Considerations of Flow Cytometric Lymphocyte Subset Analysis in Korea Based on a Survey of Current Clinical Laboratory Practice

Abstract

Flow cytometric lymphocyte subset analysis (FCLSA) is essential for assessing immune status across various diseases and clinical settings. We surveyed current clinical laboratory practices related to FCLSA to establish a baseline reference for future standardization in Korea. Nine university hospitals actively performing FCLSA responded to the 22-question survey, which covered seven categories of laboratory practice. These hospitals used commercial reagent antibody kits from either Beckton Dickinson Biosciences (N=4) or Beckman Coulter Diagnostics (N=5). Most hospitals performed daily instrument setup and scheduled maintenance every 2–6 months. Two levels of commercial quality control materials were routinely used each day. Sample and reagent antibody volumes varied across hospitals, even when the same reagent kit was used. Acquired cell counts ranged from 5×10³ to 5×10⁴ cells, with two hospitals adjusting counts based on the cell type analyzed. Most laboratories reported percentages and general opinions; some additionally reported white blood cell and lymphocyte counts, along with lymphocyte percentages. This is the first comprehensive survey on the clinical laboratory practice of FCLSA in Korea. Standardization of FCLSA should be accelerated to ensure reliable and reproducible results.

Body

The discovery of the human immunodeficiency virus in the 1980s accelerated the clinical application of flow cytometry (FCM) for cluster of differentiation (CD)4⁺ T cells [1, 2]. Since then, flow cytometric lymphocyte subset analysis (FCLSA) has been widely used to assess immune status and diagnose lymphoid neoplasms [1–4]. We surveyed current clinical laboratory practice of FCLSA to understand the prevailing status and establish a baseline reference for future standardization in Korea.

Among over 50 hospitals implementing FCLSA in Korea, nine representative university hospitals were selected based on the number of FCM tests conducted, licensed bed count, and geographic distribution. The FCM study group, comprising laboratory medicine specialists from the Korean Society of Diagnostic Immunology, designed and critically reviewed the survey questionnaire. The survey comprised 22 questions in seven categories (Supplemental Data Table S1).

Four hospitals used flow cytometers (FCRs) from Beckton Dickinson Biosciences (BD) (BD, San Jose, CA, USA), and five used FCRs from Beckman Coulter Diagnostics (BC) (BC, Miami, FL, USA) (Table 1). Eight hospitals performed daily instrument setup, and one hospital conducted weekly setup. Compensation was performed every two months by six hospitals, and monthly, daily, or every six months by the remaining three. Eight hospitals performed maintenance at regular intervals of 2–6 months, and one hospital performed maintenance as needed.

Table 1. Result summary of the survey of nine university hospitals performing FCLSA in Korea.

