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Guidelines for Performing Single-Platform Absolute
CD4+ T-Cell Determinations with CD45 Gating for Persons Infected
with Human Immunodeficiency Virus
Francis F. Mandy, Ph.D.1 Janet K.A. Nicholson, Ph.D.2 J. Steven McDougal, M.D.3
1Bureau of HIV/AIDS, STD and TB
Population and Public Health Branch
Ottawa, Ontario, Canada
2Office of the Director
and 3Division of AIDS, STD, and TB Laboratory Research
National Center for Infectious Diseases, CDC
The material in this report originated in the National Center for Infectious Diseases, James M. Hughes, M.D., Director; Division of AIDS, STD, and
TB Laboratory Research, Jonathan E. Kaplan, M.D., Acting Director.
These guidelines were developed by CDC for laboratorians who perform immunophenotyping for detection and
enumeration of CD4+ T-cells and other lymphocyte subsets in persons infected with human immunodeficiency virus (HIV). The
guidelines describe single-platform technology (SPT), a process in which absolute counts of lymphocyte subsets are measured from a
single tube by a single instrument. SPT incorporates internal calibrator beads of known quantity in the analysis of specimens
by three- or four-color flow cytometry. With CD45 gating, the relative numbers of beads and lymphocyte subsets are
enumerated, and their absolute numbers and percentage values are calculated. This report supplements previous recommendations
published in 1997 (CDC. 1997 revised guidelines for performing CD4+ T-cell determinations in persons infected with
human immunodeficiency virus [HIV]. MMWR 1997;46[No.
RR-2]) that describe dual-platform technology, a method in
which absolute counts are derived from measurements obtained from two instruments---a flow cytometer and hematology
analyzer. The new recommendations address concerns specific to the implementation of SPT as well as other general topics such
as laboratory safety and specimen handling.
Obtaining accurate and reliable measures of
CD4+ T lymphocytes (CD4+ T cells) is essential to assessing the immune
system and managing the health care of persons infected with human immunodeficiency virus (HIV)
(1--4). The pathogenesis of acquired immunodeficiency syndrome (AIDS) is largely attributable to the decrease in the number of T cells that bear the
CD4 receptor (5--9). Progressive depletion of
CD4+ T cells is associated with an increased likelihood of severe HIV disease and
an unfavorable prognosis (10--13). Accordingly, the U.S. Public Health Service (PHS) has recommended that
CD4+ T-cell levels be monitored every 3--6 months in all HIV-infected persons
(14). Measurement of CD4+ T-cell levels has been used
to establish decision points for initiating prophylaxis for
Pneumocystis carinii pneumonia and other opportunistic infections
and for initiating and monitoring antiretroviral therapy
(15--20). CD4+ T-cell levels are also a criterion for categorizing
HIV-related clinical conditions according to CDC's classification system for HIV infection and surveillance case definition of
AIDS among adults and adolescents (21).
Single-platform technology (SPT) is designed to enable determinations of both absolute and percentage lymphocyte
subset values using a single tube. Until recently, most absolute T-cell numbers were derived from three measurements determined
with two different instruments, a hematology analyzer and a flow cytometer (dual-platform technology [DPT]). Hence, the
CD4+ T-cell number is the product of three laboratory measurements: the white blood cell count, the percentage of white blood
cells that are lymphocytes (differential), and the percentage of lymphocytes that are
CD4+ T cells (determined by flow
cytometry). In 1997, CDC published guidelines addressing concerns related to DPT
(22); those guidelines remain appropriate
for laboratories performing CD4+ T-cell counts with this technology.
On November 14--15, 2001, a third national conference on
CD4+ immunophenotyping was held in Orlando, Florida,
to discuss scientific and technologic advances in the development and production of reagents, instrumentation, and software
that have occurred since publication of the 1997 guidelines. The conference was attended by representatives from public
health, private, and academic laboratories as well as product manufacturers. These guidelines reflect a consensus of that
conference, reviewed by attendees, and specifically related to the performance of SPT.
