web monitor

Flow Cytometry Guide and Troubleshooting


 - Flow Cytometry
 - How a flow cytometer works
 - Sample preparation
 - Data analysis
 - Applications
 - No stain or weak signal detected
 - Non-specific staining (background)
 - Too much stain
 - Sample will not run
 - High side-scatter background
(Download PDF)


Flow Cytometry

Flow cytometry is a laboratory technique used to quantify cellular or particle properties based on the scattered incident laser light and the fluorescent emissions from a beam of light projected on a focused stream of fluid.  Particles that cause light to scatter range in size from a few microns to several hundreds of microns in diameter and are traveling as a stream of single particles at a rate of thousands per second.  This technique is commonly used by life science research laboratories measuring physical and chemical characteristics of cells, as well as by diagnostic laboratories measuring patient blood/tissue samples for health related disorders.  To analyze cellular/particle properties a number of detectors along with an aligned optics system of filters and lenses are used to collect scattered light from the point where the laser beam strikes the stream of sample fluid.  Detectors are then able to record the intensity of the light and transmit this information to a computer for further analyses using flow cytometry software.  This information can ultimately provide insight on cellular populations, size, inner-structure, or be used for sorting cells.

[back to top]


How a flow cytometer works:

The major components of a flow cytometer are the fluidics system, lasers, optics, detectors and computer for analysis.  The fluidics system is responsible for arranging the cells or particles in single file as they flow in front of the laser beam.  This is achieved through a technique called hydrodynamic focusing and is the result of an inner chamber of sample fluid and an outer (surrounding) chamber of sheath fluid.  The outer sheath fluid moves faster than the inner sample fluid and when the two fluids are introduced at the point of contact called the flow chamber the particles or cells within the inner sample fluid are aligned in single file.  There are two types of flow chambers: a flow cell and a nozzle tip.  The flow cell chamber is used in bench top cytometers and when the laser beam hits the sample fluid it is contained within this flow cell.  A nozzle tip flow chamber is used in stream-in-air cytometers and when the laser beam hits the sample fluid it is exposed to the open air.  

At the flow chamber the laser light is beamed on the sample and this point of interception generates two types of scattered light: forward-scattered light (FSC) and side-scattered light (SSC).  Forward scattered light is projected in the forward direction just off the axis of the incident laser beam and side-scattered light is projected roughly 90 degrees off of the laser beam.  Forward scattered light in general provides information on the size of the cell or particle, while side-scattered light provides information regarding the internal components of the cell and its granularity.

Once the light is scattered, a series of collecting optics are used to direct and filter the light as it’s targeted toward different detectors.  There are three types of filters: long pass, short pass and band pass.  Each has the ability to selectively filter light based on its wavelength.  For example, a band pass filter filtering light on its way to a detector setup for Fluorescein Isothiocyanate (FITC) would allow light between 515 nm and 545 nm to pass and wavelengths outside of this range would not pass.  Another example is R-Phycoerythrin (R-PE), which would use a band pass filter to allow light between 564 nm and 606 nm to pass.  Short pass and long pass filters are used to selectively filter wavelengths greater than or less than a specified wavelength.  It’s also sometimes necessary to have dichroic mirrors that can be used to reflect light, in addition to filtering light.  This is necessary to split light of shorter and longer wavelengths from the same signal to different detectors.  These are setup at a 45 degree angle to the incident light.  

After passing through the collecting optics the filtered wavelengths of light will come into contact with a detector.  Detectors are used to capture light signals and convert them to an electronic format for processing by a computer.  There are two types of detectors: photodiodes and photomultiplier tubes (PMTs).  Photodiodes are used to detect forward-scattered light, while photomultiplier tubes are used to detect side-scattered light.  The number of light photons is proportional to the intensity of the signal generated by the detector.

