Flow cytometry is a high-throughput technology that analyzes individual cells suspended in a fluid stream. As cells pass through a laser, the system measures multiple physical and chemical characteristics simultaneously. This method allows for detailed investigation of specific cell populations within the complex environment of the central nervous system (CNS).
Microglia are the primary resident immune cells of the CNS, responsible for its immune surveillance and response. Combining the study of microglial cell biology with flow cytometric analysis enables researchers to quantify and characterize these cells with a high degree of precision. This approach helps in understanding the roles microglia play in maintaining brain homeostasis and in the progression of neurological diseases.
Applications of Microglia Flow Cytometry
Flow cytometry has several applications in the study of microglia. It is used to quantify microglial populations within specific brain regions, providing precise cell counts that can be compared across different experimental conditions. This is useful for studying conditions involving microglial proliferation or cell death, offering direct evidence of population changes.
A primary application is the detailed phenotyping of microglia to understand their functional state. Microglia are dynamic cells that adopt various activation profiles, such as pro-inflammatory or anti-inflammatory. By using antibodies against specific proteins, flow cytometry can identify and quantify the proportion of cells expressing markers associated with these different states, providing a nuanced view of the neuroinflammatory landscape.
This analytical approach is widely used in neuroscience to investigate the role of microglia in neurodegenerative disorders. In studies of Alzheimer’s disease, flow cytometry can track changes in microglial activation markers in response to amyloid-beta plaque deposition. Similarly, in models of Parkinson’s disease or multiple sclerosis, the technique is used to characterize the microglial response during disease progression.
The method is also applied in the preclinical assessment of new drugs. Researchers can use flow cytometry to determine if a candidate drug alters the number or activation state of microglia in a disease model. For example, it can assess whether a compound shifts microglia from a pro-inflammatory phenotype to a more protective, anti-inflammatory one, providing information about the drug’s mechanism of action.
Isolating Microglia from Brain Tissue
The initial step in performing flow cytometry on microglia is the isolation of a viable single-cell suspension from brain tissue. The process begins with cardiac perfusion of the animal with a cold saline solution, such as phosphate-buffered saline (PBS). This procedure flushes blood out of the brain’s vasculature, which prevents contamination of the final cell preparation with circulating blood cells.
Following perfusion and brain extraction, the tissue undergoes a two-part dissociation process. The brain is first mechanically minced into small pieces and then transferred into a solution containing enzymes that break down the extracellular matrix holding the cells together. This enzymatic digestion is performed at 37°C for a controlled period to ensure tissue dissociation without compromising cell health.
The resulting cell slurry contains a mixture of all CNS cell types and lipid-rich myelin debris. To enrich for microglia, this mixture is processed through density gradient centrifugation using a solution like Percoll, which can be layered to create different densities. The cell suspension is layered on top of a multi-density Percoll gradient.
When the gradient is centrifuged, the different components separate based on their buoyant density. Myelin and cellular debris collect at the top, while red blood cells and other dense debris form a pellet at the bottom. Microglia and other mononuclear cells collect at the interphase between the 70% and 37% Percoll layers. This layer is carefully collected, washed to remove the Percoll, and prepared for the next stage.
Throughout this isolation procedure, maintaining cell viability is a priority. All buffers and solutions are kept ice-cold, except during the brief enzymatic incubation, and centrifugation is performed at low temperatures. The final cell pellet is resuspended in a specialized buffer designed to support cell health until antibody staining and analysis.
Antibody Panels and Staining Protocols
Once a single-cell suspension is obtained, the cells are labeled with fluorescently-tagged antibodies for identification by the flow cytometer. This process relies on an antibody panel, which is a selected cocktail of different antibodies that target specific proteins, or markers, on or inside the microglia. Each antibody is conjugated to a unique fluorophore that emits light at a specific wavelength when excited by a laser, allowing for the simultaneous detection of multiple markers on a single cell.
The foundation of a microglia-focused panel involves antibodies that can distinguish them from other myeloid cells like infiltrating macrophages. Pan-myeloid markers, such as CD11b and CD45, are used, as microglia have a unique expression pattern of these markers. To achieve greater specificity, the panel includes antibodies against homeostatic microglia markers like TMEM119 and P2RY12, which are proteins predominantly expressed by microglia in a resting state.
To investigate the functional state of microglia, the antibody panel is expanded to include activation markers. These markers help characterize the phenotype of the cells. Common markers for a pro-inflammatory or reactive state include CD68 and CD86. Conversely, a marker associated with an anti-inflammatory or tissue-repair phenotype is CD206, the mannose receptor.
Building a multi-color antibody panel requires careful planning to avoid technical issues like spectral overlap, where the emission of one fluorophore spills into the detection channel of another. Researchers must select fluorophores with distinct emission peaks. The use of Fc block is also a standard part of the staining protocol. This reagent is added before the antibodies to block Fc receptors on the microglia, preventing non-specific binding and reducing background signal.
Gating Strategies and Data Analysis
After the stained cells are run through the flow cytometer, the data is analyzed through a process called gating. This is a sequential method of electronically selecting specific cell populations for further analysis based on their measured properties. The process begins with several cleanup steps to ensure the final analysis is performed only on high-quality, relevant cells.
- First, a gate is drawn on a plot of forward scatter (FSC) versus side scatter (SSC) to isolate cells from instrument noise and small debris.
- Next, cell clumps or doublets, which can be mistaken for single cells with higher fluorescence, are excluded by gating on a plot of forward scatter area (FSC-A) against forward scatter height (FSC-H).
- A viability dye is used to exclude dead cells, which can non-specifically bind antibodies. A gate is drawn to select only this live population for all subsequent analysis.
With the data cleaned of debris, doublets, and dead cells, the hierarchical gating strategy to identify microglia can begin. From the live, single-cell population, a plot of CD45 versus CD11b fluorescence is created. Microglia are identified by their expression pattern of being positive for CD11b but expressing a low to intermediate level of CD45 (CD11b+/CD45low).
In contrast, other myeloid cells like macrophages that may enter the brain from the blood express high levels of CD45 (CD11b+/CD45high). By drawing a gate around the distinct CD11b+/CD45low population, researchers can isolate the microglia for further characterization. Once this primary microglia gate is established, additional gates can be applied to analyze the expression of other markers within that population, such as quantifying the percentage of microglia positive for activation markers like CD68 or CD206.
Methodological Controls and Validation
To ensure that flow cytometry data is accurate and reproducible, several types of controls are necessary. For multi-color analysis, Fluorescence Minus One (FMO) controls are used. An FMO control is a sample stained with all the antibodies in the panel except for one. This is done for each fluorophore in the experiment. The purpose of an FMO control is to reveal the amount of fluorescence spread, or spillover, from all the other fluorophores into the empty channel. This information is used for accurately setting the gate between a negative and positive population.
Another component of validation is compensation. When using multiple fluorophores, the light emitted by one can bleed into the detector intended for another, a phenomenon known as spectral overlap. To correct for this, single-stain controls are run for each fluorophore in the panel. These controls provide a bright positive signal that the flow cytometer’s software can use to calculate the percentage of spectral overlap between channels. This calculation generates a compensation matrix, which is then applied to the experimental data to mathematically remove the spillover.
Isotype controls were historically used to estimate non-specific antibody binding. An isotype control is an antibody of the same class and fluorophore as the primary antibody but has no known specificity for any target in the sample. However, their use has become less favored because no two antibodies have the exact same non-specific binding characteristics. For setting gates in multi-color experiments, FMO controls are now considered the more appropriate and accurate tool.