High-Dimensionality Flow Cytometry for Immune Function Analysis of Dissected Implant Tissues

Isolation of cells from dissected implants and their characterization by flow cytometry can significantly contribute to understanding the pattern of immune response against implants. This paper describes a precise method for the isolation of cells from dissected implants and their staining for flow cytometric analysis.

This protocol can help us characterizing host immune response against different biomaterials which can subsequently assist in designing better future medical implants. Flow cytometry gives us information about immune cells infiltrating a biomaterial which helps us determine mechanisms of how cells respond to injury and material implantation as well as targets for improved therapeutics. The same protocol can be modified to characterize immune response in different settings.

Dissect the quad muscle of mice that received an ECM implant one week ago and collected in a 50 milliliter tube containing five milliliters of serum free media. Finally diced the tissue using scissors. Next, add five milliliters of digestive enzyme media to the tube.

Place the tube containing digestive media and diced tissues in a shaking incubator for 45 minutes at 37 degrees Celsius and 100 RPM. At the end of the incubation filter the digested tissue suspension through a 70 micron strainer into an individual 50 milliliter tube. Use a five milliliter syringe head to mash any solid chunk of tissues and wash the strainer with room temperature PBS.

Discard any remaining residue in the strainer and adjust the volume of the tube to 50 milliliters with room temperature PBS. Collect the cells by centrifugation and resuspend the pellets in 10 milliliters of cold five millimolar EDTA solution in PBS. If blood cells are present in the sample, resuspend the pellet in one milliliter of RBC lysis buffer, incubate for 10 minutes, then add nine milliliters of EDTA.

Leave the suspension on ice for 10 minutes. Then adjust the volume to 50 milliliters with cold PBS. Collect the cells by centrifugation and resuspend the pellets and 100 microliters of cold PBS.

Transfer 10 microliters of the suspension to individual micro centrifuge tubes and mix with 10 microliters of trip and blue for counting. Dispense the remaining volume of cells into each well of a V bottom 96 well plate. Remove 20 microliters of cells from the well and add them into a new well to use as unstained control.

Then use PBS to bring the final volume in each well to 200 microliters. Collect the cells at the bottom of the plate wells by centrifugation and resuspend the cells in 100 microliters of a one to 1000 concentration of viability dye per well. At the end of the incubation, wash each well with 100 microliters of fresh PBS and resuspend the pellets in 200 microliters of staining buffer per well.

After the second centrifugation, resuspend the cells in 50 microliters of one to 100 dilution of monocyte blocker. Incubate the cell suspension for five minutes on ice and then add 50 microliters of antibody cocktail. Incubate for 30 minutes on ice protected from light.

Then wash the wells with 100 microliters of staining buffer per well. Centrifuge the cells again and wash with 200 microliters of staining buffer. Repeat the process two more times followed by resuspending the cells in 400 microliters of staining buffer.

Before analyzing the sample, run an unstained sample to allow the cell population to be adjusted on a side scatter versus a forward scatter plot. Next, run the stained sample. Extract the autofluorescent signature.

Then import the FCS files to use them as a control for unmixing. Click on the unmixing icon to open the unmixing wizard and perform unmixing using all single color controls by gating the positive and negative populations. Next, click on the QC section and look at the complexity index.

The complexity index is a measure of how distinguishable a collection of spectral signatures is when the spectral signatures are unmixed together. Finally, unmixed the sample by clicking live unmix. Here, the results of a 14 color facts on control mouse tissue can be observed.

In this analysis, several lobbyist populations, such as ly6g positive neutrophils and ly6c medium and high expressing monocyte classes could be observed. CD11c high MHC two positive dendritic cells were readily apparent when gated against CD11b to rule out macrophages and other myeloid lineage cells such as neutrophils and monocytes. A subset of CD206 positive CD86 positive dendritic cells could be identified by focusing on this CD11c positive population, which includes both CD86 high M1 like dendritic cells and CD206 high M2 like dendritic cell populations.

F4/80 demonstrated a gradient of expression as is commonly observed in various macrophage populations. Siglec-F was present on both F4-80 positive and negative populations most likely corresponding to a macrophage subset and eosinophils respectively. Although staining with cell surface markers allows these designations of cell types to be made, it is important to note that cells expressing different markers should be viewed in a functional manner as opposed to a more binary classification.

When attempting this protocol, keep in mind that understanding the autofluorescent signature of your understained samples before designing your panel is key in achieving a better and mixing. Besides analyzing cells, the isolated cells can be sorted into specific populations for downstream evaluations with cellular assays, in vitro, transcriptomic analysis, or for microscopic analysis to evaluate cellular morphology. Increasing complexity of flow cytometry analyses of biomaterials helps bridge the gap between basic immunology studies and the engineering of new therapeutics.