Development of new guidelines was driven by advances in knowledge and experience with new approaches to
enumerate CD4+ T cells. First, a gating strategy for identifying lymphocytes using CD45 fluorescence and side-scattering characteristics
is now the preferred method for identifying lymphocytes accurately and reproducibly. Second, three- or four-color
flow cytometry has been demonstrated to be superior to two-color methods for measuring
CD4+ and CD8+ T-cell counts
(23). Finally, the availability of Food and Drug Administration (FDA)-approved commercial microfluorosphere counting
reagents for SPT has resulted in decreased interlaboratory variability
(24,25). Consequently, SPT is the preferred method in
an increasing number of laboratories (4).
I. Laboratory Safety
A. Use universal precautions with all specimens
B. Adhere to the following safety practices
Wear laboratory coats and gloves when processing and analyzing specimens, including reading specimens on
the flow cytometer.
Never pipette by mouth. Use safety pipetting devices.
Never recap needles. Dispose of needles and syringes in puncture-proof containers designed for this purpose.
Handle and manipulate specimens (e.g., aliquot, add reagents, vortex, and aspirate) in a class I or II
biological safety cabinet.
Centrifuge specimens in safety carriers.
After working with specimens, remove gloves and wash hands with soap and water.
For stream-in-air flow cytometers, follow the manufacturer's recommended procedures to eliminate
the operator's exposure to any aerosols or droplets of sample material.
Disinfect flow cytometer wastes. Before adding waste materials to the waste container, add a sufficient volume
of undiluted household bleach (5% sodium hypochlorite) so that the final concentration of bleach will be
10% (0.5% sodium hypochlorite) when the container is full (e.g., add 100 mL of undiluted bleach to an
empty 1,000-mL container).
Disinfect the flow cytometer as recommended by the manufacturer. One method is to flush the flow
cytometer fluidic chambers with a 10% bleach solution for 5--10 minutes at the end of the day and then flush with
water or saline for at least 10 minutes to remove excess bleach, which is corrosive.
Disinfect spills with household bleach or an appropriate dilution of mycobactericidal disinfectant.
Note: Organic matter will reduce the ability of bleach to disinfect infectious agents. NCCLS
recommendations regarding how to disinfect specific areas should be followed
(30). For use on smooth, hard surfaces, a
1% solution of bleach is usually adequate for disinfection; for porous surfaces, a 10% solution is needed
Ensure that all samples have been properly fixed after staining and lysing but before analysis.
Note: Some commercial reagents employ a single-step, lyse and fix method that reduces the infectious activity of
cell-associated HIV by 3--5 logs (31,32); however, these reagents have not been evaluated for their
effectiveness against other agents (e.g., hepatitis virus). Cell-free HIV can be inactivated with 1% paraformaldehyde
within 30 minutes (33--35).
II. Specimen Collection for Single-Platform
Use tripotassium ethylenediamine tetraacetate
(K3EDTA, 1.5 ± 0.15 mg/mL blood) or heparin
(36--39), and perform the test within the time frame allowed by the SPT manufacturer. Because acid citrate dextrose is
as a liquid to blood collection tubes, its use would make calculating accurate final sample volume difficult and
is not recommended. With this absolute counting technology, use of an accurate sample volume is critical.
Reject specimens that cannot be processed within 72 hours.
B. Collect blood specimens by venipuncture
(40) into evacuated tubes containing
K3EDTA anticoagulant, completely expending the vacuum in the tubes.
Use pediatric tubes to obtain specimens from children, and ensure that the tube is full.
Mix the blood well with the anticoagulant to prevent clotting.
C. Label all specimens with the date, time of collection, and a unique patient identifier. Ensure that patient
information and test results are accorded confidentiality.
III. Specimen Transport
Maintain and transport specimens at room temperature (64°--72°F [18°--22°C])
(39,41--43). Specimens should not be exposed to extreme temperatures that could allow them to freeze or become too hot. Temperatures >99°F
(37°C) might cause cellular destruction and affect flow cytometry measurements
(39). In hot weather, pack the specimen in an insulated container. If necessary, place this container inside another containing an ice pack and
absorbent material. This method helps retain the specimen at ambient temperature. The effect of cool temperatures (i.e.,
<39°F [4°C]) on CD45 gate-based immunophenotyping results is not clear
Transport specimens to the immunophenotyping laboratory as soon as possible.
