Definition: Flow cytometry is a technique allowing for the examination of large numbers of single cells at high speed. The principle involved that cells can be passed within a sheath of solvent so that they pass a laser as individual units. The laser is employed to capture a scatter profile of the cells that gives information about the size and internal complexity of the cells and may also excite fluorescent probes that identify specific surface or internal structures.
Cytometers record information about each individual cell across a number of characteristics. To accomplish this, cells are titrated to run at a speed (measured by cells/second – often around 10,000 cells/sec) that is within the capacity of the machine to read. Physically, a constant flow of ‘sheath fluid’ is run across the detector’s path. The cell suspension runs as a separate stream within the sheath fluid.
Data points can be represented very clearly as values for each characteristic measured, and may be listed as a series of numbers as the table below. Here, cells were ‘labeled’ with antibodies against three known proteins, Btk, CD3, and CD19. Each antibody also carried fluorophores that emit known wavelengths of light when excited by (a) laser(s) of specific wavelength(s). These antibodies are illustrated in the figure to the side. Each type of antibody binds to a specific ‘antigen’ and carries several fluorophores that have been chemically linked to them (illustrated by different colored stars). Alternatively, secondary reagents can be used to bind to the primary antibodies to allow more freedom of color choice or to amplify weaker signals.
Cells are labeled or ‘stained’ with these antibodies by incubating cells with the antibodies for a period of time, followed by washes to remove excess, unbound antibodies. Typically, all stains can be done together in a single incubation unless secondary antibodies are employed to amplify weak signals or adjust the colors used or intracellular staining is required (see below).
If we measured data from each of the three cells above, this might be sufficient to illustrate the identity of each cell type without further analysis. However, if thousands of cells are measured for each condition in an experiment (done in triplicate), tables of numbers lose their value as effective illustrations of the data.
To account for this, scatter plots or density plots (similar to topographical maps) are regularly used to illustrate these larger datasets. Because it is only practical to present values in two dimensions at a time, plots are often drawn such that a population is identified in one plot and then those ‘gated’ cells are then redrawn in subsequent plots to illustrate values in new categories. Cells may also be examined for just one characteristic using a histogram.
Forward Scatter (FSC) and Side Scatter (SSC)
FSC and SSC are (very often) the primary measures of the physical properties of cells as they pass through the cytometer’s laser. FSC provides information about the size of the cell, while SSC provides information about the internal complexity of the cell. These data are presented for a sample dataset of white blood cells below. The more numerous Red Blood Cells (RBCs) and platelets have been eliminated prior to analysis.
The cells illustrated in the FSC / SSC plot above fall into identifiable subsets of white blood cells based on their size and complexity. The gated cells are known as lymphocytes, which includes both B and T Cells. Gating is a way of selecting a group of cells to analyze further.
Here, the lymphocyte population is now distinguished by the presence of identifying surface proteins, CD19 (found on B Cells) and CD3 (found on T Cells). By plotting the fluorescence emitted by antibodies to these receptors, we can not only identify the two major populations but gate each of them for further analysis for another protein, the intercellular protein kinase, Btk.
Looking at the Btk expression requires a slightly different technique because this protein is located inside the cell. For antibodies to access to Btk, we have to punch holes in the cell that let antibodies permeate cells. This is done chemically after all surface staining is complete and cells are ‘fixed.’ Otherwise, the protocol is very similar to surface binding.
In the last panel, both B Cells and T Cells (individually identified previously) are assessed for the presence of Btk and the results are represented as the number of events (cells) exhibiting high or low expression (illustrated below).
Here we can see that the B Cells express uniformly high levels of Btk, while T Cells express little or none. It would also be possible for us to see if only a subpopulation of either B or T cells expressed the kinase. In that case, we could gate expressers vs non-expressers to see if there are any other indications that these cells are different such as cell size or expression levels of the other receptors (CD19 or CD3).
It is possible to use staining to examine other features of the cells as well. For instance, if a treatment of cells might result in cell division, this can be tracked by using a non-toxic dye which is added to cells prior to treatment and then assessed afterward (typically 3-5 days). Because the dye is added only once, cells that divide will each take only half the quantity of the original dye. It is possible to distinguish up to 4-5 divisions clearly.
Data from these proliferation assays is often viewed in histograms to see the proportion of cells at each division, or with another label to see if the dividing cells up- or down-regulate certain receptors. It is also common to use a vitality dye that would demonstrate if cells that don’t divide die, vice versa, or exhibit some other pattern. The cells illustrated below are CD4 T Cells that were induced to divide by a ‘mitogen,’ possibly IL-2. The histograms depict cells in each generation, where the generation farthest to the right is the parent generation (i.e. undivided cells – this would be confirmed by a control population grown without mitogen). The next peak to the left represents cells that have divided once, the next represents cells that have divided twice, and so on. In quantitating the number of cells that have divided, it is important to consider that ONE parental cell is responsible for TWO cells that have divided once or FOUR cells that have divided twice, etc. (Note that the CellTrace dye is plotted along a log axis)
The Scatter plot illustrates the same cells, also plotted by cell division on the X- axis, however, this time the Y-axis separates cells according to their expression of CD4. These data show that the most actively divided cells are divided between CD4 expressors and non-expressers. We can also see that CD4 expression spikes in expressors upon division.
Fixing – chemically attaching antibodies to their targets in a reaction that kills the cells. This is required for longer term storage of cells and if further processes such as intracellular staining must be done.
Gating – Drawing a limit around a group of cells or an area where cells might appear for further analysis and/ or quantitation. Gating will always result in a calculation of the percentage of the total cells that are included within the gate
Labeling / Staining – to add fluorescent reagents to cells that will bind to specific elements.
Mitogen – a substance that induced cell division.
Washing – to repeatedly add a solvent to cells, spin to pellet the cells and remove unbound materials with the solvent.