The overall goal of this method is to use cell tracking dyes to directly quantify the extent of cell division for different immune cell subsets within complex populations. This is achieved by first labeling the parent cell population with a fluorescent dye known to give stable binding to cellular proteins or to intercalate non covalently into cellular membranes for tracking dye labeled cells that respond to stimulation counters, staining with fluorescent antibodies and analysis using multi-parameter flow cytometry allows detection of differential responses among the immune cell types of interest stimulated, but tracking dye unstained controls are used to establish the lowest daughter cell fluorescence intensity that can be distinguished from autofluorescence tracking dye labeled, but unstimulated controls are then used to establish the fluorescence intensity at which the undivided cells will be detected. Stimulation typically results in a broad range of intensities as cells proliferating in response become progressively dimmer, but non-responders do not.
Ultimately, the use of peak modeling software to estimate the relative frequency of the cells in different generations can allow quantitative comparison of the proliferative responses across different populations or stimuli using metrics such as proliferation index or precursor frequency. The main advantage of this technique over existing methods like tritiated, thy incorporation, or KI 67 expression, is that the DI dilution provides a direct measure of cell division that can be, can be combined with other ME flow cytometric measures like immunophenotyping and cytokine production. Visual demonstration of the method of membrane dye staining is helpful as dye uptake by cells is extremely rapid.
Consequently, good technique is the key to obtaining bright homogenous staining. After human peripheral blood mononuclear cells are isolated centrifuge for five minutes at 300 times G and room temperature to minimize platelet contamination. Then resuspend the pellet at 10 to the seventh cells per milliliter in HBSS plus 1%BSA, and place them on ice.
Remove and reserve a 500 microliter Eloqua, then transfer another 500 microliter aliquot of cells into a 12 by 75 millimeter conical polypropylene tube. Add 3.5 milliliters of HBSS and spin down the cells for five minutes at 400 times G and room temperature during the wash add 0.5 milliliters of diluent C labeling vehicle from the PKH 26 GL kit to another 12 by 75 milliliter conical polypropylene tube carefully aspirate the super natin leaving no more than 15 to 25 microliters of residual fluid and taking care not to remove any cells. Next, add 0.5 milliliters of diluent C labeling vehicle to the cell pellet and aspirate and dispense the solution several times to obtain a single cell suspension.
Now prepare the two XD stock by adding two microliters of PKH 26 to the tube containing diluent C.Only now immediately pipette the cell suspension into freshly prepared two XPKH 26 dye solution and simultaneously aspirate and dispense the admixture several times to uniformly disperse the cells throughout the dye. After one minute, add one milliliter of heat inactivated serum to stop uptake of the dye into the cell membranes. After spinning down the cells carefully aspirate the supernatant without removing the pellet.
Then disperse the pellet in four milliliters of complete medium containing 10%heat inactivated serum, and transfer the cell suspension to a fresh polypropylene tube. Wash the cells twice in HBSS plus 1%BSA adequately stained cells will exhibit a distinct pink tinge in the pellet After resus suspending the cell pellet in one milliliter HBSS plus 1%BSA, count the cells and then adjust the volume to give a final concentration of 10 to the seventh cells per milliliter. After staining the cells according to the flow cytometric protocol outlined in the table, use the first five tubes to set the forward scatter and side scatter parameters and the signals in all four fluorescence detectors.
In particular for the detector used to monitor the PKH 26 fluorescence in tube two. Verify that all PKH 26 stained cells appear on the scale as a single symmetrical peak in the third to fourth decade with few to no cells in the last channel. Next, use the color compensation software and the list mode files collected for tubes one to five to establish a color overlap matrix for each fluro in the detectors being used to monitor the three other fluorochromes.
Then apply this matrix to the list mode file for sample six. To verify that the presence of PKH 26 labeling does not alter the ability to detect seven A a D positive cells. Now apply this just established color overlap matrix to the list mode file for tube seven.
Identify the CD three positive CD four negative and CD three positive CD four positive populations on a FE versus a PC plot. Confirm that the presence of anti CD three, FE and anti CD four A PC does not alter the ability to detect seven A a D positive cells. Finally, apply the color overlap matrix for tube seven to the list mode File for tube eight.