For transport to locations outside the collection
facility, follow state or local guidelines. One method for
packaging such specimens is to place the tube containing the specimen in a leakproof container (e.g., a sealed plastic bag)
and to pack this container inside a cardboard canister containing sufficient material to absorb all the contents should
the tube break or leak. Cap the canister tightly. Fasten the request slip securely to the outside of this canister with
a rubber band. For mailing, this canister should be placed inside another canister containing the mailing label.
For interstate shipment, follow federal guidelines for transporting diagnostic specimens (available at
Note: Use overnight carriers with an established record of
consistent overnight delivery to ensure arrival the following day. Check with these carriers for their specific
Obtain specific protocols and arrange appropriate times of collection and transport from the facility collecting
IV. Specimen Integrity
Inspect the tube and its contents immediately upon arrival.
Take corrective actions if any of the following
If the specimen is hot or cold to the touch but not obviously hemolyzed or frozen, process it but note
the temperature condition on the worksheet and report form. Do not rapidly warm or chill specimens to bring
them to room temperature because this may adversely affect the immunophenotyping results
(39). Abnormalities in light-scattering patterns may reveal a compromised specimen.
If blood is hemolyzed or frozen, reject the specimen and request another.
If clots are visible, reject the specimen and request another.
If the specimen is received >72 hours after collection, reject it and request another.
V. Specimen Processing
Perform the test within 48 hours (preferred), but no later than 72 hours after drawing the blood specimen
Place the samples on a gentle blood rocker for 5 minutes to ensure that the samples are uniformly distributed.
Pipette blood volumes accurately and in a reproducible manner. A reverse pipetting technique is
Vortex sample tubes to mix the blood and reagents and break up cell aggregates. In addition, vortex
samples immediately after the lyse/fixation step and before analysis to disperse cells optimally.
Incubate all tubes in the dark during the staining procedure.
A lyse/no-wash method is required for SPT. Follow directions provided by the manufacturer.
Immediately after processing the specimens, cap the tubes and store all stained samples in the dark and
under refrigeration (39°--50°F [4°--10°C]) until flow cytometric analysis. These specimens should not be stored for
longer than 24 hours unless the laboratory can demonstrate that scatter and fluorescence patterns do not change
for specimens for stored longer periods.
VI. Monoclonal Antibody Panels
CD45 is required to aid in the identification of lymphocytes. Lymphocytes are brightly positive for CD45 and
have low light-scattering characteristics.
Monoclonal antibody panels must contain appropriate antibody combinations to enumerate
CD4+ and CD8+ T-cells and to ensure the quality of the results (Table 1).
CD4 T-cells are identified as being positive for CD3 and CD4.
CD8 T-cells are identified as being positive for CD3 and CD8.
Three-color monoclonal antibody panels
Three-color monoclonal antibody panels should fulfill the following basic requirements: enumerate
CD4+ and CD8+ T-cells, validate the CD45 gate used, and provide some assessment of tube-to-tube variability.
Three-color monoclonal antibody panels must consist of at least two tubes, each with the same lineage
marker. For the examples described previously, CD3 is the common lineage marker in each tube. Differences
between replicate CD3 results should be
CD19+ B-cell values may be important in
assessing immune status of pediatric patients.
Four-color monoclonal antibody panels
Addition of CD45 to a single tube containing CD3, CD4, and CD8 allows the identification of
lymphocytes based on CD45 and side scatter and the enumeration of
CD4+ and CD8+ T-lymphocytes.
CD19+ B-cell values may be essential for
assessing the immune status of pediatric patients.
Use of a second tube containing a natural killer (NK) cell marker together with CD3 and CD19 can help
to assess the recovery and purity of the lymphocytes within the CD45/side-scatter gate.
VII. Negative and Positive Controls for Immunophenotyping
Note: An isotype control is not needed.
Positive methodologic control
Use the methodologic control to determine whether procedures for preparing and processing the specimens
are optimal. Prepare this control each time specimens from patients are prepared.
Use either a whole-blood specimen from a control donor or commercial materials validated for this purpose.
If the methodologic control falls outside established normal ranges, determine the reason.
