Flow cytometry is a laser-based laboratory technique that analyzes the characteristics of individual cells or particles suspended in a fluid. This method rapidly measures physical and chemical properties of a cell population, providing detailed insights into cellular features. It serves a broad purpose, from counting cells to categorizing them based on specific properties.
This technology is used across various scientific and medical fields, including immunology, oncology, and microbiology. It helps researchers and clinicians identify and quantify different cell types within a mixed sample, assess cell health, and detect specific biomarkers. Its ability to analyze thousands of cells per second makes it a powerful tool for both fundamental research and clinical diagnostics.
Understanding the Basics
At its core, flow cytometry operates by suspending cells in a fluid and directing them through a narrow channel so they pass one by one through a laser beam. This process, known as hydrodynamic focusing, uses a surrounding sheath fluid to align the cells into a single stream. The precise alignment ensures that each cell is individually illuminated by the laser.
When a cell intercepts the laser, it scatters light in different directions, and any fluorescent molecules within or on the cell emit light. Detectors capture two main types of light signals: scattered light and fluorescent light. Forward scatter (FSC) measures the amount of light scattered in the forward direction, which generally correlates with the cell’s size. Side scatter (SSC) measures light scattered at a 90-degree angle, indicating the cell’s internal complexity or granularity.
Fluorescence signals come from specific dyes or antibodies that have been used to label cellular components. These fluorescent emissions are collected by additional detectors, each tuned to a specific wavelength of light. The intensity of these fluorescent signals reveals the presence and quantity of the labeled molecules. All these light signals are then converted into electronic signals, amplified, and processed by a computer, creating data points for each analyzed cell.
Preparing Your Samples
Successful flow cytometry begins with meticulous sample preparation, ensuring cells are in a single-cell suspension. For tissues or adherent cell cultures, this often involves enzymatic or mechanical dissociation to break them down into individual cells. Blood samples might require red blood cell lysis to isolate specific cell types.
Once a single-cell suspension is obtained, cells are typically stained with fluorescent reagents. These reagents often include fluorescent dyes or antibodies conjugated to fluorochromes, which bind to specific cellular targets like proteins on the cell surface or inside the cell. Direct staining uses an antibody directly conjugated to a fluorochrome, while indirect staining uses an unlabeled primary antibody followed by a fluorochrome-conjugated secondary antibody.
Proper controls are used for accurate data interpretation. Unstained cells help identify background autofluorescence, while single-stained controls set up compensation for spectral overlap between different fluorochromes. Isotype controls, antibodies with no specific target but matching the primary antibody’s class, help distinguish specific from non-specific binding. Reagents must be handled and stored carefully, typically in the dark and at low temperatures, to maintain stability and fluorescent properties.
Operating the Instrument
Before running samples, the flow cytometer requires proper setup and calibration. This involves powering on the instrument, checking the fluidics system for sufficient sheath fluid and waste capacity, and verifying clear sample lines. Many instruments perform an automated startup sequence to prime the system.
Daily quality control (QC) is performed using standardized beads with known fluorescent properties. Running these beads ensures consistent instrument performance over time, verifying laser alignment and detector sensitivity. This QC process allows for adjustments to maintain optimal performance and data comparability across experiments.
Setting instrument parameters involves adjusting the voltage or gain for each detector, controlling light signal amplification. This helps separate different cell populations effectively. When using multiple fluorochromes, compensation corrects for spectral overlap, where one fluorochrome’s emission spills into another’s detection channel. Samples are then loaded, and data acquisition begins, with the flow cytometer rapidly processing and recording information from each cell at rates typically ranging from hundreds to thousands per second.
Interpreting Your Results
After data acquisition, the raw data, often in a standardized .fcs file format, needs to be visualized and analyzed. Common methods for data visualization include dot plots and histograms. A histogram displays the distribution of a single parameter, such as fluorescence intensity, while a dot plot shows the relationship between two parameters, with each dot representing an individual cell.
Gating is a key process in flow cytometry data analysis, involving drawing regions or “gates” around specific populations of interest on these plots. This allows researchers to isolate and analyze subsets of cells based on their unique light scatter and fluorescence characteristics. For example, a gate might be drawn around a population of cells expressing a particular surface marker to quantify their percentage within the total sample.
Analysis involves quantifying characteristics of the gated populations, such as the percentage of cells within a gate or the mean fluorescence intensity (MFI) for a specific marker. MFI indicates the average amount of a particular molecule expressed by cells in that population. Occasionally, issues like sample clogs, poor staining, or instrument malfunctions can affect results, leading to unusual scatter patterns or dim fluorescence, which may require troubleshooting and re-running samples.