protocol.
References

1. Holmes, K., Lantz, L.M., Fowlkes, B.J., Schmid, I., and Giorgi, J.V. (2001) Preparation of Cells and Reagents for Flow Cytometry. Coligan et al. (Eds.) In Current Protocols in Immunology, pp. 5.3.1-5.3.24. John Wiley & Sons, Inc.

2. Strober, W. (1997). Common Immunologic Techniques – Monitoring Cell Growth. Coligan et al. (Eds.) In Current Protocols in Immunology, pp. A.3A.1-A.3A.2. John Wiley & Sons, Inc.

Materials
  • Staining medium (SM) [1X HBSS; 2% (v/v) calf serum; 10mM NaN3, 10 mM HEPES [pH 7.2]
  • 1X PBS
  • BD Cytofix (contains 4% paraformadehyde), Cat # 554655OR
  • Paraformaldehyde solution, 16%, Canemco Inc, Cat #017B
  • Filtered calf serum (CS) [0.45 m TC sterile filtered to avoid bacterial growth]
  • 0.1% Trypan Blue
  • Nitex mesh, 85-µm mesh size
  • 4 ml conical tubes [Diamed STK 8550]
  • 5 ml round bottom tubes [Falcon 2052]

Sample note
For each FACS sample use 0.5-2×106 cells/tube. A considerable amount of cell loss will occur during the multiple washes and spins. The CS is used to underlay the cell suspension during washes. DO NOT add PI to the SM used in step 5 (BD protocol) or step 6 (alternate protocol), because the cells are now permeabilized and will all be PI positive.

BD Cytofix protocol
  1. Perform staining for extracellular markers according to our recommended Extracellular Staining Protocol (see FMCF web page) or any other standard FACS staining protocol.
  2. Wash cells by adding 500-1000µl SM, underlay with 250-300 l CS and collect by centrifugation at 400xg for 5 minutes at 4°C (1500 rpm in Beckmann GS6-KR). Aspirate supernatant.
  3. Fix cells by re-suspending cell pellet in 200µl of BD Cytofix Buffer™ per tube. Incubate cells for 20-30 minutes on ice. (Note: Cell aggregation can be avoided by briefly vortexing cells prior to addition of the fixation buffer).
  4. Wash cells by adding 500-1000µl SM (omit the CS underlay at this point because of the fixative in the buffer) and collect by centrifugation as described above. Aspirate supernatant.
  5. Re-suspend cells in 500µl SM, filter through Nitex mesh and store at 4°C, protected from light until ready to run samples on the cytometer.
Alternative protocol
  1. Perform staining for extracellular markers according to our recommended Extracellular Staining Protocol (see FMCF web page) or any other standard FACS staining protocol.
  2. Wash cells in SM by adding 500-1000µl SM, underlay with 250-300µl CS and collect by centrifugation as described above. Aspirate supernatant.
  3. In the meantime, prepare 2% paraformaldehyde solution in 1xPBS (from16% paraformaldehyde stock). This working solution is stable for a week when stored at 4°C, in an airtight container, protected from light, provided that the solution remains at pH 7.
  4. Fix cells by re-suspending the pellet in 500-1000µl of 2% paraformaldehyde. Alternatively, clumping may be best avoided first resuspending cells in 900µl 1xPBS and then adding 100µl 16% parafromaldehyde for a final concentration of 1.6%. Incubate at room temperature for 10 minutes or for 20-30 minutes on ice.
  5. Wash cells in 2ml of 1xHBSS or 1xPBS SM (omit the CS underlay at this point because of the fixative in the buffer) and collect by centrifugation. Aspirate supernatant.
  6. Re-suspend cells in 500 lµSM, filter thru Nitex mesh and store at 4°C, protected from light until ready to run samples on the cytometer.

Note: works well with all fluorochromes, including PE and APC based tandem conjugates. It is recommended that fixed samples be run on the cytometer within 1 week, preferably the next day. Autofluorescence tends to increase and sample quality generally declines with long-term storage of fixed samples.

Objective

To determine optimal voltage settings for forward scatter (FSC) and side scatter (SSC) parameters and to use beads to set these voltage parameters consistently from experiment to experiment.

Optimal voltage settings will allow you to view the populations of interest on scale and to maximally resolve populations that differ in these parameters.

