Perhaps the most common, and indeed most famous, use for flow cytometry is in immunophenotyping . This is used by research scientists to defi ne populations of cells, but also used by clinical scientists to identify immunodeficiencies and genetic disorders. One of the first uses of flow cytometry was during the 1980s when the number of cases of acquired immunodeficiency syndrome ( AIDS) was rapidly increasing. In this context, flow cytometry was used to assess lymphocyte depletion in AIDS patients.

In this section, we will use an example of simple immunophenotyping of one of the most commonly investigated tissues – human blood – to understand the basic protocols involved in flow cytometry, as well as the practicalities and limitations of this technique. Sample preparation for flow cytometric analysis can be very straight forward, but in some cases may also involve highly specialised tissue preparation protocols. Examining cell populations in human blood is considered a simple protocol and the basic outline is shown in Table 1.

Table1. General protocol for preparation of human blood for flow cytometry

Sample Processing

 One of the most important aspects of acquiring good flow cytometry data for immunophenotyping is the quality of the sample. Typically, it is best to process the sample as rapidly as possible following isolation of cells. Extracted cells should be kept on ice at all times during the staining procedure.

The tissue of interest then needs to be prepared so as to generate a suspension containing only a single cell type. This can be straightforward and involve simply lysing red blood cells in human blood or specialised protocols where physical disruption or enzymatic digestion is employed to break down the structure of a tissue, generating a single-cell suspension. This sample preparation step is important, not only for cells to be analysed in isolation by flow cytometry, but also to ensure the sample is homogeneous and the absence of interference with the normal fluidics flow of the flow cytometer. Human blood is usually taken in tubes containing EDTA or heparin to prevent clotting. Typically, the red blood cells are lysed (using water or ammonium-chloride–potassium buffer) as only white blood cells are of interest for phenotyping. If one is interested in examining a specific type of cell, then extracted cells can be separated by density-gradient centrifugation prior to investigation by flow cytometry. In that case, there is no need for red blood cell lysis as these will be removed by the preceding separation step. Frequently, such a separation step is used to separate mononuclear cells (termed peripheral blood mononuclear cells or PBMCs) from granular cells such as neutrophils and eosinophils.

Once a single-cell suspension has been achieved, cell samples are first incubated with blocking agents, such as Fc block (also known as anti-CD16/32), mouse and/ or human serum. These blocking agents are incubated with the sample to prevent non-specific binding of antibodies to cells that express surface receptors capable of binding to antibodies, even if they are not expressing the surface marker of interest, as is the case in macrophages and neutrophils. Another way of thinking about this is that these cells are sticky and without Fc block or serum are likely to have high levels of background binding. Following this incubation, cells are stained for specific surface markers alongside a marker such as 4′,6-diamidino-2-phenylindole (DAPI) that will stain only dead cells in the preparation and hence allow for discrimination between live and dead cells. This is particularly important when examining immune cells isolated from tissue preparations, where cell viability can be influenced by the tissue processing protocol.

Gating

 In flow cytometry, the discrimination of a particular cell population from all cells present is called gating. A gated population has been purposefully selected by the person acquiring or analysing the data. The expression of a specific marker is then analysed only for a certain cell population rather than all cell types present in a sample. This selection not only ensures efficient analysis, but also increases the chances of identifying potential variations in marker expression.

When acquiring data on a sample, there are a few important points to bear in mind to ensure the validity of the data:

• The sample should not be processed through the cytometer too fast. This can be determined by the flow rate or events per second that the instrument is able to detect. Typically, one should not exceed 10 000 events per second, as otherwise the instrument may not be able to read all the events and discard some of them.

• The number of events per second should be stable and remain reasonably constant throughout data acquisition. If this is not the case, either the fluidics of the instrument are blocked or the sample is not of sufficient homogeneity.

First, it is common to use the ratio of forward- and side-scatter parameters to either gate on cell populations of interest or to exclude very small cells, such as remaining red blood cells or lysed cellular debris. Subsequently, the ratio of forward-scatter height to area is employed to identify and remove any events that do not indicate the presence of single cells. This gate is sometimes called the singlet gate and will remove any cells that are stuck together. Examining cells based on forward and side scatter can inform about the cell populations within the sample (Figure 1a). Certain populations can be distinguished based upon this analysis, which provides an approximation of cellular morphology based on size and internal structures of the cell. Although this should never be used as a definitive way to identify cell populations, it is a helpful indicator.

Fig1. Assessment of forward- and side-scatter profile, as well as live cell discrimination via flow cytometry. (a) The correlation between forward and side scatter can be used to provide an estimation of cellular morphology and size. The plot shows forward- against side-scatter intensity on whole human blood after red blood cell lysis. Multiple cell populations can be distinguished based on these simple parameters. (b) The plot shows side-scatter intensity against a dead cell marker, in this case a fixable UV dye. Live and dead cell populations are indicated.

