Flow Cytometry Protocol
Flow cytometry is a laser based technique that measures physical and chemical properties of cells or particles in a fluid stream, as detailed in the NCBI Bookshelf authoritative technical references. This guide provides a rigorous protocol for designing, executing, and troubleshooting flow cytometry experiments. It is written for researchers and laboratory technicians who need a source bounded, practical framework that covers core concepts, decision points, workflow steps, quality checks, common mistakes, and limits of interpretation. It complements the training resources available from EMBL-EBI Training for biological data analysis.
At a Glance
| Component | Description |
|---|---|
| Core Concepts | Light scatter, fluorescence, compensation, gates |
| Decision Points | Panel design, fluorophore selection, control strategy |
| Workflow Steps | Sample preparation, staining, acquisition, analysis |
| Quality Checks | Viability assessment, compensation controls, FMO controls |
| Common Mistakes | Poor compensation, antibody under titration, too many colors |
| Limits of Interpretation | Spectral overlap, autofluorescence, statistical power |
Core Concepts and Instrument Setup
Flow cytometry relies on the principle that cells in a focused stream intersect a laser beam. The scattered light and emitted fluorescence are collected by detectors. Forward scatter correlates with cell size, and side scatter correlates with granularity. Fluorescence detection depends on fluorophore excitation and emission spectra. Compensation is the mathematical process that corrects for spectral overlap between fluorophore emission patterns. Improper compensation leads to false positive or false negative signals.
Instrument setup requires laser alignment, photomultiplier tube voltage calibration, and threshold adjustment. Use reference beads to verify instrument performance daily. The Galaxy Training Network offers open training materials on instrument quality control and data acquisition workflows. Set appropriate forward scatter and side scatter gains to capture the target cell population without saturation. For fluorescence channels, adjust voltages so that the unstained population appears in the first decade of a logarithmic scale. Avoid spreading or clipping signals.
Decision Points in Panel Design
Panel design is the most critical decision making step. It involves choosing which antigens to detect and which fluorophores to pair with each antibody. The guiding principle is to select fluorophores that are bright enough for antigens expressed at low levels and to avoid spectral overlap between channels used for coexpressed markers. Use a systematic approach.
First, list all target antigens based on your biological question. Rank them by expected expression density. Assign the brightest fluorophores (e.g., PE, APC) to dim antigens. Assign dimmer fluorophores (e.g., FITC, Pacific Blue) to bright antigens. Check spectral overlap using online tools or reference charts. The Bioconductor project provides software packages, such as flowCore and CATALYST, that include functions for panel design and spillover matrix evaluation. Second, consider reagent availability. Validate that each antibody clone is tested for flow cytometry and works with your sample type. Third, titrate every antibody to determine the optimal concentration that maximizes the signal to noise ratio. Undertitration produces poor separation, and overtitration increases background.
Include at least one viability dye to exclude dead cells. Choose a dye that emits in a channel not used for key markers. For example, if you use a green laser based channel for a viability dye, avoid that channel for other fluorophores.
Practical Workflow for Sample Preparation and Acquisition
The workflow follows a defined sequence: sample collection, cell isolation, staining, washing, acquisition, and analysis. Each step requires careful execution.
Start with fresh or properly cryopreserved samples. For suspension cells, wash with cold phosphate buffered saline containing bovine serum albumin or fetal bovine serum. Resuspend and filter through a 40 or 70 micron cell strainer to remove clumps. Count cells and adjust to a concentration of 1 x 10^6 to 1 x 10^7 cells per milliliter. For adherent cells, harvest using a non enzymatic reagent or trypsin, and wash immediately. Note that trypsin can affect surface epitope detection.
Block nonspecific binding by incubating cells with Fc receptor blocking reagent for 10 minutes at room temperature. Then add the appropriate antibody master mix. Incubate in the dark for 20 to 30 minutes at 4 degrees Celsius. Wash twice with staining buffer by centrifugation at 300 x g for 5 minutes. Resuspend in a small volume of buffer for acquisition. For intracellular staining, fix and permeabilize the cells first. Follow a validated protocol such as the one described for caspase activated pannexin 1 channels in Methods Cell Biol, which uses flow cytometry to assess intracellular activity.
Acquisition parameters include setting a live gate based on forward and side scatter, then setting viability gate. Acquire at least 10,000 events per sample in the region of interest. For rare populations, collect more events to achieve statistical robustness. Use low flow rate to reduce coincidence and improve resolution. The NCBI Sequence Read Archive stores flow cytometry data in standard formats, but note that it is primarily a sequencing repository. For flow data, use repositories such as FlowRepository.
Data analysis involves gating using software like FlowJo or open source alternatives. Define gates sequentially: singlet gate, live gate, then marker specific gates. Apply compensation based on single stained controls.
Quality Checks and Controls
Quality checks ensure data reliability. Include the following controls in every experiment.
