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• W1982327519 abstract "Since the publication of the “Three-color supplement to the NIAID/DAIDS Guideline for Flow Cytometric Immunophenotyping” in 1996 (1), significant scientific and technological advances in the development and production of reagents, instrumentation, and software have increased the use of multicolor flow cytometry in both research and clinical laboratories. With the increased adoption of three and four-color flow cytometry as the preferred methodology in determining patients' CD4 and CD8 T-cell counts, it has become apparent that a gating strategy that integrates the bright CD45 cells reduces interlaboratory and intralaboratory variability. Traditionally, a lymphocyte gate for immunophenotyping is derived from a bivariate frequency distribution histogram that includes 90° side scatter (SSC) and forward light scatter (FSC) frequency patterns. This type of histogram configuration is called a homogenous gating protocol (2). The advantage of the combination of bright CD45 fluorescence and light scatter, a heterogeneous gating protocol, was first reported in 1993 (3). Over the past few years, it has been determined that this alternative approach provides a more reproducible and accurate lymphocyte gate (4-6). This heterogeneous method will be referred to as the CD45 gating method. The purpose of this article is to update the 1996 NIAID/DAIDS Guideline and include the use of the CD45 gating method to minimize measurement variability with multicolor flow cytometry for the enumeration of T-cell subsets. Some information is provided about the advantages of four-color flow cytometry and the use of single-platform bead-based technology for determining absolute and percentage of lymphocytes. The addition of integrated fluorosphere counting provides a single-platform protocol that facilitates the simultaneous determination of both absolute and percentage of lymphocyte subsets. The specifications and recommendations were developed for use in laboratories supporting clinical trials and epidemiological studies carried out under the auspices of the National Institute of Allergy and Infectious Diseases, Division of AIDS (NIAID/DAIDS). Efforts currently underway by the Centers for Disease Control and Prevention, as well as the National Committee for Clinical Laboratory Standards (NCCLS) and Health Canada's National Laboratory for HIV Immunology, will likely provide additional guidance in this rapidly changing arena. The antibody selection includes several commercially available premixed and numerous “theoretically applicable” combinations. A three-color panel for the routine measurement of lymphocyte subsets includes CD45/CD3/CD4 and CD45/CD3/CD8 combinations so that dual positive CD3+CD4+ and CD3+CD8+ cell percentages can be reported. A minimum four-color panel consists of a single tube containing CD45/CD3/CD4/CD8. If not using commercially premixed or Food and Drug Administration (FDA)-cleared combinations of antibodies, the laboratory must follow Subpart K of the Clinical Laboratory Improvement Amendments of 1988 (CLIA) to validate their reagents (7). The NIAID/DAIDS Flow Cytometry Advisory Committee recommends the multicolor panels using a fluorescein isothiocyanate (FITC)/phycoerythrin (PE)/third color/fourth color fluorochrome assignment order (Tables 1, 2). For all NIAID/DAIDS adult studies, it is mandatory to report both percent and absolute CD4+ and CD8+ T-cell counts and these subsets should be assayed as CD3+CD4+ cells and CD3+CD8+ cells, respectively. For all NIAID/DAIDS pediatric studies, in addition to reporting percent and absolute CD4+ and CD8+ T-cell counts as above, it is also mandatory to report total B cells, measured as CD19+ cells. NIAID/DAIDS adult studies do not require inclusion of markers for B and natural killer (NK) cells. However, these markers can be useful in accounting for additional cell populations for quality control purposes. For three and four-color NK cell analysis, it is recommended that laboratories stain with CD56 or CD16 antibodies conjugated to the same fluorochrome. The CD56 and CD16 antibodies are used in combination with CD3 and CD45 antibodies to identify the majority of NK cells (8, 9). Lymphocyte gates are set using linear 90° SSC and log CD45 fluorescence. Lymphocytes are defined as CD45bright with low SSC (Fig. 1) (3). CD45 gating is highly recommended by NIAID DAIDS Flow Cytometry Advisory Committee for all CD3+CD4+ and CD3+CD8+ determinations for NIAID-sponsored trials. When using a single-platform method for absolute subset counts, CD45 gating is required. In this case, the setting of the signal discriminator or the signal threshold is critical because the beads used by the two manufacturers are of different size and fluorescence intensity. Therefore, BD Bioscience (San Jose, CA) uses a fluorescence signal discriminator whereas Beckman Coulter (Hialeah, FL) uses a forward angle light scatter signal threshold for generating CD45 versus SSC histograms. a,b: Dual-parameter histograms of SSC and CD45 fluorescence. Displays show the gated lymphocyte population among granulocytes, monocytes, basophils, and debris. Recent studies suggest that CD45 gating provides more consistent and precise analyses. However, there are times when