Method and Results Two common cell lines, U937 and Jurkat (available through ATCC), were used. Initial cultures for each line were seeded at both a high and low density, so that the final cell concentration at analysis would be dense or light, respectively. After one week of culture, cells were resuspended and 1 mL transferred directly from each tissue culture flask into a 1.5 mL tube. 7-Amino-actinomycin D (7-AAD, Cayman Chemical), a fluorescent dye which is excluded by viable cells, was used as a marker of cells with compromised outer membranes.
Five (5.0) µL of a 1 mg/mL stock 7-AAD solution, along with 50 µL of AccuCount Fluorescent Particles (Spherotech; ACFP-50-5) were added to each sample tube and mixed thoroughly. Tubes were kept at room temperature (RT) in the dark, and sampled between 5 and 30 minutes after addition of 7-AAD, with gentle mixing immediately prior to analysis.
A hemacytometer was used to perform the microscopic cell counts. Appropriate dilutions of cell samples were made into phosphate-buffered saline containing Trypan Blue. Counts were performed in triplicate. At least 100 nonblue cells were counted for each replicate.
With the C6, only a single 2-D density plot of forward light scatter (FSC-A) versus 7-AAD fluorescence (7-AAD FL3-A) was required for data analysis. A negative control sample for each cell type (no added 7-AAD) was used to define the viable-cell gate (Figure 2A, gate P5). This gate included events with high FSC-A, and defined the FL3-A background fluorescence. The dead-cell gate (Figure 2B, gate P6) was set using a cell sample containing 7-AAD. This gate only needs to be set once for each cell type within a given experiment.
Using the Statistics Tab in CFlow, the events/µL for these two gates were displayed for triplicate samples (Figure 2C). This data was copied and pasted into a spreadsheet program to determine the standard deviation (SD) and coefficient of variation (CV) for triplicate measurements (Figure 2D).
Small Particle Measurements
Method and Results Aliquots of whole blood (1 or 2 µL), collected in sodium citrate tubes, were diluted 1:10 into HEPES-buffered saline with 1% formaldehyde. Twenty (20) µL aliquots of diluted blood were incubated in 1.5 mL tubes at RT, 20 minutes, with 20 µL of CD41-PE antibody (DAKO clone 5B12). Samples were then diluted with 1 mL HEPES-buffered saline with 1% formaldehyde. Five (5) µL of RFP-50-5 beads (Spherotech) were added to allow comparison of two counting methods. Samples were well mixed and read, without washing, on the C6.
For this assay, data collection was triggered by the positive fluorescence signal of CD41-PE labeled platelets (read in FL2-H), in order to improve discrimination of platelets from debris and to increase counting accuracy. The appropriate FL2-H threshold channel was determined by first triggering on FSC-H, determining where CD41-PE+ events fell relative to the negatives, and setting this value (FL2-H channel=1,000) as the primary threshold in CFlow Plus (Figure 3A). All subsequent samples were collected with this threshold in place (Figure 3B). Platelet counts per µL of sample were copied from the CFlow Statistics Tab into a spreadsheet program, and dilution factors were applied to determine the platelets per µL of original whole-blood sample (Figure 3C).