AFCG Lymphocyte Immunophenotyping Standard







Mr Greg Bryson, Dr Margaret Cooley, Ms Susan Francis, Mr Stephen Hunter

Mr Lyndsay Peters, Dr Henry Preston, Mr Joseph Webster


Ms Susan Francis, Mr Peter Hobson, Mr John Zaunders

Terms of Reference for the AFCG Lymphocyte Immunophenotyping Standards Committee

1. Committee composition: To be broadly based with a variety of experience

2. Quality control: To examine methods for inter- and intra­laboratory quality assurance for all areas of flow cytometry.

3. To report to the AFCG members on the recommendations for minimum standards for lymphocyte immunophenotyping.

4. To work towards establishing a joint AFCG /RCPA Quality Assurance program directed specifically at clinical flow cytometry.



Laboratory Safety

Specimen Collection

Specimen Transport

Specimen Integrity

Specimen Processing

Controls for Immunophenotyping

Flow Cytometer Quality Control

Sample Analysis

Data Storage

Data Reporting

Quality Assurance


Appendix 1: Determination of Reference Ranges

Appendix 2: Log Book Forms


These recommendations are presented with a view to being a minimum standard. These recommendations should not be seen to restrict the ability of any individual. This document will be reviewed regularly to ensure that these recommendations embrace current accepted laboratory practices.

The aim of lymphocyte immunophenotyping is to enumerate and identify specific sets or subsets of lymphocytes. This phenotypic analysis is usually performed on blood specimens; however, other body fluids may have to be examined.

These guidelines are written in broad terms as a gesture to indicate and not to dictate.


Each laboratory will adopt internal procedures and policies for the safe handling of biological specimens.

1. Use universal precautions1 with all specimens.

2. Develop appropriate internal procedures to cope with accidents such as spillages.

3. Handle and manipulate specimens in a safe biological confinement area wherever possible.

4. Fix cell samples with a 0.5% available formaldehyde-based solution for 15 mins before leaving a BSC.

5. Final cell suspension should be in a 0.5% ­ 1% available reactive formaldehyde-based solution.

6. Unfixed samples outside the safe handling area should be capped.

7. Appropriate safety devices such as gloves, gowns, goggles, centrifuge carriers, automatic pipetting are recommended whenever handling and processing specimens. Use disposable plastic equipment wherever possible.

8. Wash hands with medicated soap after working with specimens, removing gloves, or when leaving the laboratory, and as in accordance with usual local laboratory policy and universal precautions.

9. For decontamination of flow cytometers refer to the instrument manufacturer's recommended procedures.

10. Liquid waste should be treated with sodium hypochlorite. Solid waste should be handled carefully in appropriate robust containers.

11. Laser safety: Most benchtop flow cytometers use visible lasers which pose very little risk of injury to the operator. Operators should be aware of the potential dangers of lasers and the need for safety devices such as shields and goggles in given circumstances. The operator is referred to the manufacturer of the instrument and to AS 22111 with regard to safety of lasers.


1. Universal precautions1 should be strictly observed when collecting blood samples.

2. Each specimen should be labelled with the patient name or a unique patient identifier, and date and time of collection. If a pre-printed label is used, the signature or initials of the collector should appear on the label to verify that the information relates to the patient from whom the blood was collected.

3. Each specimen should be accompanied by a test requisition which should include the patient name or unique patient identifier, date and time of collection, age, sex, pertinent medication and presumptive diagnosis of the patient, name of requesting physician, and address for return of results.

4. The request form and specimen tube(s) should carry identical patient information. Both should be checked on receipt in the laboratory; in case of discrepancy or doubt, a clear, documented protocol approved by the Director/Scientist in Charge of the laboratory should be followed. Unlabelled samples and forms should be discarded.

5. A total white cell count and differential and/or stained blood film should be performed at the laboratory initiating the request within the time frame specified by the manufacturer of the haematology instrument used. For distant laboratories and dispatch centres, a total white cell count and unstained blood film should accompany each specimen.

6. EDTA anticoagulated specimens are suitable if the specimens are to be processed within a 12 hour period after collection.

7. Heparin or ACD anticoagulated specimens may be processed immediately or up to 48 hours after collection.

8. Any specimen over 48 hours old, unlabelled or incorrectly labelled, or of insufficient volume should be re-collected.

9. Bone marrow specimens should be received and processed within 12 hours of collection.


1. Packaging, labelling, and transport of specimens should comply with all current local, state, national, and international regulations for the regions through which the specimens will pass.

