Few methods allow to quickly assess the functionality of hematopoietic progenitors. This protocol measures the impact of the first cell divisions on cell fate and commitment. This protocol allows to measure simultaneously division and differentiation at single cell level and with high throughput without requiring complex and extensive manipulation.
This method is suitable to any hematopoietic population that can be maintained in culture. So it can be easily applied to cancer biology, immunology, and developmental biology. To begin, aliquot 250 microliters of the CD34+fraction equally in four 15 milliliters polypropylene tubes.
Label the tubes as described in the text manuscript. Aliquot 250 microliters of the CD34-fraction into another four 15 milliliter polypropylene tubes and label the tubes. Prepare CFSE high and CSFE low solutions as described in the text manuscript.
Add 250 microliters of the CFSE high solution to the CF and CV tube, 250 microliters of the CFSE low solution to the VC tube, and 250 microliters of Dulbecco's Modified Eagle Medium, or DMEM, without fetal bovine serum, or FBS, to the VI tube. To ensure an efficient mix of cell suspension in cell dye, incline the tube by 90 degrees and deposit the CFSE solutions on the tube wall. Then, hold the tube vertically and mix three or four times to ensure a fast mixing of the CFSE solutions with the resuspended cells.
Incubate the suspension at 37 degrees Celsius for eight minutes. After incubation, add five milliliters of DMEM containing 10%FBS and place the tubes at 37 degrees Celsius for five minutes. Spin the tubes at 300 G for five minutes.
Remove the supernatant via aspiration without disturbing the pellet and wash the pellet with five milliliters of PBS, ETDA. Spin the tubes again and discard the supernatant before resuspending the pellet in 40 microliters of PBS, ETDA. To stain the CD34+cells, prepare a master mix of antibodies into a single 0.5 milliliter tube.
Add seven microliters from the antibody master mix to each of the four CD34+conditions. Resuspend the cells in staining buffer using approximately 500 microliters each for the beads in CD34+cells and one milliliter for the CD34-tubes. For cell sorting, set the gating strategy by creating dot plot diagrams.
First, visualize the cells on an FSCA versus SSCA dot plot and double click on the Polygon gating tool to select the population with low side scatter. Label this newly created gate as Cells. In the following dot plot, FSCA versus FSCH, right-click on the plot and select the gate Cells from the drop-down menu by clicking on it.
Use the same gating tool to select a tight population on the diagonal between the two axes. Label this gate as Single Cells. In the third dot plot, APC versus FSH-H, display the population Single Cells and gate the cells negative for the expression of APC lineage.
Label this newly created population as Lineage Negative. In the fourth plot, CFSE versus CTV, display the population Lineage Negative and create four separate gates, one for each dye combination. Gate VI is shown here as an example.
Use the fifth and sixth plots to identify the progenitors of interest. Generously gate CD34+CD38-population and the CD34+CD38+in the fifth plot. Then, select the CD34+CD38-population in the sixth plot and draw three gates for LMPP, HSC, and MPP.
Run the CD34+fractions, recording at least 5, 000 events in the single cell gate. Adjust the gate for each dye combination, setting a tight gate to select a homogenous population. Prepare the Plate Sorting Template using the Experiment Sort layout.
The wells named CD34-contain 5, 000 to 10, 000 cells sorted on the CF, CV, VC, VI gate. The bulk wells contain 500 cells sorted on the gate CD34+CD38-The single cell wells contain only one event per cell division dye combination per well, so four events per well in total. Next, start with sorting the CD34-CF in Yield Purity mode.
Click on the Acquire button and then the Sort button. Click Acquire, then on the Sort button, ensuring to have ticked 0160 as purity grade. Finally, sort the cells of interest, one cell per well, in single cell purity, ensuring to have ticked the Index Sorting option.
Centrifuge the plate at 300 G for five minutes and remove the supernatant by rapidly inverting the plate under the hood over a paper towel. Add eight microliters of standing buffer to wells A1 to A4, followed by adding eight microliters of the mix to the other wells. Wash the cells by adding 100 microliters of staining buffer per well using a multi-channel pipette.
Centrifuge and remove the supernatant before resuspending the cells in 85 microliters of staining buffer. Start the analysis on the flow cytometer in Acquisition mode using the dedicated template and clicking on Custom. After selecting the fluorophores of interest from the list, set the plate setup following the plate template corrected for the number of wells containing at least one cell.
Tick the Agitation option and select 100 microliters as the acquisition volume limit. Then, set the acquisition rate not superior to one microliter per second, as lower speed improves the total volume analyzed per well. For representative gating, concatenate the different bulk wells in a single file.
After clicking on the Concatenate Populations option, select All uncompensated parameters from the Parameters menu, then click on Concatenate. Upload the concatenated file to the workspace, then apply the compensation matrix via drag and drop. Cells labels with CV and VC need a transformed value.
Click on Tools and Derive Parameter to get the transformed value. Paste the indicated formula into the formula box. Apply the gating to each color individually as a histogram plot.
For CF and VI, set CFSE-A and CTV-A on the X-axis, respectively. For CV and VC, set the newly derived parameter on the X-axis, then set gates corresponding to each peak. Apply the gating to each individual single cell well.
Ensure to add the derived parameter to every analyzed well. Manually verify each color gate for each well to detect events that are incorrectly assigned to a given peak. The flow cytometry analysis after cell culture requires precise gating to establish the kinship of each individual cell along with the cellular phenotyping.
Peak definition and assignment are crucial aspects of flow cytometry analysis. The histograms display two examples of families spread on multiple peaks, one of two similar intensity peaks and one with two different intensity peaks. Different types of data representation and statistical testing are shown here for two separate experiments performed after 72 hours for HSCs and MPPs.
The heat map for the condition HSC differentiation represents different cell families, both in cell fates and cell divisions. The histogram comparing the fate HSCs and MPPs derived cell families is shown here. For both cell types, the comparison between the condition GT and the condition differentiation is displayed together with the statistical tests performed for MPPs.
Histograms comparing the percentage of cell families per generation and the type of symmetry or asymmetry in fate for the first division are shown here. The distribution of fates per generation for MPPs and the condition differentiation are shown here. This procedure requires to isolate cells in bulk to set the appropriate gating strategy.
It is therefore better to start the staining with at least 10, 000 cells. Combining this method with continuous measurement, like live cells imagining, can be very powerful, as the two together can provide a detailed description of the early progenitors of cell dynamics. This approach was first developed for T cell biology by the Hopkins Labs.
This version, adapted for blood stem cells, is now expanding to cancer and stem cell biology.