it is not easy to resolve monocytes from lymphocytes as is the case in b. One should run a dual-color dot plot with CD3 and CD4 from the gated area A, inspect the CD4+CD3− quadrant, and make adjustments to gate A until the CD4+CD3− events become insignificant. Laboratories that desire to switch from a two-color immunophenotyping method to a three or four-color method will be required to incorporate the CD45 gating method. (Consult the AACTG web site at http://aactg.s-3.com/iqa.htm for additional information.) Laboratories that switched to three or four-color immunophenotyping protocol prior to the establishment of this guideline may continue to perform their current methodology only if they maintain a satisfactory performance rating as determined by the DAIDS Immunology Quality Assessment (IQA) Program. If a laboratory that is currently performing three or four-color immunophenotyping without CD45 gating begins to perform unsatisfactorily, the laboratory will be required to switch to CD45 gating. Recent studies reported that non-CD45 gating laboratories were significantly more likely to have an unacceptable interlaboratory result than CD45 gating laboratories (4, 10). As shown in Figure 2, the deterioration of sample integrity that is usually observed with traditional dual light scatter gating is minimal for up to 4 days when CD45 gating is used (11). Until there is substantial published evidence from multisite evaluations that supports the robustness of CD45 gating for up to 4 days, efforts should be made to process as soon as possible samples that are more than 24 h old. Comparing sample integrity at Day 0 and Day 4 using heterogeneous and homogeneous gating strategies. Note the debris in the lower left-hand corner of the Day 4 homogeneous gate dot plot. Heterogeneous gating options, such as T-cell gating (CD3) and primary CD4 T-cell gating, are not recommended for NIAID/DAIDS-sponsored trials because T-cell percentages cannot be obtained from the flow cytometer using these gating strategies. Use of a single CD3/CD4/CD8 tube with CD3 T-cell gating may affect both the accuracy and the reproducibility of T-cell subsets when reporting results as lymphocyte percentages (4). In addition, this panel cannot be used with the single-platform methods for T-cell subsets because these methods use a lyse/no-wash technique that makes lymphocyte identification impossible without the use of CD45. Other heterogeneous gating strategies incorporating CD45, which aim at reducing costs and allow for both absolute counts and subset percentage calculations, may be practical and are currently being validated. The advantages of CD45 gating are that 1) lymphocytes are distinguished easily based on CD45 fluorescence and 90° (side) scatter even in the presence of a large amount of debris, which allows the use of the lyse/no-wash methodology; 2) an external isotype control is not required; 3) gating on “bright” CD45 for each tube helps ensure inclusion of only lymphocytes, eliminating the need to correct for lymphocyte gate purity; and 4) many nonlymphocyte contaminants in a traditional homogeneous FSC/SSC lymphocyte gate (e.g., unlysed red blood cells) can be excluded easily with the heterogeneous CD45 gate. The disadvantages are that 1) the distinction among NK cells, monocytes, B cells, and debris may not always be clear-cut when drawing the heterogeneous gate; and 2) the traditional lymphocyte recovery value cannot be determined if antibody panels only include CD3, CD4, and CD8. Antibody preparations and software are commercially available for use with whole blood staining that do not require a wash and centrifugation step after the lyse/fixative reagent is added. This development shortens preparation time and eliminates the need for a centrifuge. This is mandatory for single-platform methodologies, which require that lymphocyte subsets be maintained in the original blood volume for an absolute cell count determination. The presence of nonlymphocytes within the boundaries of the lymphocyte gate is assumed to be negligible. By using a heterogeneous histogram (low SSC and CD45bright fluorescence) for identification of lymphocytes, an assumption is made that the only cells meeting these criteria are lymphocytes, and therefore, no correction is needed for the lymphocyte subset percentages (2). The proportion of all lymphocytes (T, B, and NK cells) present in the specimen and contained within the boundaries of the homogeneous FSC/SSC lymphocyte gate or the heterogeneous CD45 gate cannot be determined using mAb panels that include only CD3, CD4, and CD8. To determine lymphocyte recovery (T, B, and NK cells), the antibody panel must include tubes for B and NK cell determinations (12). Fluorescence data should be displayed as dual-parameter histograms of CD3 log fluorescence versus log fluorescence of CD4, CD8, or CD19 as outlined in the 1993 Guideline, section 3.09a (13). An isotype control is not required when using the CD45 gating strategy for CD3+CD4+ and CD3+CD8+ subset determinations because the unlabeled (negative) lymphocyte populations in each tube serve as an internal immunological control for nonspecific binding. Cursor settings are determined by the fluorescence patterns from the negative and positive populations for CD3, CD4, and CD8. Because the