Note: The purpose of the methodologic control is to detect problems in preparing and processing the specimens. Biologic factors
that cause only the whole-blood methodologic control to fall outside normal ranges do not invalidate the results
from other specimens processed at the same time. Poor lysis or poor labeling in all specimens, including
the methodologic control, invalidates results.
Positive control for evaluating reagents
Use the positive control to test the labeling efficiency of new lots of reagents or when the
labeling efficiency of the current lot is questioned. Prepare this control only when needed (i.e., when reagents are in question)
in parallel with reagent lots of known acceptable performance.
Note: New reagents must demonstrate similar results to those of known acceptable performance.
Use a whole-blood specimen or other human lymphocyte preparation (e.g., cryopreserved or
commercially obtained lyophilized lymphocytes or stabilized whole blood).
VIII. Flow Cytometer Quality Control
Verify optical alignment daily. Usually, clinical flow cytometers that are capable of three- and
four-color immunophenotyping have fixed optical systems, i.e., the relative position of the flow cell with respect to the
elements is fixed. In such systems, the instrument operator cannot optimize alignment but must verify that
the instrument meets the manufacturer's specifications for optical alignment. Regardless of whether the alignment is
user adjustable, it should be checked with alignment standards, such as wide-spectrum fluorescent
microfluorospheres with measurable light-scatter characteristics. Daily monitoring of optical alignment ensures that the cytometer
gives acceptably bright fluorescence measurements and that homogeneous peaks are produced for all parameters to be
used in sample analysis (45).
Use a stable calibration material (e.g., microfluorospheres labeled with fluorochromes) that has measurable
and known forward-scatter, side-scatter, and fluorescence properties in each channel to be used for sample analysis.
Verify acceptable optical alignment by establishing that calibration particles meet manufacturer- or
laboratory-defined criteria for brightness and homogeneity.
Align stream-in-air flow cytometers daily (at a minimum) and stream-in-cuvette flow cytometers (most
clinical flow cytometers are this type) as recommended by the manufacturer.
Standardize fluorescence and light-scatter signals daily. This ensures that the flow cytometer is operating
within manufacturer- or laboratory-defined acceptance ranges under test-specific conditions each day and that
its performance is consistent from day to day.
Select machine settings that are appropriate for antibody/fluorochrome-labeled, whole-blood specimens.
Use microfluorospheres or other stable standardization material to place the scatter and fluorescence peaks in
the same narrow range of scatter and fluorescence channels each day.
Adjust the flow cytometer as needed.
Retain machine standardization settings for the remaining quality control procedures (sensitivity and
color compensation) and for reading the specimens.
Determine fluorescence resolution daily. The flow cytometer must differentiate between the dim peak
and autofluorescence in each fluorescence channel
Unstained and lysed fresh whole blood is suitable for adjusting the photomultiplier tube (PMT) voltages.
The autofluorescence from the unstained lymphocytes should be completely on scale (i.e., <5% of cells within
the lymphocyte light-scatter gate fall in channel 0 in each fluorescence scale) and should fall within the lower
left quadrant of the dot plot for every PMT/detector in use.
Evaluate standardization/calibration material or cells to verify that cells with low-level fluorescence can
be resolved from autofluorescence (e.g., microbeads with low-level and negative fluorescence,
CD56-labeled lymphocytes, or dim cells in CD8-labeled lymphocytes).
Establish a minimal acceptable distance between peaks; monitor this difference, and correct any daily deviations.
Compensate for spectral overlap daily (Figure 1). Compensation is the process of correcting for spectral overlap
of one fluorochrome into the filter window being used to monitor another fluorochrome. In most instruments
used clinically, this correction is done by adjusting the electronic compensation circuits on the flow cytometer to
place populations not expected to be dual positive for two fluorochromes into orthogonal fluorescence quadrants with
no overlap into the double-positive quadrant. At the same time, avoiding overcompensation is
essential because this may cause dual-positive cells to be incorrectly classified as single positive. The following procedures may be
performed manually, or the software on the flow cytometer may perform the spectral compensation automatically.