This protocol uses unlabeled BD Calibrite beads (cat# 349502) and 10 micron size beads from Invitrogen’s Flow Cytometry Size Calibration Kit (cat# F-13838) to set voltages for lymphocytes, however a different bead type could be used if your cells are substantially different in size or refractive index from normal lymphocytes. The goal is to use a standardized particle that will appear on scale with the populations of interest. That being said, beads often exhibit a much higher side scatter property than cells of the same size because they have different refractive indices. A particle with a refractive index that is very different from that of the surrounding medium will scatter light more. While the beads mentioned above are on scale on the forward scatter parameter the beads are off scale on side scatter. The side scatter voltage was therefore lowered to record the beads.

  1. Run cells and adjust FSC and SSC voltages so that all populations of interest are on scale and roughly centred. Ensure that populations with different scatter properties are sufficiently resolved from each other. (e.g., lymphocytes and macrophages/granulocytes in murine bone marrow or human peripheral blood). Also consider whether you anticipate running several different cell types of substantially different sizes in the same experiment. In that case make sure to choose FSC/SSC settings that will keep both the smallest and largest cells on scale. In some cases this may require log scaling for FSC or SSC or both parameters. Note that although you can view the acquired data with linear or log scaling in FlowJo after data collection, the optimal voltage settings for placing the cells at the low-mid range of the log scale will likely be very different than those for optimal placement on a linear scale. Therefore, if log or bi-exponential scaling is desired, carry out the voltage optimization while viewing the live events.
  2. Create a worksheet template with histogram plots for FSC and SSC parameters.
  3. Run beads to capture bead target values (i.e. the MFI of the beads for each parameter). Please note that the side scatter voltage may need to be lowered to view the beads on scale. If so, note the difference in the voltage setting.
  4. Create a gate around the peak on each histogram and note MFIs. You can export statistics as a CSV file that can be viewed in or printed out from Excel. Save worksheet template for use in subsequent experiments.
  5. Export and save cytometer instrument settings by right-clicking the Cytometer Settings icon in the Browser and choose “Export”.
  6. For subsequent experiments, right-click an open experiment in the Browser and choose “Import Cytometer Settings”. Click “Yes” to overwrite the current settings. Select the settings file you want to import and click “Import”.
    Note: Cytometer settings include PMT voltages, compensation, threshold and ratio values.
  7. Run beads and adjust voltages so that they meet the bead target values established in step 4.
  8. If the side scatter voltage was adjusted to bring the beads on scale adjust the voltage back the same amount before running samples.
  9. Proceed with experiment.

(BD Biosciences protocol)

Materials
  • Staining medium (SM) [1X HBSS; 2% (v/v) calf serum; 10mM NaN3, 10 mM HEPES, pH 7.2]
  • BD Cytofix/Cytoperm Kit [554714]*
  • Sterile filtered calf serum (CS) [0.45 µm TC sterile filtered]
  • Gey’s solution
  • 0.1% Trypan Blue
  • Nitex mesh, 85-µm mesh size
  • 4 ml conical tubes [Diamed STK 8550]
  • 5 ml round bottom tubes [Falcon 2052]
  • Antibodies as required for experiment

Sample note

For each FACS sample use 0.5-2×106 cells/tube. A considerable amount of cell loss will occur during the multiple washes and spins. The CS is used to underlay the cell suspension during washes. For peripheral blood lymphocytes (PBLs) use 1-8 X 105cells/tube ie. whatever you can get.

  1. If desired, first perform staining for extracellular markers according to the FACS staining protocol.

    Optional: block EC epitopes with unconjugated antibody of choice if investigating the intracellular levels of proteins that are expressed on the cell surface (ex: TCR, CD4. 8 etc.)