Variables	A	B	C	D	E	F	G	H	I
1. Instrument
Model name*(Manufacturer)	FACSLyric (BD)	FACSCanto II (BD)	FACSCanto II (BD)	FACSLyric (BD)	Navios EX (BC)	Navios EX (BC)	Navios (BC)	Navios (BC)	Navios (BC)
Instrument setup
Alignment
Material	BD CS&T Beads	BD CS&T Beads	BD CS&T Beads	BD CS&T Beads	BC Flow-Check	BC Flow-Check	BC Flow-Check	BC Flow-Check	BC Flow-Check
Interval	Daily	Daily	Weekly	Daily	Daily	Daily	Daily	Daily	Daily
PMT voltages
Material	BD CS&T Beads	BD CS&T Beads	BD CS&T Beads	BD CS&T Beads	BC Flow-Check	BC Flow-Check	BC Flow-Check	BC Flow-Check	BC Flow-Check
Interval	Daily	Daily	Weekly	Daily	Daily	Daily	Daily	Daily	Daily
Compensation
Material	BD CS&T Beads (verification)	BD FACS 7-Color Setup Beads	BD FACS 7-Color Setup Beads	BD CS&T Beads (verification)	BC Flow-Check (verification)	BC Flow-Check (verification)	BC Flow-Check (verification)	BC Flow-Check (verification)	BC Flow-Check (verification)
Interval	Daily	Monthly	Daily	Daily	Daily	Daily	Daily	Daily	Daily
Material	BD FC Beads 2, 5, 7- color			BD FC Beads 12-color	BC CYTO-COMP Cell	BC CYTO-COMP Cell	BC CYTO-COMP Cell	BC CYTO-COMP Cell	BC CYTO-COMP Cell
Interval	2 months			2 months	6 months	2 months	2 months	2 months	2 months
Maintenance interval	3 months	6 months	6 months	As necessary	6 months	2 months	2 months	2 months	2 months
2. Technician†
Dedicated technician	Yes -rotated	Yes -rotated	Yes	Yes	Yes -rotated	Yes -rotated	Yes -rotated	Yes	Yes -rotated
Training/education	Yes -laboratory’s own training at rotation	Yes -laboratory’s own training at rotation -if necessary	Yes -laboratory’s own training	Yes -laboratory’s own training	Yes -laboratory’s own training	Yes -laboratory’s own training at rotation -manufacturer-provided education	Yes -laboratory’s own training at rotation -manufacturer-provided education	Yes -attend academic conferences -manufacturer-provided education	Yes -laboratory’s own training at rotation
3. Reagents
Reagent antibody	BD Multitest 6-color TBNK kit:CD45, CD3, CD4, CD8, CD19, CD16+56	BD Multitest 6-color TBNK kit:CD45, CD3, CD4, CD8, CD19, CD16+56	BD Multitest 6-color TBNK kit:CD45, CD3, CD4, CD8, CD19, CD16+56	BD Multitest 6-color TBNK kit: CD45, CD3, CD4, CD8, CD19, CD16+56	BC CYTO-STAT tetraCHROM kit: -CD45, CD3, CD4, CD8 -CD45, CD3, CD19, CD56	BC CYTO-STAT tetraCHROM kit: -CD45, CD3, CD4, CD8 -CD45, CD3, CD19, CD56	BC CYTO-STAT tetraCHROM kit: -CD45, CD3, CD4, CD8 -CD45, CD3, CD19, CD56	BC CYTO-STAT tetraCHROM kit: -CD45, CD3, CD4, CD8 -CD45, CD3, CD19, CD56	BC CYTO-STAT tetraCHROM kit: -CD45, CD3, CD4, CD8 -CD45, CD3, CD19, CD56
Additional antibody	None	None	None	None	CD16+56‡ or CD20§	CD16‡	CD16‡	CD16‡	None
Fluorophores	6 colors APC, APC-Cy7, FITC, PE, PE-Cy7, PerCP-Cy5.5	6 colors APC, APC-Cy7, FITC, PE, PE-Cy7, PerCP-Cy5.5	6 colors APC, APC-Cy7, FITC, PE, PE-Cy7, PerCP-Cy5.5	6 colors APC, APC-Cy7, FITC, PE, PE-Cy7, PerCP-Cy5.5	4 colors ECD, FITC, PC5, RD1∥, PE (CD16+56, CD20)	4 colors ECD, FITC, PC5, RD1∥, PE (CD16)	4 colors ECD, FITC, PC5, RD1∥, PE (CD16)	4 colors ECD, FITC, PC5, RD1∥, PE (CD16)	4 colors ECD, FITC, PC5, RD1∥
4. Internal QC