Select the compensation control so it will match the brightest specimen signal. Use either microbead or
cellular compensation material containing four populations for three-color immunofluorescence (no
fluorescence, phycoerithrin [PE] fluorescence only, fluorescein isothiocyanate [FITC] fluorescence only, and a population
that is positive for only the third color) or five populations for four-color (the four described previously and
a population that is positive for only the fourth color).
Analyze this material, and adjust the electronic compensation circuits on the flow cytometer to place
the fluorescent populations in their respective fluorescence quadrants with no overlap into the
double-positive quadrant (Figure 1). With three fluorochromes, compensation must be carried out in an appropriate
sequence: FITC, PE, and the third color, respectively
(46). For four-color monoclonal antibody panels, follow the
flow cytometer manufacturer's instructions for four fluorochromes. Avoid overcompensation.
If standardization or calibration particles (microbeads) have been used to set compensation, confirm
proper calibration by using lymphocytes labeled with FITC- and PE-labeled monoclonal antibodies and a
or fourth-color-labeled monoclonal antibody for three-color or four-color panels, respectively. So that
separate cell populations can be recognized without overlap, cells in individual tubes may be separately stained with
each different fluorochrome-labeled antibody and then combined in a single tube for analysis. These
populations should have the brightest expected signals.
Note: Using a dimmer-than-expected signal to set compensation
can result in suboptimal compensation for the brightest signal.
Reset compensation when photomultiplier tube voltages or optical filters are changed.
Commercially available software can analyze data without compensation and perform the
compensation automatically. When using this software, follow manufacturer's instructions for this procedure.
Repeat all four instrument quality control procedures (section VIII A--C) whenever instrument problems occur or
if the instrument has been serviced.
Maintain instrument quality control logbooks and monitor them continually for changes in any of the
parameters. In the logbook, record instrument settings, peak channels, and coefficient of variation (CV) values for materials
used to monitor or verify optical alignment, standardization, fluorescence resolution, and spectral
compensation. Reestablish target fluorescence levels for each quality control procedure when lot numbers of beads are changed
or the instrument has been serviced.
IX. Sample Analyses
With single-platform absolute count determination, use of the lyse/no-wash sample processing is mandatory.
The lymphocyte population is identified as having bright CD45 fluorescence and low side-scattering properties
(Figure 2). Set the threshold or discriminator as recommended by the manufacturer.
Adjust side scatter so that all leukocyte populations are visible. Draw a gate on the bright
CD45+ cell population and analyze the cells in that population
Count at least 2,500 gated lymphocytes in each sample to ensure that enough cells and beads have been counted
to provide an accurate absolute lymphocyte value.
X. Data Analysis
A. CD45 gating
Lymphocytes are identified by being brightly labeled with CD45 monoclonal antibody and
having low side-scattering properties. Two typical examples of a four-color SPT analysis based on CD45 gating are illustrated
Establish criteria for cluster identification based on a clear definition of lymphocytes that does not
include basophils (less bright CD45, low side scatter) or monocytes (less bright CD45, moderate side scatter).
Note: Care must be taken to include all lymphocytes. CD45 fluorescence may be slightly less with B cells than with
T cells (the major cluster of lymphocytes). NK cells have bright CD45 fluorescence but have slightly more
side-scattering properties than the majority of the lymphocytes.
CD45/side-scatter gates for lymphocytes are assumed to contain >95% lymphocytes. Lymphocyte purity
is assumed to be high with the CD45/side-scatter gating strategy; therefore, correction of lymphocyte subset
values is not needed (47).
If an estimate of lymphocyte recovery is needed (i.e., percentage of total lymphocytes within the
CD45/side scatter gate), all the B and NK cells must be immunophenotyped as well.
Note: Validation of a CD45/side-scatter gate is recommended during its initial use to help determine the CD45 and side-scatter characteristics
of T, B, and NK cells and to ensure their inclusion in the gate.
Set cursors based on the tube containing CD3/CD4 and CD3/CD8 so that the negative and positive cells in
the histogram are clearly separated.
Analyze each patient or control specimen with lymphocyte gates and cursors for positivity set for that
particular patient or control.