2. Specimens should be maintained at 18o ­ 22o C in a light-proof container.

3. Temperatures below 10oC should be avoided, and above 37oC must be avoided.


1. Visually inspect the specimen for clots, haemolysis, insufficient sample volume, or container defects. Re-collect the sample if the specimen shows any visible signs of deterioration.

2. Specimens which have been collected inappropriately may be processed by the laboratory according to a local approved, documented policy. The deficiencies in the sample should be noted and the report should reflect the effect that these deficiencies may have on the results.


1. The whole blood lysis method is generally recommended because it does not employ density gradients or lengthy centrifugation which may lead to differential losses of specific subpopulations. However, using this procedure assumes that all leucocyte subsets are equally tolerant to the lysis method used.

2. Several lysing techniques are available. These include water, tris-buffered ammonium chloride, and hypotonic buffer (4,5). Several proprietary lysing reagents are also available from instrument and monoclonal antibody manufacturers. For commercial reagents, the manufacturer's recommended protocol should always be followed unless data are available confirming that any modifications do not adversely affect results.

3. Where possible a full blood count and differential must be performed before processing, and the cell concentration adjusted accordingly. One should aim for a cell concentration of 1 x 106/test tube.

Specimens with pronounced leucopaenia may have insufficient cells for flow cytometric analysis, thus requiring a larger volume of sample or a buffy coat preparation. Conversely, normal concentrations of antibody reagents may be insufficient to saturate all binding sites in specimens with leucocytosis, leading to possible false negative results. Therefore, samples with leucocytosis may need to be diluted before testing. A balance between cell and antibody concentration should therefore be found.

4. Any panel of antibodies must include:

    4.1 Gating controls to allow for correction due to contaminating cells and/or particles (see Appendix 3 for lymphocyte gating).

    4.2 Isotype controls appropriate for the antibodies in the panel.

    4.3 A suitable panel of antibodies for investigating the presumptive diagnosis. The selection of antibodies used in the panel should be referenced.

5. Any deviation from the manufacturer's recommended protocol should be documented in a laboratory protocol book and only used when mean channel fluorescence or aberrant cell populations are not being studied. Such deviations should show that the results are comparable with those obtained using the recommended procedure.

NOTE: Excess reagent may cause increased nonspecific staining of negatives and may result in decrease of positive/negative resolution.


1. Isotype controls are recognised as an essential part of any monoclonal antibody panel for the purpose of establishing levels of non-specific binding and autofluorescence.

In many cases the isotype control may not be optimal for controlling non-specific fluorescence because of differences in fluorochrome / protein ratio and antibody concentration between the isotype control and the test reagents. This is particularly important in certain causes of leukaemia (in particular, myelomonocytic) where there is a high degree of species crossreactivity due to the presence of Fc receptors. At this time there is no solution to this problem.

2. A method control must be prepared and run on a daily basis in parallel with patient samples. At a minimum, a positive reagent control should be prepared and run whenever a new batch of any reagent used in cell preparation and staining is initiated.


These procedures should be carried out when the flow cytometer is first received, or when major maintenance or repair is performed.

1. Alignment of the optical components of the flow cytometer (laser, focusing lenses, collecting lenses, photodetectors, etc.) should be performed according to the manufacturer's recommended alignment procedures. These procedures should use the recommended alignment particles, which are typically uniform plastic beads incorporating a fluorescent dye. (Other materials may be recommended by the manufacturer). The laboratory must determine optimum settings for their own instrument / alignment particle combination and establish their own expected values. The expected range along with relevant instrument settings should be recorded in an instrument log book for subsequent use and daily monitoring. (See Appendix 2, Optical Alignment Log ).

Optical alignment can be verified by:

    1.1 Running alignment particles at instrument settings determined at time of initial instrument set-up.

    1.2 Daily recording of the mean channel fluorescence and CV for all parameters that will be analysed for test specimens in the daily log book and/or on Levy­Jennings plots (Optical Alignment Log).

    1.3 If particle values are not within acceptable range, alignment should be optimised before proceeding.

2. Regular verification of instrument sensitivity and spectral overlap compensation settings should be determined and recorded using cells or fluorescent microparticles.

    2.1 Instrument sensitivity is verified by:

      (1) Using freshly stained lymphocytes, establishing that positive­negative separations are acceptable, OR

      (2) If sensitivity particles (e.g. fluorochrome­labelled beads or nuclei) are used, run them at test­specific settings established at the time of initial set­up.

