Guide to Flow Cytometry

Flow cytometry is a commonly used technique to rapidly detect and measure the physical and molecular characteristics of single cells in solutions of mixed cellular populations. Flow cytometry uses lasers to produce scattered light signals indicative of cell size and composition, with the option of including fluorophore-labeled antibodies or fluorescent dyes to detect specific proteins or ligands within a cell or on cell membranes. Researchers then use these fluorescent marker signals to sort or quantify cells for countless biological processes, from cellular proliferation and signaling to programmed cell death.

But, despite its vast number of uses, the intricacies of flow cytometry and how to ensure successful, informative, and robust experiments with this powerful technology could seem overwhelming if you’re unfamiliar with some of its essential aspects.

Here, we break down how flow cytometry works, what it’s commonly used for, how to design a multicolor flow cytometry panel for more complicated analyses, and some handy tips to help you troubleshoot any problems you might have.

What is Flow Cytometry?

The first flow cytometers became commercially available over 40 years ago, allowing scientists to measure only one or two cellular parameters. Now, many years of multi-disciplinary collaboration between biologists, physicists, chemists, and engineers have provided staggering advances in instrument technology, fluorescently conjugated antibodies, and fluorescent dyes. Modern flow cytometers now use multiple lasers and advanced detectors to routinely measure over 20 fluorescent parameters simultaneously at rates of over ten thousand cells per second (Robinson, 2022).

Information about the morphological properties of a cell or the localization of proteins is generally lost with flow cytometry, unlike in immunofluorescence staining followed by microscopy. However, the ability of flow cytometry to rapidly analyze millions of cells at the single-cell level provides scientists with unprecedented sensitivity and statistical power to detect rare cells or potentially important biological phenomena difficult to detect with lower throughput techniques.

Uses of Flow Cytometry

Flow cytometry has extensive uses in research and diagnostics. Researchers can use surface and intracellular markers to define and characterize different cell types in heterogeneous cell populations, particularly in fluids like blood. For instance, immunophenotyping analyses routinely screen human peripheral blood to identify subtypes of T-cells, B-cells, granulocytes, and monocytes via cell size, granularity, and different marker combinations. They are commonly used in the clinic for disease diagnosis or monitoring.

Similarly, other flow cytometry advances have provided extensive insight into a process of programmed cell death called apoptosis. Dysregulation of apoptosis is central to the development of various cancers, autoimmune disorders, and many other aspects of disease. The innovation of various novel reagents combined with advances in flow cytometry allows researchers to assess the level and types of apoptosis via cellular “flags” marking dying but not healthy cells.

For instance, ImmunoChemistry Technologies provides a range of over 20 non-cytotoxic Fluorescent Labeled Inhibitors of CAspases (FLICA®) reagents to covalently bind active caspase enzymes with different fluorescent read-outs available, suitable for multicolor flow cytometry experiments. Caspase enzymes are core components of the apoptosis pathway and directly respond to pro-apoptotic signals to cleave proteins, leading to cell death. Directly monitoring intracellular caspase activity with FLICA® reagents avoids false positives common with other approaches like TUNEL staining and is central to many peer-reviewed studies. For instance, our Green Fluorescent FAM-FLICA® Caspase-1 (YVAD) Assay Kit is supported by over 80 peer-reviewed citations in diverse biological settings and diseases. Researchers can also use our necrosis vs apoptosis assay kit to differentiate between the two types of cell death.

Flow cytometry also excels at determining cell types in complex mixtures. For instance, studies in the mammalian brain have used flow cytometry to identify and study individual synapses based on their isolation by size and with panels of synaptic markers (Prieto et al., 2017). The Anti-VGlut1 Antibody (N28/9) from Antibodies Inc. was instrumental to these findings with flow cytometry, as the VGlut1 protein is core to the uptake of glutamate into synaptic vesicles in excitatory neural cells.

These findings were possible thanks to the well-thought-out design of flow cytometry experiments using the most appropriate reagents available. But, a great experimental design is only possible when researchers thoroughly understand the intricacies of the flow cytometer system.

How Flow Cytometry Works

Flow cytometers come in all shapes and sizes depending on their specific usage.

Broadly, traditional flow cytometers depend on three integrated systems: fluidics, optics, and electronics (McKinnon 2018).

The fluidics system relies on a pressurized stream of buffered saline solution known as sheath fluid surrounding a central core injected with the mixture of cells under investigation. The sheath fluid focuses the sample into a single stream of individual cells, known as hydrodynamic focusing. This ensures that the lasers and optics of the instrument system can analyze each cell separately.

The instrument's optics section comprises one or more beams of focused lasers alongside mirrors, lenses, and filters used to move, collect, and detect light. As each cell passes through one or more lasers, the instrument records it as an event.

The electronics component then detects the photon pulse emitted as a cell passes through the laser beam. The detectors convert these signals into voltage pulses interpreted by the flow cytometer as digital footprints containing information about each cell's different properties.