Include the following analytic reliability checks, when available:
With SPT, an additional analytical tool can be used to check the accuracy of the absolute count; time can
be used as a parameter to determine how long it takes to obtain a microfluorosphere count that represents a
unit volume of blood analyzed. Optimally, if blood pipetting was performed without noticeable error and the
were accurately added to the tubes, the time required to analyze a microliter of whole blood should be
constant. Follow manufacturer's instructions to set time as an active parameter. If more or less time is required for
a sample to accumulate the usual number of microspheres, this may indicate a serious counting problem
and specimen processing should be repeated.
Optimally, the sum of the percentages of
CD3+CD8+ cells should equal the total percentage
of CD3+ cells + 5%, with a maximum variability of <10%.
Note: For specimens containing a considerable
number of T T-cells (48,49), this reliability check may exceed the maximum variability.
XI. Data Storage
Store list-mode data for all specimens analyzed. This allows for reanalysis of the raw data, including redrawing
of gates. At a minimum, retain hard copies of the CD45/side-scatter gate and correlated dual-histogram data of
each sample's fluorescence.
Retain all primary files, worksheets, and report forms for 2 years or as required by state or local regulation,
whichever is longer. Data can be stored electronically. Disposal after the retention period is at the discretion of the
XII. Data Reporting
Report all data in terms of CD designation, with a short description of what that designation means.
Note: CD4+ T cells are T-helper cells. The correct cells to report for this value are those that are positive for both CD3 and
CD4. Similarly, CD8+ T-cells are T-suppressor/cytotoxic cells and are positive for both CD3 and CD8. Do not
include other cell types (non-T cells) in CD4 and CD8 T-cell determinations.
Report lymphocyte subset values as follows:
Report both percentages and absolute counts.
With SPT, determine the absolute counts directly from the flow cytometers. These calculations are
usually handled by software that reports calculated results. The following formula should be used:
Report data from all relevant monoclonal antibody combinations with corresponding reference limits of
expected normal values (e.g., CD4+ T-cell absolute number and percentage). Reference limits for immunophenotyping
test results must be determined for each laboratory
(45). Separate reference ranges must be established for adults
and children, and the appropriate ranges must be reported for patient specimens.
XIII. Quality Assurance
Ensure the overall quality of the laboratory's
CD4+ T-cell testing by monitoring and evaluating the effectiveness
of the laboratory policies and procedures for the preanalytic, analytic, and postanalytic testing phases. The practices
and processes to be monitored and evaluated include the following:
methods for collecting, handling, transporting, identifying, processing, and storing specimens;
information provided on report forms for test requests and results;
instrument performance, quality control protocols, and maintenance;
reagent quality control protocols;
process for reviewing and reporting results.
employee training and education, which should consist of the following:
--- basic training by flow cytometer manufacturers and additional training involving hands-on workshops
for flow cytometer operators and supervisors;
--- education of laboratory directors regarding flow cytometric immunophenotyping through workshops
and other programs;
--- continuing education regarding new developments for all flow cytometric immunophenotyping
personnel through meetings and workshops;
--- adherence to federal and state regulations for training and education;
assurance of satisfactory performance. Laboratories must fully participate in a performance evaluation
program and demonstrate acceptable level of performance. When proficiency testing programs have been approved by
the Centers for Medicare & Medicaid Services (formerly, the Health Care Financing Administration) as meeting
requirements of the Clinical Laboratory Improvement Amendments of 1988 (CLIA '88) (none are
currently approved for CD4+ T-cell testing), laboratories must satisfactorily participate.
review and revision (as necessary or at established intervals) of the laboratory's policies and procedures to
ensure adherence to the quality assurance program. All staff involved in the testing should be informed of any
problems identified during the quality assurance review, and corrective actions should be taken to prevent recurrences.
Document all quality assurance activities.
Evaluation and Validation of a Newly Adopted SPT in the Laboratory
When a laboratory adopts the new SPT, specimens should be tested in parallel by using both the current and the
new method to characterize any systematic differences in the methods. Laboratorians should use statistical tools that provide
useful information for the comparison studies. Linear least squares regression analyses are helpful in establishing good
correlations between the new and established methods. If no error is detected with the new method, the
r2 value will approach 1.0. However, regression-type scatter plots provide inadequate resolution when the errors are small in comparison with
the analytical range and do not characterize the relationship between the two methods
A bias scatter plot provides laboratorians with a more useful tool for determining bias. These simple, high-resolution
graphs plot the differences in the individual measurements of each method (result of old method---result of new method)
against measurements obtained with one of the methods (result of old method)
(50). Such graphs provide an easy means of determining if bias is present and distinguishing whether bias is systematic, proportional, or random/nonconstant.
