FAQ

International Clinical Cytometry Society Guidelines (ICCS) and multiple other experts suggest that the clinical presentations in several conditions increase the likelihood of paroxysmal nocturnal haemoglobinuria (PNH) as follows: bone marrow failure syndrome, aplastic anaemia, myelodysplastic syndrome (MDS), haemolytic anaemia, haemoglobinuria, unexplained cytopenias and unexplained thrombosis.^1-24 Positive test results for any of these conditions may increase the likelihood of PNH and lead to reflex confirmatory testing for PNH clones by flow cytometry.

The correct flow cytometer set-up is crucial for producing accurate and reliable results. For PNH analysis, we recommend carrying out a daily performance check using standard beads (Flow Check beads, calibration beads, Cytometer Setup and Tracking beads) to establish appropriate application settings (photomultiplier tube setting, time delay and colour compensation).

Ideally, application setting should be performed with non-labelled target cells (red blood cells [RBCs], neutrophils, monocytes) and cells labelled with all conjugates used in the assay. Alternatively, commercial compensation beads (calibration beads, CYTO-COMP beads) or cells (lymphocytes, CYTO-COMP cells) can be used. Spectral overlap compensation can be performed manually; however, we advise using a digital compensation set-up.

Three-colour combinations are no longer recommended for PNH testing, as these combinations can only include either CD45 or a lineage-specific gating antibody, but cannot include both. For high-quality PNH testing, consensus guidelines recommend using 4–7-colour combinations.²⁴

EDTA is the preferred anticoagulant, although heparin or acid citrate dextrose can be used.²⁴ For RBC analysis, samples should be kept at 4–8°C. For neutrophil analysis, samples should be kept and transferred at room temperature.

Analysis should be performed within 24–48 hours, providing the sample has been stored at 4°C.²⁵ Older samples may be used for baseline samples, although testing results should be treated with caution as neutrophils may be significantly decreased such that there are insufficient numbers for analysis.

Samples should be packaged with the utmost care. Ideally, specialised sample transfer containers with temperature recorders should be used. The sample tube should be placed in a covered tube or cup and should not have direct contact with the ice pack.

Peripheral blood is the recommended specimen for PNH testing and diagnosis.²⁴ Bone marrow is not a suitable specimen, as immature myeloid populations may express lower levels of glycosylphosphatidylinositol (GPI)-anchored proteins, including CD14, making interpretation of results difficult.^1,24

Guidelines recommend testing both RBCs and WBCs taken from peripheral blood when trying to detect a PNH clone.¹ PNH testing of RBCs does not provide adequate information to fully evaluate patients with PNH routinely, as the effects of haemolysis and blood transfusions mean that tests on RBCs can underestimate clone size.¹ In addition, testing of RBCs alone does not exclude the possibility that the symptoms are the result of an inherited, isolated deficiency in one GPI-anchored protein.

Testing of WBCs and RBCs is widely recognised as the best approach to assessing the true size of a PNH clone. Ideally, both neutrophils and monocytes should be assessed to gain an accurate idea of true PNH clone size.²² Lymphocytes are not suitable for PNH testing due to their long lifespan and variable GPI-linked proteins, but they can be used as an internal control.^1,26

The neutrophil clone size is typically used to assess the WBC clone size; however, the monocyte clone size often differs and sometimes shows a higher PNH clone than is found in the neutrophils. The reason for this is not well understood at present; therefore, both neutrophils and monocytes should be analysed to get a more thorough assessment of the PNH clone size in the WBCs and thereby not underestimate the true PNH clone size.²²

Lymphocytes are not a suitable target for PNH testing because of their long lifespan.¹ However, lymphocyte analysis is suitable for discrimination of B and T lymphocytes, natural killer cells and basophils. Moreover, lymphocytes are useful as internal controls for proper voltage/compensation settings and assessment of the performance of the antibodies and conjugates used in the assay.^24,26

The number of cells to be analysed depends on the desired sensitivity. For most assays, the acquisition of 50,000 neutrophils is sufficient to achieve a 0.1% sensitivity (50 PNH events). In some instances, more cells may need to be acquired to detect smaller PNH populations (0.01–0.1%). If the result is reaching your laboratory’s sensitivity level (eg 0.01%), you should set up the test with more sample volume to acquire more PNH cells and see if this finding is reproducible. We recommend acquiring 20–100 definitive GPI-deficient events to be able to report a PNH population,²² but in some instances you may report on fewer as long as the cells are clustered; backgating confirms that these cells are in the ‘right location’ for PNH cells and that they are also seen in other lineages.