Light Scattering

For each cell, detectors positioned directly in line with the laser detect the “forward scatter” used to determine cell size. In contrast, other detectors are perpendicular to the cells to measure their “side scatter”, which results from the light hitting physical structures within a cell. This provides readouts of cellular granularity.

Combined forward and side scatter profiles of cells can be used to identify and analyze different cell populations. For instance, flow cytometry can determine granulocytes from monocytes in whole blood based solely on their forward and side scatter profiles. Monocytes are large but not granular, so they have a high forward scatter but lower side scatter, whereas granulocytes are both large and granular, so they have a high forward scatter and high side scatter.

The Power of Fluorescence in Flow Cytometry

For more fine-tuned detection of rarer cell types or broad events in complex cell populations, using fluorescently labeled cellular or functional markers is a potent tool in biomarker detection, cell cycle analyses, cell viability, assessing cellular effects of drug treatments, or countless other applications.

Advances in the design of fluorescent reagents now offer researchers a broad toolbox of sensitive flow cytometry solutions. This includes classical dyes such as Hoechst 33342, which emits blue fluorescence when bound to double-stranded DNA to more advanced dyes responsive to chemical reactions with the cells themselves that detect diverse cellular processes such as oxidative stress or apoptosis.

These read-outs of cellular stress or cell death could be crucial for drug discovery pipelines monitoring the toxicity of novel compounds. This has led ImmunoChemistry Technologies to develop a comprehensive tapestry of flow cytometry reagents suitable for detecting different aspects of the cellular stress and apoptotic processes. For instance, our intracellular glutathione (GSH) assay kit uses a proprietary thiol-sensitive dye to monitor changes in the concentration of reduced GSH, which is highly indicative of the onset of apoptosis.

Similarly, for studies investigating free nitric oxide or nitric oxide synthase activity levels, our nitric oxide synthase assay uses a probe called DAF-2DA to enter the cells. Once internalized, the probe undergoes reactions to release the non-fluorescent DAF-2 dye. The DAF-2 dye then reacts with nitric oxide and O2 to produce its fluorescent derivative, detectable by flow cytometry at a wavelength of 515 nm.

If a cell is fluorescently labeled, lasers excite fluorophores of different wavelengths, which then emit light that is collected by the detectors known as photomultiplier tubes. Specific photomultiplier tubes detect light at particular wavelengths after it passes through a series of lenses, mirrors, and filters that allow specific wavelengths to pass while blocking others.

This enables the simultaneous detection of multiple signals for extensive multicolor flow cytometry experiments. Still, it is important to know a few intricacies in the design of these advanced multicolor panels before starting.

How To Design a Multicolor Flow Cytometry Panel

If you want to assess multiple colors in your flow cytometry panel, there are some essential things to consider when preparing. As the adage goes, “failing to prepare is preparing to fail.”

Here, we lay out the core components of a successful multicolor flow cytometry panel.

Why Use Multicolor Flow Cytometry?

One of the main advantages of multicolor flow cytometry is the ability to interrogate single cells with multiple markers. When conducting exploratory or diagnostic research, especially in a clinical setting, extracting as much information as possible from limited samples, such as liquid biopsies taken from blood or bone marrow, is imperative. Investigating multiple biomarkers simultaneously could be crucial in identifying the specific type of leukemia or lymphoma a patient may suffer from. Multicolor flow cytometry also improves efficiency and reproducibility thanks to smaller sample volumes and increased sample throughput while maximizing data output to define cell populations or cellular processes more accurately.

But, as the number of colors and antigens increases, so too does experimental complexity, and considerations such as fluorophore combinations in your flow cytometry panel are crucial.

Not All Flow Cytometers Are Made Equal

Each flow cytometer is slightly different to one another, so it’s crucial to know the number and type of lasers, the number of detectors, and the type of filters available before starting your multicolor panel design. This will dictate the fluorophore combinations available to you, as fluorophores are only excited by the corresponding wavelength of light from the laser.

Selecting Fluorophore Combinations

The most common lasers used in traditional flow cytometers are 355 nm (ultraviolet), 405 nm (violet), 488 nm (blue), 532 nm (green), 552 nm (green), 561 nm (green-yellow) and 640 nm (red). Emission spectra for fluorophores are shifted rightwards compared to the excitation wavelength. For instance, the commonly used fluorescein isothiocyanate (FITC) dye is excited with a 490 nm laser but has a maximal emission of around 525 nm.

Importantly, there are clear overlaps with the emission spectra of some fluorophores, so these should be considered when selecting a multicolor panel. When overlap in wavelengths occurs, spillover of one fluorophore into the detection channel of another might be observed, making it difficult to reliably identify distinct cell populations. A more robust approach is to select fluorophores with little to no overlap where possible.