Now open a program containing a proliferation analysis module. Load the stimulated PKH 26 stained file from the dataset to be analyzed. Next, select the parameters for analysis in this case, PKH 26 gated on viable seven a a d negative cd, three positive lymphocytes and forward scatter versus side scatter for exclusion of small debris and large aggregates.
In defining these regions, be careful to include the high forward scatter area where blasts are typically found. Next, open the proliferation wizard. Click the start tab to open the data file and then load the unstimulated PKH 26 positive control.
Define the location of the parental peak corresponding to undivided cells. Analyze the file noting the values for parental peak position and width. If a fixed peak width is desired, check lock SD load the P KH 26 negative control and adjust the number of generations by setting the peak channel for the dimus daughter generation above the PKH 26 negative cells.
This sets the number of daughter generations. The model can accurately fit when completed. Reload the stimulated PKH 26 positive file.
If distinguishable generational peaks are evident, choose the floating setting under model option. If log decades are not exactly 4.0, adjust the generational spacing as discussed in the article. Now, analyze the stimulated sample and confirm that the region for the parental peak position remains unchanged.
Finally, select analyze for each experimental file in the dataset to apply. The model just created and record the desired proliferation metrics resulting from the best fit for each experimental file in the dataset. This first series of data illustrates the effect of the mixing technique on PKH 26 fluorescence distributions.
When cultured, U 9 37 myeloid cells were stained with the cell tracking dye using the method just described this first histogram shows the unstained control data. The cytometer settings were adjusted to place the autofluorescence from these cells in the first decade with no or very few cells accumulating in the first channel. This second histogram illustrates good PKH 26 staining.
Rapid mixing of two x cells with two x dye gave a fluorescence distribution that was bright, homogeneous and symmetrical, but had no cells off scale at the instrument settings used for the previous histogram. When two x cells were added to two x dye without immediate mixing, a more heterogeneous fluorescence distribution was obtained. This small subpopulation of dimly labeled cells would be erroneously interpreted as daughter cells if they were present at a later time point in the final histogram from this experiment, poor mixing also occurred when three microliters of concentrated ethanol dye stock was accidentally added directly to 0.5 milliliters of two x cell suspension yielding an extremely dim and right skewed fluorescence intensity distribution.
Here typical results from a proliferation experiment in which PKH 20 stained human peripheral blood mononuclear cells were cultured with or without stimulatory. Anti CD three and IL two reagents are shown. After 96 hours of culture, the cells were harvested and counterstain with anti CD three FE anti CD 19 A PC and seven A a d.
Two different gating strategies were compared in the first gating strategy, A dot plot of CD three versus seven. A A D was used to select live seven a a d negative CD three positive T cells. A forward and side scatter region was then used to gate out large aggregates while still including blasting lymphocytes.
The R one plus R two gated events for the unstained control are represented by the gray histogram and the PKH 26 stained, but unstimulated cells are in blue note, the bright symmetrical homogenous staining for viable but undivided T cells in the unstimulated control. These data were gated in the same manner as the previous figure, but in this case, the cells were stimulated with anti CD three and IL two. The PKH 26 histogram for R one plus R two gated events now contains mostly divided cells with reduced intensity represented by colored peaks for each daughter generation.
Although some bright undivided cells still remain in the blue parental peak. Note that the intensity of the highly divided cells does not yet overlap with the region where unstained cells are measured, which would confound the estimation of metrics based on number of cells in the lowest intensity daughter generations here. The same data were analyzed as the previous two sets of figures, but only light scatter and CD three were used to select for viable T cells.
This gating strategy excludes many, but not all of the dead cells as demonstrated by the small residual population of seven A a D positive dead cells remaining in the CD three versus seven A A D dot plots. The choice of fluorochromes used for immunophenotyping can create challenging compensation problems when using cell tracking dies as the staining intensity of undivided cells is extremely bright. In this experiment, 24 hour old peripheral blood mononuclear cells were labeled with PKH 26 and then counterstain with antibodies to CD three and CD four plus a viability dye.