      (3) Record mean fluorescence channel and CV in the daily log book and/or on Levy­Jennings plots (Sensitivity Log).

    2.2 Spectral compensation is verified by:

      (1) Freshly stained cells using mutually exclusive antibodies, e.g. CD3-FITC and CD19-PE for lymphocytes.

      (2) If compensation particles (e.g. FITC- and PE­labelled beads) are used, run them at test­specific settings and compensation levels established at time of initial instrument set-up.

      (3) 3-colour or 4-colour compensation requires single-colour preparations for each additional fluorochrome.

      (4) Record mean channel fluorescence intensity for each population of interest (red only, green only, and negative for both) in the daily log book and/or on Levy­Jennings plots (Compensation Log).

    If particles values are not within acceptable range, compensation settings should be re­evaluated using antibody­stained leucocytes.

    Note: Overcompensation leads to fewer errors than undercompensation.

    (A) . (B)

    Figure 1

    Representation of application of correct compensation. Gated correlated display of anti CD3-FITC and anti CD19-PE. (A) Uncompensated. (B) Correctly compensated.

    3. Overall system performance can be verified by:

      (1) Running a "normal" specimen stained with an antibody reagent such as anti CD3-FITC and CD4-PE at test­specific instrument settings.

      (2) Verifying acceptable light scatter resolution of the leucocyte populations.

      (3) Verifying that the percentage of antibody­positive lymphocytes is acceptable by comparison with previous results, or with established laboratory ranges for the antigens selected (Appendix 3).

    If this positive control does not meet laboratory criteria, remedial action should be taken. Instrument performance and/or staining procedure should be checked to determine the source of the problem. Any problems identified using this sample must be rectified prior to analysis of test specimens.


    1. Sample order. Run and check all control specimens first, before running the patient samples according to laboratory priority.

    2. Test order within any panel. The first tube should be a gating control to maximise the cells of interest and minimise contamination. The appropriate isotype controls should be run next, followed by the subsequent test panel to investigate the provisional diagnosis.

    3. Assessment of specimen viability is desirable; however, because of biohazard concerns, it is recommended that all samples be appropriately fixed prior to analysis on the flow cytometer. It is not presently possible, on a routine large-scale basis, to distinguish those cells which were non-viable prior to fixation. However, this can be performed using ethidium monoazide (EMA) as described by K. Muirhead, 2nd AFCG Methods Course, 1989.

    Definition of a lymphocyte gate

    Figure 2

    Representation of common ways of displaying correlated low angle versus 90o angle light scatter seen from lysed whole blood preparations. (L = predominantly lymphocytes, M = predominantly monocytes, P = predominantly polymorphonuclear leucocytes).

    4. Set leucocyte gates as broadly as possible consistent with acceptable levels of contamination to minimise contaminating cells and maximise the inclusion of the cells of interest (see Appendix 3).

    5. Each laboratory should establish limits of contaminating cells and debris, based on documentation that their inclusion does not significantly affect the measurement of interest. If levels of contamination exceed established laboratory limits, the corrective actions taken are to adjust the light scatter gates and reanalyse the immunofluorescent correlated two-colour plot.

    Typical satisfactory values for lymphocytes are 95% (minimum 90%) of all lymphocytes and 90% (minimum 85%) purity in the gate as determined by CD45-FITC/CD14-PE gating control [Appendix 3].

    6. If levels of contamination by non-lymphocytes cannot be minimised to within acceptable limits, then test results may be suspect.

    If this contamination cannot be explained by reinterpretation of the data or by clinical diagnostic reasons, a second specimen should be requested.

    7. Count at least 2000 gated events in each sample. This number assures with 95% confidence that the result is within 2% of the "true" value (binomial sampling). NB: This sample mode assumes that the variability of determining replicates is < 2%.

    8. The counting of 2000 gated events to ensure reasonable statistical confidence may not be achievable in severely leucocytopaenic specimens.

    9. Most commercially available directly conjugated reagents give good resolution between low intensity negative and higher intensity positive cell populations. When simultaneous two-colour immunofluorescent correlated data is analysed, boundaries must be set to define four distinct regions: cells labelled with neither antibody, cells labelled with antibody #1 but not antibody #2, cells labelled with antibody #2 but not #1, and cells labelled with both antibodies.


