Characterization of Immune Cells in Human Adipose Tissue by Using Flow Cytometry

The overall goal of this procedure is to use flow cytometry to isolate the stromal vascular fraction from human adipose tissue for phenotyping of the different immune cell populations found within the adipose tissue. This method can help answer key questions in metabolic research, such as which immune cells accumulate in the adipose tissue of obese individuals and to which extent. The main advantage of this technique is that it allows for simultaneous measurement of multiple markers on immune cells and the identification and quantification of these immune cells in adipose tissue.

Although this technique can provide crucial insight into adipose tissue inflammation, it can also be applied to other organs and tissues. Generally, individuals new to this technique will mainly struggle with stromal vascular cell contamination and loss of cells during the staining procedure. Two of my PhD candidates, Suzan Wetzels and Mitchell Bijnen, will be executing the protocol.

Begin by using a scalpel to mince one gram of adipose tissue biopsy into approximately two millimeters squared pieces. Transfer the pieces into a 50 milliliter centrifuge tube and add 10 milliliters of collagenase solution to the samples. Incubate the tissue fragments for 60 minutes in a 37 degree Celsius water bath at 60 cycles per minute.

Followed by filtration through a 200 micrometer strainer into a new 50 milliliter conical tube. Rinse the filter with seven milliliters of PBS before centrifugation. Then use a pipette to remove the supernatant without submerging the entire pipette tip into the solution.

The pellet should be clearly visible and remove nearly all supernatant to avoid contamination of the stromal vascular fraction with adipose cells. We suspend the stromal vascular cell pellet with five milliliters of fresh PBS and filter the stromal vascular fraction through a 70 micrometer strainer into a new tube. After centrifugation, we suspend the washed stromal vascular fraction pellet in three milliliters of erythrocyte lysis buffer for five minutes on ice.

Then arrest the lysis with seven milliliters of PBS and pellet the cells by centrifugation. To stain the stromal vascular fraction for flow cytometric analysis, resuspend the pellet in 90 microliters of four degree Celsius fluorescence activated cell sorting, or FACS buffer, and block the non-specific staining with ten microliters of human IgG. Split the cell suspension between two wells of a 96-well v-bottomed plate and place the cells on ice for 15 minutes.

At the end of the incubation, wash each sample with 100 microliters of FACS buffer, turning the plate upside down in one smooth motion without tapping to discard the supernatants at the end of the centrifugation. Make sure you can clearly see the pellet at the bottom of the plate before turning it upside down to prevent loss of your cells during the procedure. Resuspend the first pellet in 29.5 microliters of antibody cocktail for the first FACS panel and the second pellet in 23 microliters of antibody cocktail for the second FACS panel.

After 30 minutes on ice in the dark, wash the cells with 150 microliters of FACS buffer per well and fix the cell pellets in 150 microliters of one percent formaldehyde solution. Transfer the cell suspensions from each well into the appropriate corresponding FACS tubes and place the samples on ice. Before their analysis, load an unstained, negative control onto the flow cytometer to set the forward scatter and side scatter parameters.

Then adjust the voltages of the flow cytometer according to the manufacturer’s instructions until all of the populations of interest are visible in the forward scatter and side scatter graphs, and a distinction can be made between the debris and the live cells. When all of the parameters have been set, vortex the first experimental sample at 800 rpm to thoroughly resuspend the cells and load the sample onto the cytometer. Read a minimum of 50, 000 events in the live gate, then vortex and load the second sample for analysis as just demonstrated.

To allow analysis of the adipose tissue macrophage populations in this representative experiment, other immune cells, such as T cells, B cells, neutrophils and natural killer cells were excluded to reveal the presence of CD11b+CD11c+macrophages, CD11B+CD11C-macrophages, and CD11Blow-CD11C+dendritic cells. Quantification of the main fluorescence intensity of these cells revealed the expression of plasma cytoid dendritic cell and generic dendritic cell markers on the CD11B+CD11C+and CD11Blow-CD11C+cells, with a higher expression of both markers observed on the CD11Blow-CD11C+cells, confirming that the CD11Blow-CD11C+cells were in fact dendritic cells. Gating of the CD45 positive cells revealed CD19 positive B and CD3 positive T cell populations, the latter of which could be further subdivided into CD3+CD4+T helper, and CD3+CD8+cytotoxic T cell subsets, and CD3-CD56+natural killer cells.

The percentage of viable immune cells was then calculated for each subject, revealing a significant increase in the percentage of pro-inflammatory CD11B+CD11C+macrophages in the visceral adipose tissue of obese adult male subjects. Once mastered, this technique can be done within four hours if executed properly. While attempting this procedure, it’s important to keep the buffers and cell fractions on ice, and while labeling the cells with antibodies it’s very important to keep them in the dark.

After it’s development, this technique allowed for researchers in the field of metabolic research to explore the immune cell alterations in the adipose tissue of humans. After watching this video, you should have a good understanding of how to isolate adipose tissue immune cells and how to perform flow cytometry on these cells. Keep in mind that you’re working with potentially infectious human material and substances, such as formaldehyde, which can be dangerous.

Always take precaution wearing gloves and safety glasses.