CD3, CD4, and CD8 antibodies label lymphocytes brightly, the cutoff between the negative and positive populations is determined easily (Fig. 3). Dual-parameter histograms generated from a CD45 fluorescence and SSC gate, showing the fluorescence distributions of specimens stained with CD3 and CD4 (a,b) and with CD3 and CD8 (c,d). Cursors were positioned based on the unlabeled cells in the histograms. A standard subtraction/compensation protocol to correct the spectral overlap of one fluorochrome into the fluorescence spectrum of another is necessary for multicolor analysis (14, 15). Commercially available fluorospheres and automated software are available for this protocol. For more information on the procedure and theory behind compensation, refer to the web site at http://www.drmr.com/compensation/index.html. For reporting of values, see Table 3. The advantages of performing four-color immunophenotyping are 1) the cost is less than with a larger three-color panel; 2) less time is required for specimen processing; and 3) there are fewer tubes to aliquot and handle. The disadvantages are 1) the spectral compensation for four colors is more complicated than for three-color flow cytometry; 2) additional expertise is needed for instrument setup, data collection, and data analysis because the instrumentation is more complex; and 3) the two-tube three-color panel provides duplicate CD3 values for validation whereas the single-tube four-color method does not. When a laboratory considers switching from three to four-color immunophenotyping, there is a need to document the effect (or lack of effect) that this will have on laboratory values. NIAID/DAIDS laboratories that seek approval for implementation of four-color immunophenotyping must demonstrate equivalence of CD3+CD4+ and CD3+CD8+ percentages measured by the proposed four-color method compared with the laboratory's current three-color method. Using their current three-color method and their proposed four-color method, laboratories are required to evaluate 60 different, sequential patient specimens with CD4+ T cells ≤30% as determined by their three-color method. The instructions for performing the three to four-color comparison study are located on the AACTG web site http://aactg.s-3.com/iqa.htm under the heading, “Immunology Quality Assessment Program (IQA).” It is important that the switch study data be reflective of the data generated in the laboratory and not a select subset of comparisons. Laboratories are to keep list mode files from their switch study in case there is a need to look at them for reanalysis. There may be instances where a laboratory wishes to switch directly from two to four-color immunophenotyping. Although NIAID/DAIDS guidelines allow for this switch, experience has shown that laboratories have difficulties demonstrating equivalence because of the dramatic difference in immunophenotypic analyses. Laboratories are much more likely to show equivalence if they perform a stepwise switch from two to three-color and then a three to four-color switch. The single-platform counting technology has emerged as the preferred absolute cell enumeration method for clinical flow cytometric applications (16). The absolute cell subset number is calculated directly from the original blood volume to which suspended microspheres have been added. Briefly, analysis tubes contain a known number of fluorescent beads and a precise volume of blood. By gating the bead population during analysis, absolute cell counts can be determined readily by a simple calculation. The pipetting of blood and beads must be accurate and precise during the application of the single-platform method. Washing of the blood is not allowed because the exact starting volume must be maintained. Advantages of the single-platform internal bead-based methodology for measuring CD3+CD4+ and CD3+CD8+ subset percentages and absolute counts are 1) the precision for absolute cell counts for both fresh and 24-h specimens is superior to those obtained using a two-platform method (5, 6, 17); 2) time is saved with reduced sample manipulation, resulting in a faster turn-around-time for the data; 3) single-platform technology may be more cost-effective because the cost of white blood cells and lymphocyte differential counts is eliminated; and 4) single-platform absolute counts are not affected by gate purity. The disadvantages are 1) the problems with pipetting affect the results to a greater degree with single-platform methods and 2) the flow cytometry laboratory staff must educate patient care providers regarding the advantages of the single-platform methodology because it is new and unfamiliar. When a laboratory considers switching from the conventional two-platform to the more robust single-platform methodology, there is a need to document the effect (or lack of effect) that this will have on laboratory values. It is not unusual to find a bias in the absolute count when such a switch occurs (5). This expected change should be reported to the clinicians so that they are better able to determine whether a change in CD4 count occurred due to improvement in methodology or due to disease progression or a change in therapy. Therefore, NIAID/DAIDS laboratories that seek approval for implementation of single-platform immunophenotyping