This article discusses in vivo brain cancer imaging techniques using fluorescently-labeled glioma cells in mouse models. The methodology involves the use of lentiviral vectors to express infrared fluorescent proteins, enabling deep tissue imaging of tumor development.
In vivo brain cancer imaging involves injecting fluorescently-labeled cancer cells into the mouse brain to observe tumor development. For fluorescent labeling, incubate glioma cells with lentivirus particles. The lentiviral vector carries the gene for infrared fluorescent protein, iRFP.
The virus binds to the glioma cell surface and releases RNA into the cell. Cellular enzymes convert the viral RNA to double-stranded DNA, which integrates into the cellular genome and allows iRFP expression. iRFP's excitation and emission maxima lie within a near-infrared region where tissues have minimal absorbance and scattering. This allows accurate deep tissue imaging.
Next, remove the spent medium and add trypsin. Trypsin, a protease enzyme, digests the adhesion proteins, detaching the cells from the flask surface. Pipette repeatedly to separate the individual cells. Collect the cell suspension into a tube and centrifuge. Now, add a suitable buffer to resuspend the pellet.
Dispense the cell suspension into flow cytometry tubes and stain the cells with a fluorescent dye, DAPI. DAPI stains the nucleus of dead cells bright blue, distinguishing them from live cells. Using a fluorescence-activated cell sorter or FACS, sort the live iRFP positive cells from other cell populations based on their fluorescence. The isolated cells can be used for injection into mice for tumor imaging.
Begin by seeding 1 times 10 to the sixth glioblastoma cells in 5 milliliters of medium into a 10-centimeter dish for a 24-hour incubation in a cell culture incubator. The next morning, transduce the cells with lentivirus expressing an appropriate fluorescent protein of interest at a multiplicity of infection of 5 and return the cells to the incubator.
After 24 hours, replace the supernatant with 5 milliliters of fresh medium and incubate the culture for another 48 hours. At the end of the incubation, replace the medium with 3 to 5 milliliters of trypsin for 10 to 15 minutes at 37 degrees Celsius, followed by careful pipetting to fully dissociate the detached cells.
Collect the cells by centrifugation and resuspend the pellet in 500 microliters of sorting solution. Split the cells into the appropriate number of FACS tubes and costain the cells with DAPI for dead cell exclusion. Then, load the tubes onto the flow cytometer and sort the live cells according to their fluorescent construct expression into a 15-milliliter conical tube containing fresh sorting solution.
After centrifugation, resuspend the construct-positive/DAPI-negative cells in 5 milliliters of culture medium and seed them onto a new 10-centimeter dish for their culture at 37 degrees Celsius for 48 to 72 hours. At the end of the incubation, split the cells for seeding into multiple dishes for subsequent in vivo experiments.
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