Material	-BC IMMUNO-TROL Cells -BC IMMUNO-TROL Low Cells	BD Multi-Check Control	-BD Multi-Check Control -BD Multi-Check CD4 Low Control	BD Multi-Check CD4 Low Control	-BC IMMUNO-TROL Cells -BC IMMUNO-TROL Low Cells	-BC IMMUNO-TROL Cells -BC IMMUNO-TROL Low Cells	-BC IMMUNO-TROL Cells -BC IMMUNO-TROL Low Cells	-BC IMMUNO-TROL Cells -BC IMMUNO-TROL Low Cells	-BC IMMUNO-TROL Cells -BC IMMUNO-TROL Low Cells
Interval	Daily	Monthly	Daily	Daily	Daily	Daily	Weekly	Daily	Daily¶
Acceptance criteria	Within the laboratory-defined target range (mean±2SD)	Within the laboratory-defined target range (mean±2SD) Cumulative CV <15%	Within the laboratory-defined target range (mean±2SD)	Within the laboratory-defined target range (mean±2SD)	Within the laboratory-defined target range (mean±2SD)**	Within the laboratory-defined target range (mean±2SD)	Within the laboratory-defined target range (mean±2SD)	Within the laboratory-defined target range (mean±2SD)	Within the laboratory-defined target range (mean±2SD)
5. EQA	KEQAS, CAP	KEQAS	KEQAS, CAP	KEQAS, CAP	KEQAS, CAP	KEQAS	KEQAS	KEQAS	KEQAS
6. Examination phase
1) Sample
Anticoagulant	EDTA	EDTA	EDTA	EDTA	EDTA	EDTA	EDTA	EDTA	EDTA
Acceptance criteria	-No clots -Within 24 hrs after sample collection	-No clots -Cell viability ≥70%	Cell viability ≥80%	-No clots -Within 24 hrs after sample collection	-No clots -Within 24 hrs after sample collection	-No clots and hemolysis -Within 24 hrs after sample collection	-Within 24 hrs after sample collection -WBC>1×103/µL,Lymphocyte>10%	- Within 24 hrs after sample collection	-No clots and hemolysis -Within 24 hrs after sample collection -WBC>1×103/µL,Lymphocyte>10%
2) Sample processing and acquisition
Cell counts	No	Yes	Yes	Yes	No	Yes	Yes	Yes	Yes
Viability check	No	Yes(samples over 48 hrs after collection)	Yes(samples over 24 hrs after collection)	No	No	No	No	No	No
Sample amount/tube	50 µL	50 µL	80 µL	1×106 cells	100 µL	100 µL	50 µL	50 µL	100 µL
Amount of reagent antibody/tube	20 µL	20 µL	10 µL	10 µL	10 µL	10 µL	5 µL	5 µL	5 µL
Lyse
Incubation time	10 mins	15 mins	10 mins	10 mins	Automated	10 mins	10 mins	10 mins	10 min
Lysing solution	BD FACS Lysing Solution	BD FACS Lysing Solution	NH4Cl Lysing solution (in-house)	BD FACS Lysing solution	BC IMMUNOP-REP Reagent Kit	BC OptiLyse C Lysis Solution	BC VersaLyseLysing Solution	BC VersaLyseLysing solution(750 µL)	BC VersaLyseLysing solution(1 mL)
Wash after lyse	No	No	No	No	No	No	No	No	No
Acquisition cell count	2×104 cells	2×104 cells in lymphogate	2×104–5×104 cells (depending on lympho-cyte count)	1×104 cells	-T cells: 6×103 -B or NK cells:1×104	5×103 cells	-T cells: 5×103 -B or NK cells:1×104	-T cells: 6×103 -B or NK cells:1×104	5×103 cells in lymphogate
7. Post-examination phase
Gating	Manual	Automated	Manual	Manual	Automated	Automated	Automated	Automated	Automated
Reporting contents	- Subset percentage	-Subset percentage -General opinion	-Subset percentage and count†† -General opinion	-Subset percentage and count†† -WBC counta -Total lympho-cyte percentage and count††	-Subset percentage -CD4 and CD8 counts††	- Subset percentage	-Subset percentage -General opinion	-Subset percentage and count†† -General opinion	-Subset percentage -General opinion