    1. The possibility of patients' contesting the diagnostic implications derived in part from flow cytometric testing makes it incumbent upon the laboratory to be able to demonstrate and verify the process used in arriving at the reported test results.

    2. Where possible all listmode data on all samples analysed should be retained.

    At a minimum, retain correlated dual fluorescent data for each test and any interpretive comments on samples where a significant diagnosis is made.

    3. Retain all primary files, worksheets, and report forms.

    4. Minimum duration of data storage depends on state and federal regulations. These regulations may vary and each laboratory will need to remain informed of the current requirements.


    1. Analysis should include internal reliability checks of results, including:

      a) Optimally, the sum of CD3+% plus CD19+% plus CD3-CD16+ and/or CD56+ (the "lymphosum") should equal the purity of lymphocytes in the gate ± 5%, with a maximum variability of £ 10%. If the data are corrected for lymphocyte purity, then the lymphosum should be between 95 and 105% (minimally 90-110%).

      b) Optimally, the sum of the CD3+CD4+% plus CD3+CD8+% should be no more than 5% more than the CD3+%, and no more than 10-15% less than the CD3+%, depending on the number of g/d-TCR+CD3+ cells present.

      c) Replicate results within a panel (e.g. CD3+%) for the same sample should be within 5% of each other for FS v SS gating or within 3% for CD45 v SS gating.

      d) Light scatter patterns should be examined for each tube within the panel for variation from tube to tube. Similarly, the number of gated events and/or time to collect data should not vary greatly from tube to tube.

    Potential sources of error which are not necessarily covered by the above reliability checks may include inappropriate gating leading to exclusion of relevant cells, tubes in a panel run in the wrong order, inappropriate cut-offs between negative and positive cells, and calculation or transcription errors. Individual laboratories may require procedures to cover such possibilities.

    2. Each laboratory should determine the level of test variability by preparing and analysing at least six replicates. This will provide a basis when changes to methodology are introduced.

    Example 1: A sample control measure is the lymphosum2, which is the sum of T cell %, B cell %, and NK cell % ; ideally this should equal 100% for assays corrected for gate purity. Typically, lymphosum values of 95%­105% are acceptable.

    Example 2: Tube-to-tube variation can be monitored by the inclusion of the same antibody in separate tubes within the one patient test series.

    3. Regulatory bodies currently require that a laboratory keep all equipment maintenance and calibration records, staff training records, up-to-date method protocols, daily operator/reagent records, verification of transcription of results from machine printouts, procedures for amendment of results, and checks by supervisors/pathologists.

    4. Where possible, the laboratory should belong to and participate in a recognised external Quality Assurance program with regular review of the results.


    1. Report all unique patient identifiers including name/code, medical record number, laboratory ID/accession number, and collection date/time as well as print date/time.

    2. Report all data in terms of cluster of differentiation (CD) with a short description of the main antigen recognition characteristics.

    3. For unclustered antibodies, report the clone name with a short description of the main antigen binding characteristics.

    4. For blood specimens, report all data as a percentage and absolute number of the population of interest within the gate as determined by the gating control.

    5. Report data from all relevant antibody phenotyping combinations with corresponding reference limits of expected normal values, e.g.: CD3+8+ suppressor/cytotoxic T Cells ± % and/or ± absolute values.

    Reference limits for immunophenotyping test results must be determined for each laboratory.

    6. Each laboratory should establish reference limits for the antigens being tested (see Appendix 1).


    1. Universal precautions: There appears to be no single document that addresses the specific needs of flow cytometry.

    It is recommended that readers refer to the following documents:

      (i) Australian Standard AS 2211-1991, Laser Safety.

      (ii) Australian Standard AS 2243.3 ­1995, Safety in laboratories, Part 3: Microbiology.

      (iii) NCCLS M29­T, Protection of laboratory workers from infectious disease transmitted by blood, body fluids and tissue.

      (iv) MMWR 1988;37(24):377­82, 387­8. CDC Update: Universal precautions for the prevention of transmission of human immunodeficiency virus, hepatitis B virus, and other bloodborne pathogens in health care settings.

    2. NCCLS. Vol 12 No 6.

    3. MMWR MARCH 4, 1994 / Vol. 43 / No RR­3. 1994 Revised Guidelines for the Performance of CD4+ T­Cell Determinations in Persons with Human Immunodeficiency Virus (HIV) Infection.

