Absolute CD4 Counts Obtained by a Three-Color Flow-Cytometric Method without the Use of a Hematology Analyzer

Abstract

We evaluated the Ortho TRIO-Cytoronabsolute system for determining absolute CD4 counts. The CD4 counts in our blood specimens from 100 individuals ranged from 3 to 1,962; the percent CD4 ranged from 1.3 to 62.2, respectively. The TRIO system was biased toward lower absolute counts than a combination of flow cytometry and hematology but showed no bias in percent CD4 calculations.

The stage of human immunodeficiency virus (HIV) disease is determined by the number of CD4-positive T cells in an individual’s blood. This value is normally obtained by determining the percentage of lymphocytes which coexpress CD3 and CD4 by flow cytometry and multiplying that value by the absolute lymphocyte count, as determined by a hematology analyzer (dual-platform method). Single-platform flow-cytometric systems which directly determine absolute CD4 counts have been introduced. These systems include the Ortho TRIO panel, BD FACScount and TrueCount, and Coulter XL. Nicholson et al. (6) recently reviewed the TrueCount system and found that this system provided CD4 counts which averaged 11.3% higher than those obtained by a dual-platform method. The TRIO panel, along with the Ortho Cytoronabsolute flow cytometer, is a Food and Drug Administration (FDA)-approved method for determining absolute CD4 counts (3). This method was recently validated in a multicenter trial with specimens from healthy individuals and found to produce precise CD4 counts averaging 15% lower than those obtained by a dual-platform method (2). The Cytoronabsolute flow cytometer is a syringe-driven cytometer, which may be calibrated for determining the total absolute lymphocyte count (3). The software multiplies the total lymphocyte count by the percentage of the desired subset, e.g., CD4-positive T cells, obtained by immunofluorescence analysis. We evaluated the ability of the TRIO-Cytoronabsolute system to determine absolute CD4 counts in HIV-infected individuals, non-HIV-infected transplant recipients, and control patients who had not been tested for HIV.

We collected peripheral blood specimens from 100 individuals into both acid-citrate-dextrose (ACD) and EDTA tubes. Our sample consisted of patients who were presenting for immunophenotyping and healthy controls. Institutional review board approval was not necessary for this study because we were performing assays which physicians had ordered on the patients.

Specimens collected in ACD (our standard anticoagulant for two-color immunophenotyping) tubes were stained with the two-color antibodies (Coulter Corp., Hialeah, Fla.) listed in Table 1. This panel yields CD4 results comparable to those of the six-tube CDC panel for both healthy and HIV-infected individuals (1, 4). We incubated whole blood with antibody for 20 min at room temperature. Erythrocytes were lysed and leukocytes were fixed by using the Q-prep instrument (Coulter).

TABLE 1.

TRIO and two-color antibody panels^a

Antibody combination	FITC	PE	Cy-5PE
TRIO-1	IgG1	IgG1	IgG1
TRIO-2	CD4	CD8	CD3
TRIO-3	CD16	CD19	CD3
Two-color-1	IgG1	IgG1
Two-color-2	CD3	CD4
Two-color-3	CD3	CD8
Two-color-4	CD19
Two-color-5	CD16

^aAbbreviations: FITC, fluorescein isothiocyanate; PE, phycoerythrin; Cy-5PE, allophycocyanin 5–phycoerythrin; IgG1, immunoglobulin G1. 

Specimens collected in EDTA (anticoagulant required for the FDA-approved TRIO procedure) were stained with the TRIO reagents (Ortho Cytometry, Raritan, N.J.) listed in Table 1. We incubated 100 μl of EDTA-treated blood with 10 μl of antibody for 20 to 30 min at room temperature in the dark. Two milliliters of Ortho Lysing Solution (an ammonium chloride lysing reagent) was added with gentle mixing. The tubes were incubated for 15 min to ensure complete erythrocyte lysis. Specimens were kept in the dark and analyzed within 1 h of completion of the lysis step. Cells analyzed by the TRIO method for absolute counts cannot be fixed (3). Thus, samples must be analyzed within 1 h of processing and infectious particles will not be inactivated. The Cytoron system has a completely enclosed sample-handling system and an autoloader which will minimize exposure to infectious particles. Waste is collected into a container which may contain bleach.