*Model name (manufacturer) of flow cytometers.

^†Operator.

^‡CD16 or CD16+56 was added for accurately assessing NK cells (e.g., CD16⁺CD56^± NK cells).

^§CD20 was added for quantifying the B-cell subset in patients on anti-CD20 monoclonal antibody therapy.

^∥RD1 has a similar emission wavelength to PE.

^¶Each internal QC material is used alternatively for daily internal QC.

**Mean value of 10 or more measurements and SD over 6 months.

^††Results are presented as counts per microliter (/µL).

Abbreviations: FCLSA, flow cytometric lymphocyte subset analysis; BD, Becton Dickinson Biosciences; BC, Beckman Coulter Diagnostics; FACS, Fluorescence-activated cell sorting; PMT, photomultiplier tube; CD, cluster of differentiation; APC, allophycocyanin; APC-Cy7, allophycocyanin-cyanine 7; FITC, fluorescein isothiocyanate; PE, phycoerythrin; RD1, rhodamine; PE-Cy7, phycoerythrin-cyanine 7; PerCP-Cy5.5, peridinin-chlorophyll protein-cyanine 5.5; ECD, phycoerythrin-Texas Red-X; PC5, phycoerythrin-cyanine 5; EQA, external quality assessment; KEQAS, Korean Association of External Quality Assessment Service; CAP, College of American Pathologists; EDTA, ethylene-diamine-tetraacetic acid; WBC, white blood cell; NK, natural killer.

All hospitals used commercial reagent antibody (RA) kits: four used the BD Multitest 6-color TBNK (BD6) kit (BD, San Jose, CA, USA), and five used the BC CYTO-STAT tetraCHROME (BC4) kit (BC, Miami, FL, USA). The BD6 kit includes CD45, CD3, CD4, CD8, CD19, and CD16+56 antibodies (Abs) in a single tube with six fluorophores. The BC4 kit consists of one tube containing CD45, CD3, CD4, and CD8 Abs for T cells, and the other containing CD45, CD3, CD19, and CD56 Abs, with four fluorophores in each tube. CD16 is not included in the BC4 kit; therefore, three hospitals added CD16, and one added CD20 or CD16+56 optionally. Six hospitals used two levels of commercial QC materials simultaneously for internal QC (IQC), one used two levels alternatively, and two used only one level. Seven hospitals performed IQC daily, and two performed IQC weekly or monthly. All hospitals used laboratory-defined target ranges (mean±2SDs), and one additionally used the cumulative CV (<15%).

All hospitals participated in the Korean Association of External Quality Assessment Service (KEQAS) program, and four hospitals additionally participated in the College of American Pathologists (CAP) external quality assurance (EQA) program. The sample amount (SA) and RA amount (RAA) per tube varied even among hospitals using the same kit. For SA, four hospitals used 50 µL, three used 100 µL, one used 80 µL, and one adjusted the SA to 1×10⁶ cells. The RAA ranged from 5–20 µL. All hospitals used the lyse-no-wash method [5]. The acquired cell count (ACC) ranged from 5×10³ to 5×10⁴ cells. Three hospitals acquired 5×10³ to 2×10⁴ cells, two acquired 5×10³ or 2×10⁴ cells in the lymphogate, one acquired 2×10⁴ to 5×10⁴ cells depending on lymphocyte count, and three used different ACCs depending on cell type: 6×10³ cells for T cells and 1×10⁴ cells for B/natural killer (NK) cells in two hospitals, and 5×10³ cells for T cells and 1×10⁴ cells for B/NK cells in one hospital. Six hospitals used automated gating, and three used manual gating. All hospitals reported percentages. Some hospitals additionally reported general opinions, cell counts (/µL), white blood cell (WBC) and lymphocyte counts (/µL), lymphocyte percentage, or CD4/CD8 ratio.

Given the high maintenance costs, daily instrument setup is difficult, and in hospitals conducting fewer FCLSA tests, instrument setup according to the test schedule is required.

The CLSI H42-A2 guidelines recommend an Ab panel consisting of CD3, CD4, CD8, CD19, CD45, and CD56 Abs to detect major lymphocyte subsets (T, B, and NK cells); CD3, CD4, and CD8 Abs for T-cell subsets; CD19 Ab for B cells; and CD56 Ab for NK cells [5]. Since anti-CD20 monoclonal Ab (mAb) therapy can induce CD20 blockade in FCLSA, CD19 Ab is additionally required for monitoring B cells after anti-CD20 mAb therapy [6, 7]. The simultaneous use of single-color CD16 and CD56 Abs can increase the fluorescence intensity of CD16⁺CD56^± NK cells [5]. Human T, B, and NK cells exist in various subtypes with different functions, and substantially more Abs are needed to identify these subtypes in FCLSA [8]. Recently, FCM has been advanced for precise cell subpopulation analysis, and a 40-color FCM panel has been introduced for deep immunophenotyping of major lymphocyte subsets [9]. With the development of numerous immunomodulatory therapies, prospective multicenter studies have advanced standardized protocols and reagent development [1, 3, 7–9]. The number of panel Abs for FCLSA in Korea should be increased.

