Immunophenotyping by Flow Cytometry

Principles of flow cytometric immunophenotyping

The primary methods for immunophenotyping in the haematological setting are immunocytochemistry (described in previous editions of this book) and flow cytometry.

In this chapter we shall focus on flow cytometry and its application to the diagnosis and treatment response of haematological neoplasms.

Flow cytometry is a laser-based technology, which is able to identify and quantify cell populations. Cells in suspension are manipulated into a stream of fluid where single cells are interrogated by an electronic detection system. Flow cytometry has many applications and is now routinely used in the diagnosis of haematological malignancies. It can provide quantitative data on a number of cell parameters for a large number of cells, in the region of thousands of cells per second, making it a powerful diagnostic tool.

A flow cytometer has five main components:

• Lasers providing a monochromatic light

• A flow cell with a liquid stream (sheath fluid), which carries and aligns the cells so that they pass in single file through the laser beam

• Optical systems and filters regulating the light signals

• Photomultiplier detectors (PMTs) that generate data on forward light scatter (FSC) (which provides an approximation of cell size) and side light scatter (SSC) (which relates to cell complexity), as well as converting fluorescence signals from light into electrical signals that are processed by a computer

• A computer for analysis of the signals.

Flow cytometry relies on the application of fluorescent conjugates or fluorochromes attached to a monoclonal antibody (McAb) that has specific affinity for an antigen on the cell surface, in the cytoplasm or in the nucleus. Each fluorochrome has a characteristic peak excitation and emission wavelength, and the emission spectra often overlap, requiring compensation. Consequently, the combination of fluorescent labels that can be used depends on the wavelength of the laser used to excite the fluorochromes and on the detectors available.

Immunophenotyping of haematological neoplasms by flow cytometry involves the labelling of white blood cells or their precursors with fluorescent-labelled monoclonal antibodies directed against target cellular proteins or antigens. When appropriate antibodies – usually combined as ‘panels’ – are chosen, the cell lineage and stage of differentiation of leukaemic or lymphoma cells can be determined, allowing the leukaemia/lymphoma to be classified. The accurate classification of haematological neoplasms is critical for the clinician’s choice of treatment and for overall prognosis.

In summary, immunophenotyping by flow cytometry facilitates:

• The identification and quantification of cell populations within a sample

• The differentiation of normal from abnormal cells

• The differentiation of reactive from neoplastic cells

• The identification of the differentiation or maturation stage of a cell population

• The quantification of tumour infiltration.

However it is the interpretation of the data provided by the described techniques which poses the greatest challenge and the reliable diagnosis of leukaemia relies on the following:

• Knowledge of physical characteristics/antigen expression on normal cells

• The ability to distinguish between different patterns of expression of antigens

• The ability to identify aberrant antigen expression

• The identification of a robust leukaemia-associated immunophenotype (LAIP).

These will be discussed in more detail in the sections below.

Multicolour flow cytometric immunophenotyping

Flow cytometry is now a well-established technique for identifying immunophenotypic profiles in haematological neoplasms. Historically three- or four-colour antibody flow cytometry panels were used, and these panels remain of great utility. However, developments in instrumentation, namely an increase in both the number of lasers and in the availability of fluorochrome conjugates, have facilitated the design of eight- to ten-colour antibody panels. Such panels can provide data on up to 12 to 14 cellular parameters simultaneously, allowing the accurate identification and quantification of normal and abnormal cell populations in bone marrow and peripheral blood. The application of such extensive panels has provided a greater understanding of normal maturation stages in haemopoiesis and specifically the antigens expressed or down regulated at the different stages of this process. This provides a template to which abnormal cell populations can be compared. This new knowledge has increased the accuracy of diagnosis and allowed the identification of very small cell populations and subpopulations.

The advances in flow cytometry instrumentation and reagents mentioned above have facilitated the routine use of eight- to ten-colour antibody panels in the clinical laboratory setting. The use of such multicolour panels is generally what is referred to as multicolour or multiparameter flow cytometry (MFC). There are many advantages associated with the use of MFC but the corresponding pitfalls must also be considered.

Advantages of MFC:

• Increased accuracy. Using large numbers of fluorochromes is associated with an exponential increase in the information obtained from a single combination of antibodies in the same tube, permitting a more reliable identification.

• Smaller sample size. Increased number of antibodies per tube means fewer tubes and less sample needed but allows acquisition of more cellular events resulting in smaller coefficients of variation and increased data precision. This is of particular relevance to paucicellular samples such as cerebrospinal fluid (CSF) and fine needle aspirates (FNA) and also paediatric samples.

