This method can help answer key questions in the field of T-cell immunology and development of viral vector vaccines. The main advantage of this method is that only small amounts of blood are needed, which allows repeated measurements from the same mouse. Additionally, virus and transgene-specific CTLs can be detected from the same sample.
Demonstrating the procedure will be Jasmine Rinnofner, a technician, and Annika Rossler, a grad student. Measuring and analyzing the samples will be Zoltan Banki, a post-doc. To begin, safely restrain a mouse according to the text protocol.
Collect 20 microliters of blood in an EDTA-coated tube from the tail vein of a mouse. When working with splenocytes, isolate the spleen and use the plunger of a syringe to press the tissue through a cell strainer. After performing lysis on the erythrocytes, count the cells and adjust the concentration of the sample.
Prepare and label one Facs tube for each sample, and transfer 100 microliters of organ suspension, or 20 microliters of blood to the tube. When working with splenocytes, centrifuge the organ suspension for five minutes, and discard the supernatant. Then, vortex the tube to resuspend the cell pellet.
For each channel to be used, also prepare a Facs tube for a compensation sample. Prepare one additional sample as an unstained control. Prepare a tube with Facs buffers with tetramers at their optimal dilutions, and vortex to mix.
Next, add 50 microliters of the tetramer dilution to each sample, and vortex to mix. Add Facs buffer without tetramers to the compensation controls and the unstained sample. Next, incubate the samples in dark conditions at 37 degrees Celsius for 20 minutes.
While the samples incubate, add Facs buffer to a tube. Then, add the antibodies to the buffer according to the dilutions specified in Table 2 of the text protocol, and vortex the solution. Pending on the scientific question are the marker combinations, apart from the one described here, might be used.
Make sure to always include antibodies against CD3 and CD8 in the panel. For each channel, prepare a tube with 200 microliters of Facs buffer, and add one microliters of an antibody dilution against CD8 in their respective color, and vortex to mix. Then, store the antibody dilutions as specified in the text protocol.
To wash the samples, add one mililiter of Facs buffer to the samples and centrifuge. Then, discard the supernatant, and drain any remaining liquid with paper towels. When working with blood, be cautious when draining off remaining liquid.
Prior to lysis of erythrocytes, the blood will not stick to the bottom of the tube. Alternatively, you can aspirate the supernatant. Next, add 50 microliters of the antibody solution to each cell pellet, and vortex gently.
Add 50 microliters of each compensation mix to the corresponding compensation control, and 50 microliters of Facs buffer to the cell pellet of the unstained control, and vortex gently. When working with splenocytes, wash the samples after antibody staining directly with one to two milliliters of Facs buffer, and centrifuge for five minutes. Then, discard the supernatant and drain off the remaining liquid on paper towels.
Add 500 microliters of ACK buffer to each blood sample, and vortex gently. Then, incubate the samples in the dark for five minutes at room temperature. Add one milliliter of Facs buffer to the samples, and centrifuge.
Then, discard the supernatant and drain any remaining fluid with paper towels. Proper lysis of erythrocytes is a crucial step to facilitate Facs measurement, therefore, when the pellet is rather wet, repeat lysis of erythrocytes. Wash the samples again as previously described.
Then, discard the supernatant and drain any remaining fluid on a paper towel. Prior to fixation, make sure that cells are well resuspended in order to prevent formation of clumps. Add 150 to 300 microliters of Facs fixing buffer to each tube, and vortex to mix.
Then, proceed with flow cytometric measurement as quickly as possible. First, measure the compensation controls and correct for any spectral overlaps. Afterwards, set up sequential gates to select for CD3-positive and CD8-positive cells.
Gate on the lymphocytes using forward and sideward scatter area. Then, within the lymphocyte population, gate on single cells using forward scatter width versus area. Use CD3 and CD8 channels to plot single-cell lymphocytes.
Identify CD8-positive T-cells, by gating on CD3-positive and CD8-positive cells. Make sure that CD8-low cells are included in the gating. Propagating on CD3/CD8-positive cells is a critical step in this protocol.
As activation of cells often leads to down regulation of CD3 and/or CD8, CD3/CD8-low cells should also be included. After this, plot the CD8-positive cells versus the tetramer cells, gating on the CD8-positive/tetramer-positive cells. If possible, record 20, 000 cells in the CD3-positive and CD8-positive gate for each sample.
Finally, save the measurements as a FCS file. In this protocol, antigen-specific CD8+T-cells were quantitatively detected in blood and tissue samples. After gating the CD3-positive and CD8-positive cells, tetramer-positive cells were identified.
Two different tetramers were combined in the same tube for staining, allowing for simultaneous quantification of two different CTL specificities. Using this protocol, T-cell responses from the same mouse can be followed over time, as only small amounts of blood are needed for each measurement. Additionally, phenotypes of antigen-specific CTLs can be analyzed.
While attempting this procedure, it is important to avoid prolonged incubation times, as this can lead to internalization of T-cell receptors. Since the discovery of tetramer technology, tetramer staining has become an essential tool for T-cell analysis, and a broad range of applications, which include tetramers accruing. A bright future for bright tetramers.