  2. Wash cells in SM and collect them by centrifugation at 400xg for 5 minutes at 4°C (1500 rpm in Beckmann GS6-KR). Aspirate supernatant.
  3. Fix and permeabilize cells by re-suspending cell pellet in 200µl of BD Cytofix/Cytoperm Buffer™ per tube. Incubate cells for 20-30 minutes on ice.
  4. Wash cells in 1ml 1xPerm Wash Buffer™ per sample (this buffer comes as a 10x solution, should be diluted to 1x using ddH2O prior to the experiment. You can store any unused portions at 4°C). Collect cells by centrifugation at 400xg for 5 minutes at 4°C. Aspirate supernatant.
  5. Perform intracellular staining by re-suspending the cell pellet in 50µl of the appropriate antibody dilution in 1xPerm Wash Buffer™. Incubate 20-30 minutes on ice, light protected.
  6. Wash cells in 1ml 1xPerm Wash Buffer™ per sample. Collect them by centrifugation at 400xg for 5 minutes at 4°C. Aspirate supernatant.
  7. Resuspend cells in 500 µl SM, filter through Nitex mesh and acquire samples on the cytometer. DO NOT add PI to the SM used in this step, since your cells are now permeabilized and will all be PI positive.
Note
  • You can store cells at 4°C, protected from light for subsequent analysis on the cytometer. It is recommended that fixed samples be run on the cytometer within 1 week, preferably the next day. Autofluorescence tends to increase and sample quality generally declines with long-term storage of fixed samples.
  • Works well with all fluorochromes, including PE and APC based tandem conjugates. The optimal antibody concentrations for intracellular stains tend to be lower than for the same antibody used for extracellular stains. As always, the optimal antibody concentration for your specific application should be pre-determined by running a titration series – see our related “Antibody Titration protocol”.

*These buffers can be purchased separately too: BD Cytofix (contains 4% paraformadehyde), cat # 554655 and BD Perm/Wash (contains saponin and FBS), cat # 554723.

Supersaturating concentrations will increase background and non-specific binding and is not cost-effective. Non-saturating concentration may cause sample-to-sample variation and decrease resolution and sensitivity. At saturating staining concentrations the amount of antibody present is not limiting and is sufficient to stain all relevant antigens without significantly lowering the concentration of free antibody (see Kantor, A. and Roederer, M. (1997) for examples of this calculation). Therefore the antibody concentration but not the number of cells is critical for optimal staining.

The optimal antibody concentration must be determined for each application and set of experimental conditions (including staining time and temperature) and is determined by using a series of dilutions. The manufacturer’s recommended amount should only be used as a reference “starting point” since their titration conditions may not be identical.

Materials
  • Staining media (SM)
  • Sterile filtered calf serum (CS)
  • 0.1% trypan blue
  • 1 mg/ml propidium iodide (PI) [in dIH2O]
  • Nitex mesh, 85-µm mesh size
  • 4-ml conical tubes [Diamed STK 8550]
  • 5-ml round bottom tubes [Falcon 2052]
  • Serial dilution of antibody in SM
  • Cells appropriate for antibody
Protocol
  1. Select appropriate tissue from which to make a single-cell suspension for staining. When titrating antibodies, it is important to have both positive (stained) and negative (unstained) cells within the population to allow a calculation of signal-to-noise ratios. Isolate enough cells for 1-2×106 cells/tube. Remember to include a tube for unstained cells. For titration of 2° antibodies or 2nd stage reagents include a 2° alone control tube.
  2. Prepare 2-3-fold serial dilutions of antibody in SM (eg. 50 µl antibody + 100 µl SM for 1:3, then 50 µl 1:3 dilution + 100 µl SM for 1:9 dilution, etc.). The starting concentration should be ~1X or 2X that recommended by the manufacturer or ~10 µg/ml purified IgG for most antibodies. Prepare at least 5 dilutions, more if you suspect that you have a really concentrated antibody preparation.
  3. For titration of 2° antibodies or 2nd stage reagents, use a previously titrated 1° antibody at the standard dilution (2X saturating) and make serial 3-fold dilutions of the 2° antibody or 2nd stage reagent.
  4. Prepare single-cells suspension of tissues.
  5. Resuspend each cell pellet in 50-µl diluted antibody. Include a positive control for each antibody to be tested. This is usually the last lot of antibody used or, if using a new conjugate/antibody, an antibody with the same specificity, on the same cellular population. Incubate for 20-40 minutes on ice. Protect from light if fluorochrome-labeled 1° antibodies are being used.
  6. Add 0.5 ml SM to each tube (ie. ≥10 staining volumes) and underlay with 0.3 ml CS using 53/4 Pasteur pipettes.
  7. Collect cells by centrifugation at 400 xg for 5 minutes at 4°.
  8. Aspirate supernatant. Resuspend cell pellet in 50 µl 2° antibody, if necessary. Protect from light. Incubate for 20-30 minutes on ice. If biotinylated 1° antibodies were used, incubate fluorochrome-conjugated avidin 2nd stage for 30 minutes on ice.
  9. Repeat steps 6 and 7.
  10. Aspirate supernatant. Resuspend cell pellet in 0.5 ml of SM containing 1 µg/ml PI. Filter through Nitex screens into labeled 5-ml round-bottom FACS tubes [Falcon 2052].
  11. Collect data on flow cytometer.
  12. Analyze data and calculate the stain index.
  13. Select the dilution with the highest stain index.
Buffers