[back to top]

Sample Preparation:

There are numerous factors to consider when preparing your sample for flow cytometry.  These include the type of specimen, the physical and chemical properties of the targeted cell, the location of antigens within the cell (membrane, nucleus, cytoplasm, or organelle), and your downstream data analysis/application.  A range of specimens can be used including mammalian cell culture, blood and tissue, bacteria, yeast, fungi and plant cells.  To prepare these specimens for flow cytometry, the cells or particles of these specimens must be isolated in a suspension buffer that will maintain cells at an optimal density to avoid cell clumping and clogging of the flow chamber.

Generally, preparing samples from blood or cell culture (both adherent and non-adherent) is easier than preparing samples from tissue.  To isolate cells from tissue, a mechanical desegregation or enzymatic digestion procedure is necessary.  Mechanical methods are good because they can be done at cold temperatures and obtain cells from the entire tissue sample. Enzymatic digestion may only partially isolate your targeted cells leaving some cells of interest within the tissue.  Enzymatic procedures are usually done at room temperature, which can decrease cell viability.  Additionally, mechanical methods are faster than enzymatic, which require a digestion period that can take hours.  Overall, both methods have advantages and disadvantages, and the one that isolates and preserves your targeted cells the best is optimal.

If you are using probes to label specific antigens within a cell, you may also utilize a series of fixation, permeabilization, wash, centrifugation and/or incubation steps for immunological detection.  The steps you employ will depend on what antigen you are targeting and whether its located in the membrane, nucleus or cytoplasm of the cell. Fixing the sample may be necessary to stabilize the cell as alcohol and detergents used for permeabilization can cause the loss of antigens.  Common fixatives include acetone and formaldehyde.  While alcohol and detergents can cause the loss of antigen, they are sometimes necessary to expose antigens of the nucleus or cytoplasm for antibody-antigen detection.  For example, the detergent Triton-X can be used to remove the cell membrane and cytoplasm to leave only the nuclei, while milder detergents such as Saponin maintain the cell membrane in addition to exposing the nucleus for DNA detection.  Wash and centrifugation steps are used to enrich, or concentrate your cell population of interest.  Methods of enrichment include centrifugation, magnetic bead separation and density gradients.  Similar to immunohistochemistry and western blotting, indirect and direct labeling methods can be used for immunological detection.

[back to top]

Data Analysis:

As light hits the detector, the detector generates an electrical impulse based on the intensity of the light.  This intensity value or “event” is determined for each cell or particle that passes the interrogation point and is transmitted to a computer for storage.  As an industry standard, data is stored according to the flow cytometry standard (FCS) and is recognizable across multiple instrument/manufacturer platforms.  The type of flow cytometer and the number of detectors it contains will ultimately determine how many measurements you can record for each particle or cell in a given event.  For example, a flow cytometer setup with nine fluorescent detectors and two light detectors can simultaneously measure eleven values for each particle or cell.  The ability to simultaneously measure multiple parameters is what makes the flow cytometer such a powerful instrument for cell biology research and clinical laboratories.
Popular graphs to visualize flow cytometry data include histograms and dot plots.  Single-parameter histograms are usually composed of the parameter’s signal value (intensity of the signal) representing the x-axis and the number of events representing the y-axis.  Signals of equal intensity will be distributed in the same channel along the x-axis and as signals increase in intensity they will be positioned farther right along the x-axis.  It’s also feasible to compose an overlay-histogram in which two different samples measured against the same parameter are displayed on one histogram.  This can be used to verify a positive/negative identification between two samples.  A dot plot is a two-dimensional diagram that correlates intensities of two parameters for each event.  An example would be a diagram with the forward scattered light (cell or particle size) representing the x-axis and the side-scattered light (cell granularity or internal morphology) representing the y-axis.  Each individual event would be plotted within the dot plot and similar events that group together spatially within the diagram would also provide you information about the density level of cells or particles.  