As dead cells result in nonspecific binding and autofluorescence, viability testing is recommended for samples used >24 hrs after collection [5, 10]. Studies have recommended 70% or 75% as a minimum threshold of cell viability [5, 11, 12]. In our study, all hospitals used EDTA as the anticoagulant, and two hospitals assessed cell viability before FCLSA. Most hospitals met the acceptance criteria, with samples analyzed within 24 hrs of collection. Two hospitals checked cell viability using a threshold of >70% or 80%. For accurate FCLSA, sufficient lymphocytes and no marked lymphocytosis are prerequisites [5]. When the WBC count is <20×10⁹/L, RAs can saturate target antigens on 1×10⁶ cells in 100 µL samples [5, 6]. BD and BC recommend using 100 µL of whole blood; for RAA per tube, BD and BC recommend 20 µL and 10 µL, respectively [13, 14]. BC recommends WBC counts of 3–10×10⁹ cells/L for optimal staining [14]. The SA and RAA differed among hospitals, even when using the same reagents. These variations likely affect test result consistency. At least 5,000 lymphocytes should be acquired to accurately analyze small lymphocyte subsets, which comprise approximately 10% of total lymphocytes [5, 6].

Currently, over 50 domestic hospitals participate in the FCLSA KEQAS program. Unlike other FCM tests, for FCLSA, most hospitals use commercial kits, which are expected to ensure a certain level of result consistency. However, cumulative EQA data from KEQAS revealed variability in FCLSA results [15]. KEQAS data from 2018 to 2023 showed that the CV was highest for NK cells, followed by B cells, and lowest for T cells (Supplemental Data Fig. S1). Guidelines recommend reporting numeric results of major lymphocyte and T-cell subsets [5, 6]. Some NK cells express CD8, and NKT cells co-express CD3 and CD56 [5, 16]. Therefore, NK-cell subsets should be excluded to identify T-cell subsets (e.g., CD3⁺CD56^– [or CD16⁺56^–] for total T cells and CD3⁺CD8⁺CD56^– [CD16⁺56^–] for cytotoxic T cells) [16]. For NK cells, the marker composition differs between the two commercial kits; BD uses a CD16+56 combo Ab, whereas BC uses CD56 alone, which may contribute to result variability. Furthermore, CD8 often appears in regions of positive fluorescence intensity that can be separated into low- and high-intensity areas [5]. CD3⁺CD8⁺ suppressor/cytotoxic T cells fall into the high-intensity area [5]. NK cells also express CD8, but at low intensity [5]. This should be considered during analysis. Differences in ACC across hospitals may also contribute to variability in KEQAS results, as shown in this survey. This should be considered when analyzing B and NK cells, which have lower cell counts than T cells.

Survey participants provided suggestions for KEQAS operation and result reporting. EQA results are often evaluated based on consensus values, using the distribution values of most participating institutions as a reference. Standardizing the result reporting format, gating, and principles for reporting counts on a dual platform would be beneficial. Recommended reference values would also be helpful. Other suggestions included the need for standardized interpretation, which would require sharing anonymized data and establishing a joint interpretation system.

Our results indicate that standardization of sample processing and the acquisition process is the most urgent priority for FCLSA in Korea. We recommend establishing guidelines based on observed differences in SA, RAA per test tube, and variation in ACC.

FCLSA is a representative FCM test performed using commercial RA kits. Although the nine surveyed hospitals are leading institutions with high expertise in FCM, FCLSA procedures varied across hospitals. Inclusion of more institutions in the survey may have revealed greater heterogeneity. As only nine university hospitals that frequently perform FCM testing were included, the findings may not fully reflect current laboratory practices across all hospitals implementing FCLSA.

To provide fundamental data for initiating FCM standardization in Korea, we recently reported the current status of flow cytometric immunophenotyping of hematolymphoid neoplasms in Korea [17]. FCLSA standardization is also considered necessary. We have provided a laboratory practice guide for FCLSA and highlighted differences in FCR types and RA kits. FCLSA increasingly requires deep immunophenotyping, and its clinical application requires policy support from the National Health Insurance and the Ministry of Food and Drug Safety.

SUPPLEMENTARY MATERIALS

Supplementary materials can be found via https://doi.org/10.3343/alm.2025.0064.