• Cost effectiveness. Less usage of repeating backbone or gating antibodies.

• Increased efficiency. Less time is required for sample processing and acquisition.

• Increased sensitivity for minimal residual disease monitoring.

Disadvantages:

• Increased complexity of compensation. Inaccurate compensation is probably the main source of erroneous data in MFC. This can be solved by applying compensation matrices but this requires expertise.

• Challenges of antibody panel validation. It is crucial to run fluorescence minus one controls for all new antibody combinations and to check for stearic hindrance between antibodies used to label antigens that are in close proximity on the cell.

• Tandem dye conjugate issues. Tandem dyes are conjugates of two fluorochromes, but this can lead to problems in resonance excitation transfer if exposed to light. Ideally a compensation matrix should be performed for each new tandem dye conjugate lot.

• Increased need for expertise in data analysis and interpretation.

• Human error associated with pipetting a high number of antibodies into a single tube. This can be overcome by preparing in-house McAb cocktails, which have been shown be stable for up to 4 weeks.

Many advances have been made in addressing the issues outlined above and groups such as the EuroFlow Consortium, the Multicolour Immunophenotyping Group UK (MIG UK), European Group for the Immunological Characterization of Leukemias (EGIL), Harmonemia and others have made great advances in standardising MFC protocols. Software tools such as Kaluza ( www.beckmancoulter.com ) and Infinicyt ( www.infinicyt.com ) have been developed to aid in data analysis and interpretation ( Fig. 16-1 ). Currently there is a move towards commercially available kits that include lyophilised or freeze-dried antibody ‘cocktails’ to overcome the issues associated with tandem conjugates. Some kits also include the use of standardised instrument set up facilitating the use of software analysis using libraries to compare normal with neoplastic cases .

Detection of surface immunoglobulin

Lymphoproliferative disorders (LPD) of mature B cells are distinguished from their normal counterparts by identifying two main types of phenotypic abnormalities: surface immunoglobulin light chain restriction and aberrant B-cell antigen expression.

Staining for surface immunoglobulins requires some extra steps in the sample preparation. This is to avoid any nonspecific binding due either to cytophilic antibodies binding to Fc receptors (monocytes and some lymphocytes) or to the binding of antibodies to cell membranes of damaged or dying cells.

This nonspecific staining can be avoided by washing the sample with an isotonic solution prior to staining for surface immunoglobulins. Nonspecific staining can also be minimised by incubating the cells with serum prior to staining.

Finally, some B-cell LPDs such as chronic lymphocytic leukaemia (CLL) may express surface immunoglobulins very weakly and it is preferable to use polyclonal antibodies (PcAb) to detect light chain restriction in these cases ( Fig. 16-3 ).

Two methods are suitable for detecting surface membrane immunoglobulin (SmIg) of blood and bone marrow cells, according to whether a PBS wash or a lysing procedure is used as the first step.

Method 1: wash–stain–lyse–wash

• Pipette 100 μl of the specimen into a round-bottom tube.

• Add 2 ml of PBS–Azide–BSA kept at 37 °C and centrifuge for 5 min at 300 g. Using a Pasteur pipette, carefully discard the supernatant.

• Repeat the procedure and resuspend the specimen in 50 μl of PBS–Azide–BSA.

• Add the appropriate McAb/PcAb combination, (e.g. anti-kappa and anti-lambda, CD19, CD45 or any multicolour cocktail).

• Incubate at room temperature in the dark for 15 min.

• Add 1 ml of ammonium chloride-based lysing solution and incubate for 10 min at room temperature.

• Add 1 ml of PBS–Azide–BSA, centrifuge for 5 min at 2000 revolutions per minute (rpm), and discard the supernatant. Repeat this step.

• Resuspend cells in 0.2–0.5 ml of sheath fluid solution (e.g. Isoton).

• Acquire data on a flow cytometer without delay.

Method 2: lyse–stain–wash

• Pipette 100 μl of the specimen into a round bottomed tube.

• Add 2 ml of ammonium chloride lysing solution, incubate for 10 min at room temperature, and wash twice in PBS–Azide–BSA as above.

• Add the appropriate volume of McAb/PcAb combination, according to the manufacturer’s instructions.

• Incubate in the dark for 15 min at room temperature. Add 2 ml of PBS–Azide–BSA, centrifuge for 5 min at 300 g and discard the supernatant. Repeat this step.

• Resuspend cells in 0.2–0.5 ml of sheath fluid (e.g. Isoton) and acquire data on a flow cytometer without delay.