In a second discrimination, any dead cells are gated out based on the dead cell marker to ensure that only viable cells will be analysed ( Figure 1b ).

After identifying the viable cell populations by these preliminary analyses, the population of interest can be gated on, for example the CD14+CD16+ blood monocytes (a specific population of white blood cells), as illustrated in Figure2.

Fig2. Gating strategy for human blood monocytes. The plots show a sequential gating strategy to identify human blood monocytes by flow cytometry.

Problems of Spectral Overlap and Colour Compensation

a major problem faced when undertaking flow cytometry experiments is the degree of spectral overlap between the emission spectra of different fluorescent dyes (e.g. FITC and PE). In some cases, this can be taken care of at the stage of designing the experiment by carefully choosing the antibody panel employed to stain the sample such that the spectral overlap is minimised. However, in order to take advantage of the ability of flow cytometry to analyse multiple labels at once, it may not be possible to avoid spectral overlap. Indeed, panels employed to stain samples for flow cytometry can utilise up to 18 different fluorescent dyes conjugated to antibodies. The more fluorophores are included in a panel, the more complicated spectral overlap can become. Figure3 highlights the overlaps between the most commonly employed fluorescent colours in flow cytometry.

Fig3. Comparison of the emission spectra for commonly used fluorescent dyes illustrates the spectral overlaps. The emission profiles represent the probability a photon will be emitted following excitation of the dye by a specific wavelength of light.

It is possible to correct for spectral overlap in a process called colour compensation when processing the data. The correction to reduce ‘spill over’ from one fluorescence channel to another may be done manually or computationally using compensation tools included with the flow cytometry software. One common way to do this involves a control experiment with compensation beads where each fluorophore is bound to an individual aliquot of beads and their fluorescence data are acquired in separate runs so that the spectrum of each fluorescence label can be established in isolation. Examples of colour compensation are illustrated in Figure4, showing a correlation of fluorescence intensity from two fluorescence labels (FITC and PE) that are under- and over-compensated (too much of the emission from one channel is bleeding into another), as well as correctly adjusted. Clearly, correct colour compensation improves the accuracy of any results obtained.

Fig4. Illustration of colour compensation due to spectral overlap of FITC and PE emission spectra. The plots show PE under-compensated (left), over-compensated (centre) and correctly compensated (right) in the FITC channel.

Fluorescence Minus One (FMO) Controls

In order to address uncertainties in multi-colour flow cytometry as to whether a particular stain results from a true positive stain or spill over from another channel (i.e. inappropriate colour compensation), a strategy of control experiments is typically applied. Given that completely unstained cells have a very different fluorescence pro fi le to cells that have been stained with multiple reagents, the current best control is to stain with all of the labels employed except for the one of interest. This control is termed a fluorescence minus one, or FMO, control. For example, in a fi ve-colour staining panel, five FMO controls should be generated in each of which four labels are deployed, but a different one omitted each time (see Figure5). This allows for a rigorous determination of truly positive staining on a sample.

Fig5. Fluorescence minus one (FMO) staining control. The tables outline an example staining panel and the FMOs that should be employed. The comparison of two flow cytometry plots illustrates the results of one such FMO control. The top plot shows staining on a sample for all colours except FITC. The fluorescence in the FITC channel (plotted on the x -axis) in the absence of an FITC-labelled antibody indicates the background staining and therefore where the positive gate should be drawn. The bottom plot shows a sample in which all colours have been stained, including FITC. The numbers indicate the frequency of cells staining positive for FITC.

Back Gating

 Back gating is a useful technique to validate whether a specific marker can be employed to look at a defined cell population. Practically, the sample is gated on the marker of interest and the resulting forward/side-scatter profile is assessed in order to find the location of the immune cell population on the plot. This approach ascertains whether the stained cell population is homogeneous. As such, back gating is also a good quality control measure.

Analysis and Quantification

Depending on the number of labels used for an experiment, it is possible to get a lot of high-resolution information about different cell populations. The analysis of these data can be time-consuming, but is aided by commercially available specialised software that requires the operator to gate on populations of interest and allow them to query not only percentages of cell types of interest, but the extent to which a population is positive for a specific marker. For example, one could determine the frequency of monocytes within a particular cell preparation and also assess the staining intensity of a particular marker on monocytes. A major area of current software development is automated flow cytometry analysis where gates will be determined by the software rather than the user, thereby streamlining analysis with less user input.

Example1. FLUOROCHROME PANELS

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مواضيع ذات صلة

?Why use a Radioisotope

Flow Cytometry: Applications

Fluorescence Labels

Fluorescence-Activated Cell Sorting (FACS)

Flow Cytometry – Instrumentation

Antibody Preparation

Harnessing the Immune System for Antibody Production

Electrophoretic Techniques : Support Media and Buffers

Principles of Liquid Chromatography

Preliminary Purification Steps

Cell Disruption and Production of Initial Crude Extract

Determination of Protein Concentrations