The laboratorian can visually determine the magnitude of these differences over the entire range of values. When sufficient
values are plotted, outliers or samples containing interfering substances can be identified. The laboratorian can then divide the
data into ranges relevant to medical decisions and calculate the systematic error (mean of the bias) and the random error
(standard deviation of the bias) to gain insight into analytical performance at the specified decision points
Several detailed guidelines and texts provide additional information regarding quality goals, method evaluation,
estimation of bias, and bias scatter plots
(50--54). Once a new method is accepted and implemented, the laboratory will need to
confirm or redefine its normal range and should continue to monitor the correlation between the results and the patient's
clinical disease data to ensure that no problems have gone undetected by the relatively few samples typically tested during
More than 1.6 million CD4+ T-cell measurements are performed yearly by the approximately 600 testing laboratories in
the United States (55). This figure is based on the reported number of tests performed annually by laboratories participating
in CDC's Model Performance Evaluation Program (MPEP) for T-lymphocyte immunophenotyping in 1996.
These measurements are performed with flow cytometers using either multiplatform technology or SPT. SPT was introduced
for clinical application in 1996, and its wide-scale implementation is relatively new. In 2000, results of two
independent multicenter studies studies of SPT were reported
(24,25). Those and subsequent reports on SPT and CD45 gating
(56--60) have increasingly encouraged adoption of these improved testing practices
(61,62). The resulting outcomes associated
with SPT and CD45 gating include a) increased confidence in results, b) more reproducible results, c) increased ability to
resolve discrepant problems, d) decreased proportion of unacceptable specimens received for testing, e) decreased proportion
of specimens requiring reanalysis, and f) fewer incidents that could pose biohazard risks
Although these guidelines for SPT use might foster improved laboratory practices, developing comprehensive guidelines
for every aspect of CD4+ T-cell testing (including some laboratory-specific practices) is not possible. Moreover, measuring
the outcomes associated with the adoption of these guidelines is inherently difficult. First, the guidelines lack evaluation
protocols that can adequately account for the interactions among the recommendations. No weight of importance has been assigned
for the individual recommendations that address unique steps in the testing process; hence, the consequences of
incompletely following the entire set of recommendations are uncertain. Second, because published data are not available for every aspect
of the guidelines, certain recommendations are based on the experience and opinion of knowledgeable
persons. Recommendations made on this basis, in the absence of data, may be biased and inaccurate. Finally, variations in
practices and interactions among the practices (e.g., how specimens are obtained and processed, skill of laboratory
personnel [such as with pipetting], testing methods used, test-result reporting practices, and compliance with other voluntary
standards and laboratory regulations) complicate both the development of guidelines that will fit every laboratory's unique
circumstances and the assessment of the value of implementing the guidelines.
The first CDC recommendations for laboratory performance of
CD4+ T-cell testing (63) were written so as not to
impede development of new technology or investigations into better ways to assess the status of the immune system in
HIV-infected persons. Developments in the technology have resulted in an assay that is technically less complicated and more
accurate. These single-platform methods are now being implemented in as many as one fourth of the laboratories in the United
States (MPEP data). In addition, other T-cell phenotypic markers are being investigated as prognostic indicators or markers
of treatment efficacy, alone and in combination with other cellular markers
These guidelines for SPT are intended for domestic implementation. Several alternative methods are available that
require fewer reagents and involve more cost-effective gating algorithms. Some of these alternative methods may be compatible
with current U.S. clinical laboratory methods; however, to date they have not been validated for domestic applications. As
published validation data accumulate from multisite studies for methods such as
PanLeucogating (66) and primary CD4 gating
(67, 68), these potentially more cost-effective options will be considered as alternative or substitute methods. In the future,
guidelines should be harmonized to include all methods that meet domestic performance standards to ensure consistent high quality.
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