Flow cytometry is a powerful, high-throughput biological technique used to analyze characteristics of individual cells. Being able to analyze, in one sitting, hundreds of different samples for many targets of interest—including but not limited to protein expression—makes this a potent tactic. It is a versatile technique with many applications and widespread use—especially in immunology and clinical settings. It is also used in related areas, such as the analysis of signal transduction, and in microbiology to analyze bacterial cells. Experience with flow cytometry is highly sought after in both academia and biotech. This outline of the general procedure should help with understanding what flow cytometry entails.
- Calculations: Start with the worksheet where calculations and algorithms are set up.
- How many experimental samples are to be analyzed? For example, these could be cells collected from blood, lavage, or tissue.
- Cell count using hemacytometer: count the total number of cells per experimental sample. Use this number to aliquot and dilute cells to 1 million cells in 300 or 500 ul. When centrifuging, make absolutely sure that there are no chunks of tissue or globs of cells.
- How many targets are there? Primarily, fluorescently labeled antibodies are used to target proteins of interest. For each antibody, list the fluorophore used, the excitation, and emission wavelengths. With this information, organize the experiment in such a way that forms optimal panels for analysis. Another commonly used target is CFSE—a fluorescent dye used to stain cells and trace cell proliferation over time.
- These numbers will allow for the calculation of the master mix with the appropriate amounts of antibodies and buffer.
- Have aliquots for analyses as “Unstained” and “Single-stained” controls.
- Centrifuge and resuspend in buffer used for flow cytometry. Once the cells are pelleted, use the infamous technique to “flick, rake, and rack.” This is used as a wash to remove the liquid and replace with new buffer. This is a mark of experience—or perhaps a way for scientists to vent their frustrations on the poor tubes in a potentially cathartic manner.
- Incubate all samples with Fc block for 10 minutes.
- Incubate appropriately with (or without) antibody.
- Spin, flick, rake, and rack several times.
- Decide whether to fix the cells immediately, not fix the cells and analyze them immediately on the cytometer, or fix the cells at a later time (this is not recommended because cells change over time).
- For flow cytometric analysis, turn on the cytometer—likely in advance, depending on the machine, as it will take time for it to connect and perform the fluidic startup sequence.
- Make sure you have appropriate training and approval. Do not leave it running, desert it, nor let the fluid levels run out.
- When all is ready, including your samples and the flow cytometer, create the file on the computer for the new experiment.
- Before running each sample, vortex it briefly to distribute the cells for analysis. Double-check to make sure there are no clumps.
- Create “Compensation controls” and optimize by adjusting voltages per fluorophore to correct for spectral overlap. The principle is explained in technical bulletins here and here.
- Run and record measurements for the sample of unstained cells to begin gating on side-scatter (SSC) vs. forward-scatter (FSC) characteristics for the “Live cell gate.”
- Run and record single-stained cells for each of the compensation controls. Define the “Positive gate” for each. Once voltages are set, they must not be changed again. Write these down as a reference.
- Run the samples through the flow cytometer. This may take a while.
- Check with the lab or facility’s procedure for cleaning and shutting down properly.
After all is done and data is collected, proceed to analysis, using software such as FlowJo.
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