Staining Media (SM)

1X HBSS with Ca2+/Mg2+; 2% calf serum; 10mM HEPES, pH 7.2
Aseptically add sterile calf serum and sterile 1 M HEPES, pH 7.2, to sterile 1X HBSS with Ca2+ and Mg2+. Store at 4°C.

0.1% Trypan Blue

Dilute 0.4% Trypan blue (Sigma, T-8154) 1:4 in 1X PBS, pH 7.2, with 5mM NaN3. Filter through a 0.2 mm filter.

In flow cytometry, optimal voltage settings are important for resolution sensitivity (i.e. the ability to resolve dim signals from background noise). While adjusting voltages so that unstained cells appear in the first decade may be adequate for FITC and PE, this method is not optimal for fluorochromes with longer emissions (red to far-red, ~650 nm and longer). This is because the photo-multiplier tube (PMT) detectors are less sensitive to these wavelengths. In addition, unstained cells emit little autofluorescence in this part of the spectrum. Therefore, the variance of dim signals is very high in the red and far-red channels because electronic noise makes a large contribution to these measurements.

The following protocol describes a “Minimal Noise” method for optimizing PMT voltages for your application (i.e. your specific cell type) to ensure that electronic noise makes only a minimal (10-20%) contribution to the measured signals. Once these optimal voltages have been set, you should check that these settings give a good dynamic range for your experimental samples and compensation controls. If needed, PMT voltages can be reduced slightly if some stained samples or compensation controls are off-scale or if they are above the linear range for that detector. Once these adjustments have been made, we recommend running fluorescent beads and recording their MFI for each parameter. By using these bead target values to set the voltages for each PMT in future experiments, you can be confident that day-to-day variations in instrument performance are accounted for to ensure that the resolution sensitivity for each parameter will be consistent from experiment to experiment.

We suggest two shortcut alternatives for users who prefer not to manually determine the optimal voltages for their specific application.

Shortcut option 1

The first option is to use the CS&T (Cytometer Setup & Tracking) voltages that were previously determined by FCF staff using the BD CS&T software. The principles used to derive the CS&T targets are similar to those in the Minimal Noise protocol described here because both approaches determine the voltage needed to place dim particles above the level of electronic noise. However, the intrinsic CVs and optical properties of the cells are unlikely to be the same as the beads, so the CS&T PMT voltages may not be truly optimal for your application. Nonetheless, they provide reasonable starting PMT voltages for many applications.

By default when a new Experiment is created, it is automatically set to the CS&T voltages (the purple dot beside the cytometer setting icon in the software browser window indicates cytometer settings are set to CS&T voltage settings – once you alter any of the voltages the purple dot will disappear). To apply the current Cytometer Setup &Tracking voltages to an existing experiment, do the following:

  • In the experiment, right-click the experiment-level Cytometer Settings icon and choose “Apply Current CST Settings”. (If you see a warning message that the Cytometer Setup and Tracking settings have expired, click “OK” anyway.)
Please note that the CS&T voltages were determined for the filters that comprise the “New Advanced” configuration. This configuration is the default configuration and was designed for a specific set of fluorochromes in which all detectors are used simultaneously. If you would like to establish CS&T voltages for your own filter configuration please contact FMCF staff.
Shortcut option 2

The second option is to use the “Lymphocyte Minimum Target Values” that the FCF has established. These target values apply only to the filters in the default (New Advanced) configuration. Fresh ex vivo lymphocytes are commonly used to optimize PMT voltages for flow cytometers because they have measurable (but dim) autofluorescence in many (though not all – see above) wavelengths. The Lymphocyte Minimum Target Values were determined using fresh ex vivo murine lymphocytes as described below. Adjusting each PMT voltage to achieve these bead targets will ensure that lymphocytes (and other cell types that have similar or greater amounts of autofluorescence) will be at a gain where electronic noise makes a minimal contribution to the measured signal.