Yes, washing RBCs and WBCs is recommended to remove any unbound antibody that could lead to the production of false-negative results. RBCs should be washed twice with phosphate-buffered saline (PBS) following incubation with monoclonal antibodies (and following RBC lysis in the case of WBCs).^1,24,26

The recommended monoclonal antibody for PNH RBC detection is CD59. Both MEM43 and OV9A2 clones conjugated to phycoerythrin demonstrate good signal-to-noise ratio and are suitable for PNH testing. CD55 monoclonal antibody is too dim, even conjugated to phycoerythrin, and is probably not necessary for PNH testing.^1,24

The most frequently used antibody/reagent combinations for PNH neutrophil and monocyte testing are FLAER/CD24 and FLAER/CD14 or, for single-tube 5-colour assays, FLAER/CD157 for both neutrophils and monocytes.²⁴

Lineage-specific gating is crucial and the choice of gating antibody determines the level of sensitivity. CD15 works well to gate on mature CD15++ neutrophils, while CD64 works well for gating on CD64++ monocytes. CD33 has been historically used for monocytes, but is not recommended as it is not effective at delineating monocytes from basophils; some basophils show intermediate CD33 expression, as well as dim FLAER expression, and may be mistaken for GPI-deficient cells.²⁴

When testing for PNH, sample preparation is often the key stage in ensuring accurate results. A common cause of false negatives is not washing the RBCs following antibody incubation.²⁴ Washing RBCs twice reduces background noise and helps ensure that any PNH population is detected.

Another common problem encountered in sample preparation is aggregation of RBCs, which can make interpretation of results difficult. Vigorous vortexing or ‘racking’ (dragging the sample tube across a tube rack) helps prevent RBC clumps from forming and thereby minimises the presence of aggregates in the sample.²⁴

The cause could be poorly stained cells, perhaps due to the dropping of cells on the flow cytometry measurement tube during the staining procedure. Take care not to leave any blood on the edges of the tube. Other explanations include poor instrument set-up, poor antibody clones or conjugates that give a poor signal-to-noise ratio.²⁴

Another important issue in rare-event analysis is the presence of background noise (non-specific events). The reason for seeing cells in the area of PNH cells could be one of the following:

1. Contamination (bacteria, fungi) in buffers (PBS). It is very important to filter reagents to avoid artefacts during PNH testing. An empty tube with PBS, water and sheath fluid can quickly reveal the culprit.
2. Contamination in sheath fluid (crystal formation). Run a bleach solution and cleanse the instrument thoroughly. Always wash the instrument thoroughly before analysing PNH samples.

It is also important to use proper lineage-specific gating²⁴ and look at dot plots instead of single-colour histograms. For example, in the case of RBC analysis, addition of CD235a (RBC specific) would identify the presence of non-RBCs. If CD235a were not added, the single-colour histogram would show a population of non-specific events as GPI-negative events in the Type III region, which could lead to a false interpretation of ‘PNH cells’.²⁶

Improper validation of flow cytometry assays is one factor responsible for reported interlaboratory variability in PNH testing.²⁵ To validate your PNH assay, we recommend taking the following steps:

1. Include appropriate positive and negative control samples (eg normal samples from healthy donors and pathological samples from PNH patients).
2. Analyse the performance characteristics of your assay (accuracy and reproducibility).
3. Participate in an external quality-assessment programme.^25,27

PNH clone size can increase over a relatively short period in some patients.²⁸ Regular monitoring is therefore important, especially in patients with a small clone size. For patients with a clone size of <0.01%, annual monitoring should be considered, and for patients with a clone size of >0.1%, a minimum of twice-yearly monitoring is recommended.²⁸

A test can be referred to as ‘high-sensitivity analysis’ when the sensitivity level is <0.01%, whereas the standard test has a sensitivity of <0.1%. To reach the high-sensitivity threshold, 50,000–100,000 cells of a given type must be acquired.

This is easily achievable for RBCs but requires a rather large acquisition for neutrophils. It is difficult to reach high sensitivity for monocytes due to their low counts in peripheral blood; however, double or triple amounts of sample can be prepared when necessary.

Sensitivity refers to the smallest number of cells that you can specifically detect with your assay. You can determine it by means of a spiking experiment: serial dilution of a PNH clone in normal blood. Using well-calibrated protocols, it is possible to detect 20 PNH Type III RBCs in 1 million events with the 4-colour assay. The background for normal individuals and blood donors is 0–6 cells/million.²⁵

Depending on the size of the GPI-deficient population/PNH clone relative to the whole population, the following terminology should be used for reporting:

• PNH population >1% = ‘PNH clone’.
• PNH population 0.1–1% = ‘minor population of PNH cells’ or ‘minor PNH clone’.
• PNH population <0.1% = ‘rare cells with GPI deficiency’ or ‘rare cells with PNH phenotype’.

No detectable level of GPI-deficient cells should also be reported. Further testing may be indicated, in the case of a PNH clone, to inform a clinical decision and determine the next steps in patient management.²²

No, the detection of PNH Type II RBCs is an artefact. It is not possible to identify an RBC PNH clone without the simultaneous presence of PNH neutrophils and monocytes.²²

In samples with dysplasia, neutrophil and monocyte populations can be difficult to distinguish; hence, careful gating on fully matured neutrophils and monocytes is essential.

In samples with aplastic anaemia, neutrophil and monocyte counts may be very low; hence, bulk lysis of several mLs of blood may be necessary to achieve sufficient WBC numbers and statistically meaningful results.