    4. Muirhead, K.A., Wallace, P.K., Schmitt, T.C., Rescatore, R.L., Ranco, J.A., Horan, P.K. Methodological considerations for implementation of lymphocyte subset analysis in a clinical reference laboratory. In Clinical Cytometry. M. Andreeff, ed. Ann. N.Y. Acad. Sci. Vol. 468, pp 113­127, The New York Academy of Sciences, New York, N.Y., 1986.

    5. Loken, M. R., Meiners, H., Terstappen, LWM. Comparison of sample preparation techniques for flow cytometric analysis of immunofluorescence. Cytometry Supplement 2:53, 1988.

    6. Schlossman, SF, et al. (eds). Leucocyte Typing V. White Cell Differentiation Antigens, Oxford University Press, 1995.

    7. Nicholson, JKA, Hubbard, M and Jones, BM. Use of CD45 fluorescence and side-scatter characteristics for gating lymphocytes when using the whole blood lysis procedure and flow cytometry. Cytometry 26: 16-21, 1996.


    1.0 Definitions

    Reference values: Set of values for any measured quantity.

    Reference interval: Classically, the range of values found in 95% of a reference population of healthy individuals without overt clinical disease.

    NOTE: Age, sex, and race are factors known to influence reference intervals.

    2.0 Procedure for Determining Reference Ranges

    Statistical methods, both parametric and nonparametric, and graphical methods are discussed in detail in references 1­3. Only a brief summary of the steps involved is presented here.

      2.1 Parametric methods

        (1) Collect data on randomly chosen set of representative individuals (e.g. 50 healthy individuals).

        (2) Inspect frequency distribution of values obtained.

        (3) If frequency distribution is Gaussian, use appropriate statistical techniques to estimate 95% confidence interval and use endpoints of interval as the reference range.

        (4) If frequency distribution is non­gaussain, back transform endpoints of 95% confidence interval to obtain reference range, (e.g. log X, of (X + C), square root X, arcsin X), and proceed as in step 3.

        (5) If no satisfactory transformation can be identified, use nonparametric methods which do not depend on the exact distribution of the data.

      2.2 Nonparametric methods

        (1) Collect data on randomly chosen set of representative individuals.

        (2) Arrange data in ascending or descending order.

        (3) Use appropriate nonparametric techniques to identify desired limiting percentiles (e.g.  2.5 and 97.5) to desired confidence level.

      Nonparametric methods are most appropriate when data do not show a Gaussian distribution and cannot be so transformed. However, they are very sensitive to outliers, and final ranges chosen may be highly dependent on methods used for removing outliers (1­3).

      3.0 Pitfalls in Determining Flow Cytometric Reference Ranges

      Each laboratory should determine its own reference range using its particular preparation method and instrumentation, because significant laboratory­to­laboratory differences related to these variables have been reported.

      However, quite large data sets are technically required to carry the above described methods for reference range determination, typically >300 for parametric methods and >120 for establishing nonparametric interval with 90% confidence. Until more standardised methodology allows pooling of data among laboratories (hence this document), this is clearly an unrealistic expectation.

      Other confounding variables besides sample size have been described (4­5).

      One practical solution to the dilemma is to accumulate and analyse reference data in smaller sets (e.g. 10­20 individuals), which can then also be pooled and analysed. If the last two sets of pooled data are found to give the same reference range within experimental error, this gives increased confidence that the reference range selected is not unduly affected by the small sample size.


      1. Winkel, P., Statlan, B.E. Reference values. In Clinical Diagnosis and Management by Laboratory Methods (ed J.B. Henry), Philadelphia, W.B. Saunders Co., 1979, pp. 29­52.

      2. Martin, H.F. Gudzinowicz, B.J. Fanger, H. Normal Values in Clinical Chemistry, New York, Marcel Dekker, 1975, pp. 102­236.

      3. Henry, R.J., Cannon, D.C., Winkelman, J.W. Clinical Chemistry. Principles and Technics, New York, Harper and Row, 1974, pp. 343­371.

      4. Edward, B.S., Altobelli, K.K., Nolla, H.A., et al. A comprehensive quality assessment approach for flow cytometric immunophenotyping of human lymphocytes. Cytometry 10:443­441, 1989.

      5. McCarthy, R.C., Fetterhoff, T.J. Issues in Quality Assurance in Clinical Flow cytometry. Arch. Pathol. Lab. Med.113: 658­666, 1989 (in press).

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