Our FLICA® range of caspase-detecting reagents are great for multicolor flow cytometry experiments, as most are available in various colors. For instance, our Far-Red Fluorescent FLICA® 660 Caspase-1 (YVAD) Assay Kit has an emission of 760 nm but is also available as a green fluorescent FAM version with a peak emission of 515-535 nm, both highly popular options with 17 and over 90 independent citations respectively. These options provide extensive flexibility in designing multicolor panels. However, this becomes more difficult for studies including more than three fluorophores due to unavoidable overlaps in emission spectra.

Various fluorescence spectrum viewers are available online to help you design efficient multicolor flow cytometry panels while minimizing spillover.

But, where spillover is unavoidable, there are a few things that you can do.

Fluorescent Compensation for Multicolor Flow Cytometry

Where the available fluorophores overlap, a method called compensation can help address the issue of photons of one fluorophore being observed by multiple detectors. Compensation allows users to determine the relative contribution of each fluorophore to the signal in a specific detector to avoid double positive populations of cells. For example, combinations of FITC and another popular dye called phycoerythrin (PE) have some overlap in their emissions spectrum, so cells positive for FITC might also appear positive for PE.

One way for successful compensation is to include a sample stained only with a FITC-labeled antibody. The user can then adjust the settings to remove all FITC signal from the PE channel. This control should be included for each fluorophore and include positive and negative populations for the best results.

Consider Fluorophore Intensity

Not all targets of fluorescently-conjugated antibodies or fluorescent dyes are common or highly expressed. Markers might only be present on small numbers of cells or detect rare events. So, to ensure highly and lowly expressed markers are evenly detected, use bright fluorophores for low abundance or unknown antigen targets or rare cells and dimmer fluorophores for targets with higher abundance.

This will make it more likely that higher signal-to-noise ratios allow the separation of positive cells from negative populations.

Troubleshooting Tips for Flow Cytometry

So, you’ve designed your flow cytometry panel with the core considerations above, but a failed experiment has left you wondering what went wrong.

Here are a few things you can check to troubleshoot why your experiment didn’t work as expected.

Lack of Signal

  1. The target might not be expressed at all, or it might be intracellular, so it could require a different permeabilization procedure.
  2. There might not be enough cells in the sample, possibly due to loss during washing stages. Ensure cell density remains around 0.5 million cells per mL.
  3. For multicolor experiments, the fluorophore intensity and antigen expression might not be optimally matched. Be sure to use a bright dye for low-expressing antigens or rare cells and vice versa.
  4. The signal might not be correctly compensated for multicolor experiments. To ensure correct compensation, include positive control and negative populations for each fluorophore.
  5. Not enough antibody was used, primary antibodies and secondary antibodies might not be compatible, or antigen-antibody binding might be suboptimal. It’s always best to double-check antibody data sheets or previous uses in the literature and titrate reagents upon first usage.
  6. There could be too many dead cells in the sample, blocking dye or antibody signals with high autofluorescence. Always use fresh preparations of cells or cell viability assays.

High Background

  1. You might have included too much antibody or dye. Titrations of reagents upon first usage can determine the best concentrations for your application.
  2. The antibody or dye used could detect non-specific targets. For antibodies, including an isotype control can help determine background fluorescence from a true positive signal. Including blocking solutions during cell staining can also decrease non-specific binding.
  3. The instrument's gain might be set too high or too low, so a positive and negative control should be used to set this upon first use.

Cell Doublets Observed

  1. Sometimes, two cells enter the laser beam simultaneously. These doublets will show as a second cell population at twice the fluorescent intensity. Ensure cells are mixed or filtered to remove cell clumps.
  2. More than one cell population might be expressing the target protein. Another biomarker antibody or dye could further separate these cell populations.

Suboptimal Cell Scatter Profiles

  1. The instrument's settings might be incorrect for your desired application. Check the manufacturer’s instructions for optimal settings.
  2. Cells might be poorly fixed or damaged, causing cellular debris. Try a fresh batch of stained cells.


Overall, armed with the information provided in this guide, you’re now ready to take on your next flow cytometry experiment with confidence. You’ll generate robust, reliable, and reproducible results in no time, even for multicolor flow cytometry experiments, and will know what to look for if things go wrong.

For more information about flow cytometry and the reagents and solutions we provide, please get in touch with us here.


McKinnon, K.M., 2018. Flow cytometry: an overview. Current protocols in immunology, 120(1), pp.5-1.

Prieto, G.A., Trieu, B.H., Dang, C.T., Bilousova, T., Gylys, K.H., Berchtold, N.C., Lynch, G. and Cotman, C.W., 2017. Pharmacological rescue of long-term potentiation in Alzheimer diseased synapses. Journal of Neuroscience, 37(5), pp.1197-1212.

Robinson, J.P., 2022. Flow cytometry: past and future. BioTechniques, 72(4), pp.159-169.