must perform a switch study and report the CD3+CD4+ and CD3+CD8+ values measured by both the proposed single-platform method and the laboratory's current two-platform method. Laboratories are required to evaluate 30 different patients with a CD4 absolute count <200 using their current two-platform method and their proposed single-platform method. In addition, the laboratory must also evaluate 60 different patients with a CD4 absolute count ≥200 using both their current and proposed method. The instructions for the two-platform to the single-platform absolute count switch study are located on the AACTG web site at http://aactg.s-3.com/iqa.htm under the heading, “Immunology Quality Assessment Program (IQA).” It is important that the switch study data be reflective of the data generated in the laboratory and not a select subset of comparisons. Laboratories are to keep list mode files from their switch study in case there is a need for reanalysis. At present, there are two commercially available suppliers of single-platform bead- based reagents. TruCount tubes are available from BD Biosciences and Flow-Count fluorospheres are available from Beckman Coulter. In principle, it is possible to find some commercially available substitutes for these two brands of fluorospheres. However, they will require validation prior to introduction for clinical cell counting in NIAID/DAIDS-sponsored research projects. As with any laboratory assay, an appropriate quality assurance program must be in effect. Pipettes must be certified on a quarterly basis that they are dispensing accurate and precise volumes. Procedures for pipette calibration are available at http://aactg.s-3.com/iqa.htm. In addition, a stabilized preparation of normal human peripheral blood should be used daily to evaluate mAb binding, volume dispensing by pipettes, and sample analysis. There are several commercial stabilized whole blood preparations available: CD-Chex (Streck Laboratories, Lincoln, Nebraska), Immuno-Trol (Beckman Coulter), FluoroTrol (BioErgonomics, St. Paul, MN), StatusFlow (R&D Systems, Minneapolis, MN), Multi-Check (BD Biosciences), as well as other similar reagents. Manufacturers of these products provide the expected subset percentages and absolute counts. NIAID/DAIDS studies require that both percentages and absolute counts be reported for lymphocyte subsets. Both measurements are clinically important. There are studies that indicate that in some situations CD4 percentage measurements are less variable (18-20) and better predictors of progression to AIDS and death in human immunodeficiency virus (HIV)-infected individuals than the absolute CD4 T-cell counts. In pediatrics, the percentage of CD4 T cells changes less during the first 5 years of life than the absolute CD4 T-cell count (21). Variability due to diurnal variation is also less for percentages of CD4 cells than absolute CD4 counts (22). Likewise, in adults over 60 years of age (23), and also during pregnancy (24), the percentage value changes less. In response to acute exercise, percent values are more stable than absolute CD4 T cells (25). A recent study evaluated the utility of a panel of tests including HIV viral load, white blood cells, and both absolute and percent CD4 as predictors of outcome following surgery in an HIV-infected cohort (26). Only the percent CD4 measurements were observed to be independent predictors of postoperative morbidity. Therefore, most studies suggest that in addition to measuring absolute CD4 T-cell counts, the monitoring of the relative percentages of CD4 cells should be incorporated into any immunological assessment of HIV-infected persons. There are no studies yet available that compare the clinical utility of percentage with the new single-platform methodology absolute count T-cell subset values. One variation of single-platform methodology allows for the enumeration of absolute T-lymphocyte subset counts without using a lymphocyte-gating reagent (27). With this method, it is possible to obtain an absolute CD4 T-cell count in a single tube with a single antibody by counting only cells that stain positively for CD4 with appropriate light scatter parameters. From the point of cost-effective technology, this approach is attractive. However, it has not been validated outside of the original publication or rigorously tested in large clinical trials. Therefore, it is not allowed for the analysis of clinical specimens for domestic NIAID/DAIDS-sponsored studies. Currently, it is recommended that both the percent and absolute counts for T-cell subsets be reported for clinical trials designed to assess the efficacy of new drug and vaccine therapies for HIV-infected individuals. In view of the data already published, The NIAID/DAIDS Immunopohenotyping Guidelines Committee recommends switching to three or four-color imunophenotyping, employing CD45 gating, and switching to single-platform technologies that allow the quantification of both percent and absolute counts." @default.
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• W1982327519 title "Use of CD45 gating in three and four‐color flow cytometric immunophenotyping: Guideline from the national institute of allergy and infectious diseases, division of AIDS" @default.
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