The goal of our research is to understand T-cell migration patterns in highly defined environmental conditions to enable informed in vitro experimentation and decipher molecular interactions impacting migration. Live imaging such as intravital multiphoton microscopy is used for in vivo understanding of immune cell kinetics. But microfluidic devices are a common in vitro tool that offer precise control over the microenvironment that cannot be afforded in a live specimen.
Our protocol allows us to analyze intricate signaling pathways and provides better understanding for targeted in vitro experiments to further address the biological roles of individual signals in a cell type specific manner. Our method is simple, reproducible, and easy to access in research labs that have a bright field microscope featuring an attached digital camera. Therefore, our approach ensures a reliable set of methods to study dynamic cell migration events.
Our lab will focus on investigating the migration of immune cells, including T-cells and neutrophils, in a variety of disease settings, such as tumors and infection, thereby comprehending the mechanisms that are important to regulate immune cell migration. This will help in developing the therapeutic methods to help treat and prevent disease. To begin, place the euthanized mouse supine on the dissection board.
Using dissection T pins, secure the foot pads of the mouse to the board. With surgical dissection scissors incise the lower abdomen and cut up to the chin. After separating the skin from the peritoneal lining, stretch it perpendicularly away from the trunk and pin it down.
Employing blunt forceps, carefully remove cervical, axial, brachial, and inguinal lymph nodes as well as the spleen and place them in a 70-micrometer cell strainer containing 2 milliliters of R9 medium. To prepare a single cell suspension, twist the tissue disruption tool one half turn clockwise and counterclockwise repeatedly. Wash the strainer with 5 milliliters of media.
Then transfer the cell suspension to a 15-milliliter conical tube and centrifuge it. After decanting the supernatant, add 500 microliters of lysing buffer to the tube to remove red blood cells. After 1 minute, mix the cells with 9.5 milliliters of R9 medium, centrifuge the tube, and decant the supernatant thoroughly.
Next, re-suspend the cell pellet in 10 milliliters of negative selection medium containing anti-MHC II and anti-CD4 antibodies, and place the tube in a rocker for 30 minutes. After pelleting the cells at 270 G for 5 minutes, decant the supernatant. Simultaneously, in a 15-milliliter conical tube, wash 200 microliters of sheep Sheep Anti-Rat immunoglobulin G beads in 7 milliliters of medium.
Place the tube on a magnet and remove the medium. After washing two more times, re-suspend the beads in 7 milliliters of R9 medium. Then mix 7 milliliters of the bead suspension with the cell pellet and incubate the tube on a rocker.
To remove antibody bead-bound cells, place the conical tube directly on the magnet for 3 minutes. Aspirate the cell suspension into a new 15-milliliter tube, keeping the tube on the magnet. Pellet the enriched CD8-positive cells and wash three times in medium.
To begin, isolate live lymphocytes from mouse secondary lymphoid tissues and visualize the cells under a light microscope with a 4x or 10x objective. Using a 1 milliliter pipette, disrupt the activated cells and transfer them into a new 15-milliliter conical tube. Pellet the cells and add lymphocyte separation media to separate the live cells from the dead ones.
For time-lapse microscopy, plate 1 times 10 to the 5th CD8-positive T-cells in the medium in a 37 degree Celsius chamber. After performing integrin blocking, acquire brightfield and fluorescent images every 10-60 seconds for 10-60 minutes. In the time-lapse imaging, activated CD8-positive T-cells had increased average velocity, displacement, and meandering index compared to the naive cells.
To begin, perform time-lapse imaging of live T lymphocytes, isolated from neuron secondary lymphoid tissues. For T-cell migration analysis, open Velocity Software and create a new image sequence. Select and move all time lapse video microscopy movies to the blue area in Velocity.
Utilizing the contrast enhancement tool, modify the brightness and contrast. Then adjust the image sizes to 0.325 micrometers for 20x and 0.65 micrometers for 10x magnification. After configuring the sequence to set time points, go to the Measurement tab and uncheck all time points.
Next, find objects using intensity and slide the bar to the right of the peak. To avoid cell debris, exclude all cells smaller than 10 micrometers and greater than 100 micrometers in diameter. Check Ignore Static Objects followed by Automatically Join Broken Tracks.
To save a new protocol, go to Measurements, click Save Protocol, followed by Name New Protocol. In the Measurement tab, click on Measure All Time Points and sort tracks by time span from high to low. After recording the track ID numbers for the cells with good tracks, export the file as comma-separated text and transfer data to the desired analysis software.
To perform manual tracking, open ImageJ, go to Plugins and select Tracking and Manual Tracking. Set the time intervals, x/y calibration, pixel size, and z calibration. Next, click on Add Track to start individual cell tracking.
Select one cell at the first time point and continue it through all time points. Finally, click on End Track. Software-assisted cell tracking characterized migratory behaviors of activated individual T-cells, as denoted by the differently colored lines.