Unstained control measures autofluorescence and sets baseline voltages. Single stained controls are required for each fluorophore to create a compensation matrix. Use beads or cells stained with only one antibody. Fluorescence minus one (FMO) controls are tubes that contain all antibodies except one. FMO controls establish the gate boundary for a given marker by showing the background from other fluorophores. Viability controls help confirm that dead cells are excluded from analysis.
Check compensation performance by verifying that the median fluorescence of positive and negative populations aligns correctly for each fluorophore. The EMBL-EBI Training materials emphasize the importance of documenting all settings and recalibrating the instrument between experiments. Run a sample of known phenotype to validate the panel performance. If results deviate, check instrument optics, reagent stability, and sample handling.
Common Mistakes and How to Avoid Them
Several mistakes occur frequently in flow cytometry experiments.
First, poor compensation due to inadequate single stained controls. Use cells or beads with the same fluorophore for compensation, and ensure that the control population has strong enough signal to calculate spillover. Second, overlapped fluorophores causing false dual positive events. Always check spectral overlap before designing the panel. Reduce the number of colors if necessary. Third, using too many fluorophores on a limited laser instrument. Each laser can only excite fluorophores within its wavelength range. Plan your panel around your laser lines. Fourth, not titrating antibodies. Undertitrated antibodies produce weak separation, and overtitrated antibodies cause high background. Perform titration for each new lot. Fifth, ignoring cell clumps. Clumps cause false doublet signals that can mimic positive events. Gate out doublets using forward scatter area versus height.
The practical workflow described in J Vis Exp for isolating podoplanin positive mesenchymal stem cells uses an indirect magnetic bead strategy. This protocol highlights the importance of consistent cell handling to avoid activation or aggregation. Another study from BMC Microbiol uses flow cytometry to characterize microbiome changes, underscoring the need for rigorous controls in diverse sample types.
Limits of Interpretation
Flow cytometry data interpretation has well defined limits. Spectral overlap is unavoidable and compensation only approximates the correction. For highly overlapping panels, residual spread can obscure dim signals. Autofluorescence from certain cell types, such as macrophages or granulocytes, can reduce sensitivity. Statistical power depends on event count. For rare subsets, the coefficient of variation increases. Use bootstrap or probability binning for robust statistics.
Resolution is limited by instrument optics, detector sensitivity, and fluorophore brightness. Some antigens are not detectable by flow cytometry due to low expression or lack of validated antibodies. Functional assays using flow cytometry, such as assays of intracellular activity, require careful interpretation because fixation and permeabilization can alter epitope accessibility. The protocol for assessing pannexin 1 channel activity in Methods Cell Biol uses flow cytometry with activity based dyes to capture functional states, but background signals must be measured.
Gating decisions introduce subjectivity. Use automated gating algorithms from Bioconductor packages like flowClust or flowMerge to reduce bias. Always report gating strategy in publications. The limits of interpretation also include the inability to determine absolute cell numbers without counting beads or volumetric measurement.
Frequently Asked Questions
1. What is the best fluorophore for detecting a dim antigen?
Select a bright fluorophore such as PE or APC that is excited by a common laser. Confirm that the fluorophore does not overlap significantly with other channels in your panel. Titrate the antibody to its optimal concentration to maximize signal.
2. How many fluorophores can I include in one panel on a standard cytometer?
A standard cytometer with three lasers can typically detect 10 to 14 colors. The exact number depends on filter configuration and fluorophore brightness. Avoid using more colors than necessary to minimize compensation complexity.
3. Do I need an FMO control for every marker?
Yes, for robust interpretation. FMO controls show the background fluorescence in a channel from other fluorophores. They are essential for establishing gates, especially for markers with dim expression or broad spread.
4. Can I use frozen cells for flow cytometry?
Yes, but with caution. Cryopreservation can affect viability and surface antigen expression. Thaw cells using a rapid thawing protocol, wash immediately, and check viability with a dye such as DAPI or propidium iodide. Compare results to fresh cells.
References and Further Reading
- NCBI Bookshelf: Free biomedical books and technical references covering flow cytometry principles and protocols.
- EMBL-EBI Training: Official training resources for biological data analysis, including flow cytometry data handling.
- Galaxy Training Network: Open bioinformatics workflow training materials with modules on flow cytometry gating and quality control.
- Bioconductor: Open software and documentation for flow cytometry data analysis, including compensation and clustering tools.
- NCBI Sequence Read Archive: Public repository for high throughput sequencing data, also used for flow cytometry data storage in standardized formats.
- Methods Cell Biol: Protocol for assessing caspase activated pannexin 1 channel activity by flow cytometry.
- J Vis Exp: Protocol for isolation of podoplanin positive mesenchymal stem cell subpopulations using magnetic beads, applicable to sample prep.
- BMC Microbiol: Study using flow cytometry to analyze microbiome changes after fecal microbiota transplantation.