In our experience, the CS&T voltages versus the Lymphocyte Minimal Targets yield very similar MFIs for unstained lymphocytes in the blue-green-yellow-orange detectors. However, we (and BD) have found that the CS&T method consistently yields much higher MFIs (often >100) for unstained lymphocytes and dim beads in the red and far-red detectors. Although both methods produce PMT voltages that provide reasonable resolution sensitivity, we prefer to use the Lymphocyte Minimum Bead Target values so that unstained cells are not placed too high on scale in the red and far-red channels.

Protocol

Part I: Determining minimal application-specific optimal voltages

Objective

Determine a minimal desirable gain using the dimmest objects to be measured (usually unstained cells) where electronic noise does not make a significant contribution to the measured signal.

A reasonable minimal gain is one in which the variance of electronic noise (en) is not more than 20% of the total variance. To set the voltage (also known as gain) where the variance of electronic noise (en) is not more than 20% of the total variance we use a target of between 10% and 20% for unstained cells (i.e. 15%).

Since variance = SD2, we therefore want to set the gain where
SD2en=(0.15)*(SD2target)

The protocol below will guide you through a procedure in which you will adjust the gain until the rSD of your unstained cells equals the rSDtarget.

  1. Using the rSD target values from the posted chart, run unstained cells and adjust voltage until the rSD of the unstained peak roughly meets the rSD target and is within the rSD lower and rSD upper limits. If you have more than one cell type in your experiment, use the ones that you expect to have the lowest levels of autofluorescence for this step.
  2. Check dynamic range.

Put on a fully stained sample at (minimal) target gain. Choose the samples that you expect to have the brightest total signal (i.e., autofluorescence plus fluorochrome signal) for this step.

  • Ensure the positives are on scale in the linear range of the detector and not greater than ≈ 100,000 – 120,000. If they are at or above 200,000, lower the voltage until positives are ≈ 100,000 – 120,000. This will leave some “wiggle room” at the top end of the scale in case some samples are even brighter.
  • Run compensation samples to ensure that they are also within the linear range of the detector (may exceed 100,000 but not into a nonlinear region) and staining is greater than or equal to staining levels of cells. If it’s not there isn’t anything that you can do at this point! Note: The linear range for each detector can be found on the baseline report.

Part 2: Capturing bead target values for future experiments

Objective

To capture bead target values for use in future experiments. This will allow you to set the instrument in a consistent manner from experiment to experiment.

  1. Create a worksheet template with histograms plots for each parameter.
  2. Run SPHERO Rainbow Calibration particles (Spherotech, cat# RCP-30-5A-4) to capture bead target values (i.e. the MFI of the beads for each channel). These are beads that contain a mixture of fluorophores that fluoresce in all channels.
  3. Create a gate around the peak on each histogram and note MFIs. You can export statistics as a CSV file that can be viewed in or printed out from Excel. Save worksheet template for use in subsequent experiments.
  4. Export and save the cytometer instrument settings by right-clicking the cytometer settings icon in the browser and choose “Export”.
  5. For subsequent experiments, right-click an open experiment in the browser and choose “Import Cytometer Settings”. Click “Yes” to overwrite the current settings. Select the settings file you want to import and click “Import”. Note: Cytometer settings include PMT voltages, compensation, threshold and ratio values.
  6. Run beads and adjust voltages so that they meet the bead target values established in step 3.
  7. Proceed with experiment.

For cells whose autofluorescence will change from experiment to experiment (e.g. tumour cells, cultured cells etc.)

Using the Lymphocyte Bead Target Values, highly autofluorescent cells will often sit in the second decade or higher in the Pacific Blue, FITC, PE or PE-TR channels. The amount of autofluorescence is much lower in the red and far-red channels. It is recommended that you DO NOT arbitrarily lower the PMT voltages to bring the “background” down to the first or second decade. This autofluorescence is real fluorescence and should be recorded as such as long as your stained cells are well on-scale. However, if your stained cells are off-scale on the upper end, then lower the voltages just enough to bring their MFI to 100,000-200,000. After that, capture bead target values for future experiments as described above.

Other Useful Protocols