Flow cytometry is a laboratory technique used to rapidly analyze the physical and chemical characteristics of individual cells suspended in a fluid stream. This method utilizes light, typically from one or more lasers, to measure multiple features of each cell as it passes through the instrument. The purpose of this test is to count, sort, and detect specific cellular components or biomarkers. It is broadly applied in diagnosing diseases like leukemia and monitoring immune function, providing a quantitative assessment of cell populations.
Preparing the Biological Sample
The test begins with meticulous preparation of the biological sample, which must be in a single-cell suspension. Samples such as peripheral blood or bone marrow are naturally suspended, but solid tissue must first be dissociated into individual cells. This disaggregation is accomplished through mechanical methods or enzymatic digestion, which breaks down the extracellular matrix.
The next step is immunolabeling, which involves tagging the cells with fluorescent markers. Specific antibodies are selected to bind to target molecules, often proteins, located on the cell surface or inside the cell. These antibodies are chemically linked to fluorophores, which are dyes that emit light when excited by a laser.
The process requires optimizing the concentration of cells, typically aiming for $10^5$ to $10^7$ cells per milliliter, to prevent clogging the instrument and ensure accurate analysis. The cells are incubated with a cocktail of these fluorescently tagged antibodies, often using several different colors to simultaneously analyze multiple targets. After incubation, the cells are washed multiple times to remove any unbound, excess antibodies.
Removing the unbound dye minimizes background noise and ensures that only specifically tagged cells fluoresce. If the target is an internal protein, the cells must be treated with a fixative and a permeabilization agent to allow the antibodies to pass through the cell membrane. The prepared, stained cells are then ready to be injected into the flow cytometer.
How the Flow Cytometer Works
The flow cytometer is comprised of three integrated systems: fluidics, optics, and electronics.
Fluidics System
The fluidics system precisely controls the movement of the cells through the machine. The cell sample is injected into a stream of a surrounding buffer solution, known as sheath fluid. This system uses hydrodynamic focusing, where the pressure of the sheath fluid compresses the sample stream. This action forces the cells to line up single-file in the center of the fluid stream, ensuring that only one cell passes through the laser beam at a time for accurate, cell-by-cell analysis.
Optics System
The optics system performs the physical measurements as the focused cells pass through the laser beam at the interrogation point. When a cell intercepts the laser, the incident light is scattered in all directions. Detectors positioned around this point collect the scattered and emitted light signals.
Light scatter is measured as Forward Scatter (FSC) and Side Scatter (SSC). FSC measures light scattered at a low angle, which correlates with the cell’s size. SSC is placed perpendicular to the laser and measures light scattered at a 90-degree angle, providing information about the cell’s internal complexity and granularity.
Simultaneously, the laser excites the fluorophores attached to the antibodies. These fluorophores emit light at specific wavelengths, which are collected by detectors, often photomultiplier tubes (PMTs). Optical filters and dichroic mirrors steer the different fluorescent wavelengths to their appropriate detectors. This allows the machine to distinguish between the various tagged markers on a single cell.
Understanding the Results
The electronics system converts the light collected from each cell into a digital signal, processing thousands of cells per second and generating a large dataset. This raw data is then visualized by a computer, typically in the form of dot plots and histograms.
A dot plot displays two measured parameters against each other, such as FSC versus SSC or the intensity of two fluorescent markers. Each dot represents a single cell, with its position determined by its measured characteristics. Histograms visualize a single parameter, showing the frequency of cells along an axis representing a characteristic like fluorescence intensity. These visualizations allow laboratory personnel to identify and isolate specific populations of interest.
The process of isolating a specific group of cells is called “gating.” This involves drawing a boundary around the cluster of dots representing the desired population on the scatter plot. The initial gate is often placed on the FSC versus SSC plot to separate different cell types, such as lymphocytes and monocytes, based on their relative size and granularity. Subsequent gates are applied to these isolated populations to identify cells that are positive or negative for specific fluorescent markers. This systematic gating strategy allows for the accurate counting of specific cell types, providing the quantitative data necessary for a diagnosis.