In this series of data, the cells labeled with two micromolar PKH 26 were counter stain with anti CD three FZ seven A a D, and anti CD four A PC, and then analyzed using the same R one plus R two gating strategy as before, because there is little overlap of PKH 26 into the A PC channel, CD four positive and negative events were clearly resolved whether or not compensation was applied. These data illustrate how the viable CD three positive CD four positive T cells remained well resolved even when the final PKH 26 concentration was increased to four micromolar. The use of anti CD four per CP in combination with anti CD three FE two micromolar PKH 26 and the viability die to pro three gave much poorer results due to significant overlap of PKH 26 into the per CP channel.
CD four positive and negative cells could not be resolved from each other in uncompensated data and were only marginally resolved when compensation was applied. Increasing the final PKH 26 concentration to four micromolar resulted in a complete loss of the ability to resolve CD four positive from CD four negative events even after compensation was applied. Note how the samples including or lacking anti CD four per CP were essentially identical.
When a dye dilution profile is analyzed using peak modeling software to estimate the frequency of cells in successive daughter generations, positions and or widths of successive daughter peaks can be fixed or floating. With respect to values for the parental peak. These values are estimated from the unstimulated control.
For example, this PKH 26 intensity profile is from an unstimulated 96 hour culture from a moderate responder and was used to provide the mod fit proliferation wizard with the first estimate of position and width for the peak representing undivided parental cells. Use of a fixed setting for peak position requires the mean for each generational peak to be exactly half the fluorescence intensity of the previous peak. A float setting for peak position allows final positions of the generational peaks to vary from the expected intensities as needed to obtain the best fit.
Similarly, a fixed setting for peak width requires the width of each generational peak to be the same as that of the parental or unstimulated control peak. A float setting for peak width allows final widths to be independently varied as needed to obtain the best fit in this table. Results of the analyses of the previous figures for two different donors are shown for these donors where distinguishable generational peaks were evident.
The best reduced chi square values were obtained when both the peak position and the width were allowed to float for proliferation profiles where distinguishable generational peaks are not evident, a fixed setting for peak position is recommended. These data illustrate the use of two tracking dyes to separately monitor the proliferation histories of different immune cell types in a flow cytometric immune suppression assay. T effector cells shown in green were labeled with CFSE and T regulatory cells shown in blue were labeled with cell view.
Claret labeled cell populations were admixed in varying ratios and cultured in the presence of anti CD three, anti CD 28, and irradiated accessory cells for 96 hours. Thereafter, cells were harvested and stained with anti CD four, PE size seven and live dead violet.Here. Representative data for a treg to T effector ratio of 0.25 to one are shown after gating out live dead violet positive cells.
A light scatter gate that included lymphocytes and blasts, but excluded aggregates and debris was applied, followed by a gate to include CD four positive events. Cell view claret staining allowed viable treg cells to be easily distinguished from viable T effector cells that were highly proliferated and therefore CFSE DIM proliferation profiles for CFSE positive T effector cells and cell view Claret positive treg cells were analyzed using mod fit peak modeling software. Approximately 25, 000 events were analyzed.
Of these 48%were viable T effectors 6%were viable T regs with the rest being dead cells, accessory cells, and debris. The calculated proliferation index, which reflects the fold increase in cell number during the assay period was 3.85 for T effectors and 1.83 for Tregs. Here, the effect of the tregs cell concentration on the proliferation index of T effector cells calculated from the CFSE dye dilution profile is shown.
The results were the same, whether the treg cells were labeled with cell view claret or were left unstained indicating that treg function was not altered by staining. With tracking as expected, the extent of tector proliferation increased as the concentration of treg cells was decreased. Treg cell proliferation indices were also calculated from cell view claret dye dilution profiles at different treg to T effector ratios.
As expected, the Tregs underwent minimal proliferation in the absence of T effector cells as T effector cell concentration increased. However, the extent of the treg cell proliferation also increased. After watching the video, you should have a good understanding of how to successfully use cell tracking dyes to monitor cell proliferation, how to choose appropriate fluorochromes and color compensation controls, and how to select an appropriate model for quantifying cell division based on dye dilution.
Along with this procedure, other methods can be performed to answer experimental questions such as staining antigen specific T cells or flow sorting for recovery of stem-like cells that are quiescent or slowly dividing. Thanks very much for watching.