Another important feature is the use of gates and regions to selectively obtain information or screen subpopulations of interest.  Creating a region is the process of circling the desired population subset within a histogram or dot plot for analyses.  This can be used to gather statistics on the circled subset or to create a gate.  Gates can be used as a screen to select a population subset for further analysis on another plot.  All data not selected will be ignored.  An example of using a gate would be circling a boundary around the lymphocytes in a SSC vs. FSC plot from a blood sample, but not including the monocytes and granulocytes. Another two-parameter histogram based only on those lymphocytes can then be created. Gates can also be used to eliminate results from unwanted particles such as cellular debris and clumps of cells.

[back to top]


One major application of flow cytometry is the ability to sort different types of cells from a heterogeneous cell population.  To separate cells they must be identified by their unique physical properties such as their size, shape and inner structure, or identified by stains/immunodetection.  Discriminating populations based on size and structure can be achieved by creating a dot plot of the FSC vs. SSC to identify cell populations, while stains/immunodection reagents are used to separate cells based on cellular markers.  After being identified, there are multiple ways to physically sort the cells.  These include a catcher tube that swings into the sample fluid stream to collect targeted cells or a system that uses vibration to induce each cell into a single fluid drop, which can then be charged and separated using an electric field to direct the cell into a collecting tube.  Sorting cells is useful when further functional analysis or applications such as PCR will be employed on a targeted population.  
Other applications include counting the number of cells in your sample, apoptosis analysis, cell cycle analysis and the isolation of stem cells.  Staining the cells for DNA detection can be used to understand what stage of the cell cycle your targeted cells are in.  It can also be used to understand how drugs or specific treatments are affecting your cells.  Common DNA-binding dyes include Propidium Iodide, DAPI and Mithramicin.  For apoptosis, it’s possible to analyze morphological and functional/biochemical changes that are characteristic of cells marked for self-destruction.  Often cells undergoing apoptosis will be smaller in size, have a condensed cytoplasm, and display nuclear fragmentation.  In addition, their DNA is likely to be broken strands.  Isolating stem cells can also be done using dyes to detect surface antigens and analyzing their cell cycle status.

[back to top]


 No stain or weak signal detected:

Problem Possible Solution(s)
Antibodies may not be compatible.
 Verify the appropriate conjugated secondary antibody is being used with primary antibody.
Expired antibodies or faded fluorescence.
 Verify proper storage conditions used and manufacturer's expiration dates.
Concentration of antibodies too low.
 Increase concentration and/or optimize multiple test dilutions.
Problem with laser alignment.
 Ensure it's properly aligned using positive control.
Antigen is not accessible.
 Modify permeabilization and/or fixation steps.  Also, verify in literature or through the use of a positive control the antigen is expressed in the desired sample type.
Gain may be set too low.
 Optimize sensitivity/signal detection using a positive control.

[back to top]

 Non-specific staining (background):

Problem Possible Solution(s)
Could be due to autofluorescence.
 Use a negative control to check and minimize bacterial contamination, which may cause autofluorescence.
Excess antibody used.
 Ensure proper wash steps used to minimize trapped/unbound antibody.  Optimize antibody titration.
Secondary antibody cross-reacting with cells.
 Choose different secondary antibody.

[back to top]

Too much stain:

Problem Possible Solution(s)
Concentration of antibodies to high.
 Decrease concentration and/or optimize using multiple test dilutions.

[back to top]

Sample will not run:

Problem Possible Solution(s)                                                  
Clumped cell or particles.
 Filter sample.
Blocked flow chamber.
 Clean system.
Waste container has insufficient airflow causing increased pressure.
 Loosen cap.

[back to top]

High side-scattered background:

Problem Possible Solution(s)
Cells have been lysed or broken into small particles. 
 Use freshly prepared cells, optimize buffer system and/or centrifugation/wash steps during enrichment.

[back to top]


1. Shapiro HM. Practical Flow Cytometry. 4th ed. New York: Wiley-Liss; 2003.

2. Watson JV. Introduction to Flow Cytometry, First Paperback Edition.  Cambridge University Press; 2004.

3. Ormerod MG. Flow Cytometry: A Practical Approach, 3rd Edition. Oxford University Press (2000).

[back to top]