All specimens were analyzed on the Ortho Cytoronabsolute cytometer. At least 2,000 cells in the lymphocyte gate were counted. The two-color tubes were gated on lymphocytes based on light scatter, and the percentage of cells staining for both CD3 and CD4 was determined. This value was multiplied by the absolute lymphocyte count determined from EDTA-containing tubes analyzed on a Sysmex NE 8000 hematology analyzer (Toa, Inc., Chicago, Ill.) to yield an absolute CD4 lymphocyte count. The TRIO tubes were analyzed by using Ortho’s ImmunoCount II software. Appropriate counting times for absolute numbers were determined with Ortho Validation beads. Lymphocytes were detected by both light scatter and immunoscatter gates. Immunoscatter gates use an immunofluorescence marker and right-angle scatter to define a population (3, 5). Isotype controls were used for both the two- and three-color methods to set positive-negative discriminators at a 2% background level. The TRIO panel is validated by using four levels of quality control (QC), including tube-to-tube consistency, Immunosum (lymphosum), T sum, and background fluorescence. We had five specimens initially fail a QC parameter; four were corrected by regating, and one was reanalyzed by using new tubes and was probably due to a pipetting error. The TRIO panel does require accurate, precise pipetting for all QC parameters to be acceptable.

Our sample included 63 specimens with absolute CD4 counts of <500 by both methods, and 37 specimens with absolute CD4 counts of ≥500 by at least one method. The overall range of absolute CD4 counts was 3 to 1,962/mm³ (Sysmex). The absolute CD4 counts obtained by the Cytoron system correlated well with those obtained by the dual-platform method (r² value of 0.99 [Fig. 1]), although the Cytoronabsolute cytometer showed a bias of −9.5% compared to the combination of flow cytometry and hematology (Fig. 2). This bias ranged from −7% for CD4 counts of >500 to −11% for CD4 counts of <100. The percentages of CD4 by the two methods correlated well (r² value of 0.95 and bias of <1% [Fig. 3 and 4]).

FIG. 1.

Absolute CD4 counts obtained by the TRIO method and the dual-platform method. The equation and r² value represent the results of linear regression analysis.

FIG. 2.

Absolute CD4 count bias plot.

FIG. 3.

CD4 percentages obtained by the TRIO method and by the two-color method. The equation and r² value represent the results of linear regression analysis.

FIG. 4.

Percent CD4 bias plot.

Our results indicate that the three-color TRIO panel provides CD4 percentages identical to those by a two-color flow-cytometric method. The percentages for other lymphocyte phenotypes also correlated well with those by the two-color method (data not shown). The absolute counts delivered by the single-platform Cytoronabsolute system, however, have a bias toward lower numbers compared to results with the dual-platform method. These data confirm and extend data presented by Connelly et al. (2). They showed absolute CD4 counts determined by the Cytoronabsolute cytometer in specimens from healthy individuals were routinely 15% lower than those obtained by a combination of flow cytometry and hematology (2). We have shown less bias across a wider range of CD4 counts and included specimens from HIV-infected individuals. The difference in bias values may be due to different hematology analyzers being employed in Connelly’s study.

It is impossible to determine if either a single-platform or dual-platform result is “right” or “wrong,” due to different methods of determining total lymphocyte count with the hematology instrument and the Cytoronabsolute cytometer. The Sysmex analyzer determines absolute counts by using direct-current measurements for cell size and volume and radio wave scatter for cell complexity. The flow cytometer determines total lymphocytes by light scatter and immunofluorescence characteristics on a specimen in which erythrocytes have been lysed. Technical errors, as described by Nicholson et al. (6), cannot be ruled out completely; however, all instruments and pipettors were calibrated, and the assays were performed by experienced technologists. When we switched from the dual-platform method to the single-platform method for our clinical reporting, we provided both sets of numbers on patients we had been monitoring by the combination method. This allowed the clinician to have a number from the dual-platform method to compare to previous values for guiding current treatment and a new baseline number for our single-platform method which would be used to follow the patient from that time forward. An alternative method would be to constantly correct reported values for the observed bias, as previously described by Nicholson et al. (5).

In conclusion, the TRIO-Cytoronabsolute system does provide CD4 percentages which correlate well with those by a two-color method across a wide range of CD4 levels. The single-platform method does provide absolute counts biased toward a lower number than those by the combination of flow cytometry and hematology. The software provides extensive QC, including lymphosum, T sum, and tube-to-tube variation analysis (3). A laboratory switching to a single-platform method must perform in-house experiments to determine if bias is present (6) and to ensure that reports to physicians account for that bias during the transition period.

(Part of this work was presented at the 8th Annual Meeting of the Association of Medical Laboratory Immunologists.)