Reproducibility in clinical flow cytometry is essential for diagnosis, longitudinal monitoring, and interlaboratory harmonization, particularly when threshold-based lymphocyte measurements such as absolute CD4 counts guide clinical management. Although standardization of instruments, reagents, antibody panels, and acquisition protocols has reduced variability at the level of signal generation, variability introduced by human interpretation during manual gating has not been quantitatively separated from biological and technical sources. Here, we applied a two-stage workflow audit anchored to a fixed, invariant automated analysis used as an analytical reference to quantify interpretive variability in routine clinical TBNK (T-cell, B-cell, and natural killer cell) flow cytometry and to assess its impact on CD4 decision-band classification. Standardized quality-control materials and 320 consecutive clinical samples, independently analyzed by six technologists on three harmonized cytometers, were used to define an empirical analytical performance envelope (i.e., the expected range of performance) and to evaluate departures from this envelope under routine conditions. Under quality-control conditions, automated-manual differences were stable and interpretable: percentage-based endpoints showed near-zero bias with narrow dispersion, absolute counts exhibited small, consistent offsets, and CD4 decision-band assignment was preserved. In contrast, routine clinical samples showed substantially greater dispersion that was structured primarily by operator identity rather than instrument configuration. Operator-associated disagreement exceeded instrument-associated variability, was low dimensional and reproducible, and was concentrated near established CD4 thresholds, yielding discordant but adjacent decision-band classifications. These findings quantify a previously unmeasured source of analytical variability in routine hematology testing that can affect threshold-based clinical classification despite acceptable quality-control performance.
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Systemic mastocytosis (SM) is a clonal mast cell (MC) disorder characterized by aberrant immunophenotypes, including expression of CD25, CD2, and occasionally CD30. CD123, the α-subunit of the interleukin-3 receptor, is a therapeutic target in hematologic malignancies and has been reported to be expressed on neoplastic MCs by immunohistochemistry (IHC) with prognostic implications. This study aims to characterize CD123 expression in SM by flow cytometry. We retrospectively analyzed 142 bone marrow samples from 79 SM patients (81 diagnostic samples) and 25 controls with normal MC immunophenotype. Flow cytometry was performed using a clinically validated 9-color mast cell tube which included CD123. Data collected included SM subtype, clinical and laboratory features, MC burden, and marker expression. Statistical analyses were performed in R. CD123 was expressed on MCs in 91% of SM cases (ISM 92%, SM-AHN 94%, SSM 100%, ASM 100%, MCL 50%). Median percentage of MCs positive for CD123 was 53.9% (IQR 8.1-83.4). Compared to prior IHC data (overall 64% positivity), flow cytometry demonstrated more cases with CD123 expression by MCs. No significant correlations were observed between CD123 expression and serum tryptase, KIT D816V allele burden, or MC burden. CD123 is frequently expressed on neoplastic MCs in SM by flow cytometry, across all subtypes. These findings support further investigation of CD123 as a therapeutic target and warrant correlation with IHC and clinical outcomes in larger cohorts.
Clinical flow cytometry laboratories are facing rising test volumes, greater assay complexity, and increasing requirements for quality control and assay validation. In response, the International Clinical Cytometry Society (ICCS) conducted a workload survey in early 2023 to gather updated information on assay volumes, complexity, staffing, and technology. Data analysis focused on identifying correlations between length of time to introduce new assays and other factors as a means to gain insight about laboratories that seem to be either adapting or struggling. Flow cytometry assays were categorized into 3 levels of technical/interpretative complexity: high (e.g., measurable/minimal residual disease (MRD assays)), moderate (e.g., leukemia/lymphoma assays (AssaysL&L), excluding MRD assays), and low (e.g., CD4 count). Annual assays per staff member were calculated according to staff involved in case sign-out (StaffSignout) or other laboratory operations (StaffLabOps). Respondents were from 101 laboratories in the United States (69.3%), Canada (4.0%), and other countries (26.7%). Low, moderate, and high technical/interpretative complexity assays were performed in 85.1%, 97.0%, and 47.5% of all laboratories, respectively. Median annual total assays (AssaysTotal) per laboratory were 3515 and, based on complexity, were 1518.5 (low), 1808.8 (moderate), and 350 (high). Among all laboratories, the median time (interquartile range) to introduce new AssaysL&L was 6 mos. (4-12 mos.), to introduce MRD assays was 11 mos. (5-12 mos.), and to validate/go-live with new cytometers was 8 mos. (4-12 mos.); these times positively correlated with each other. This study confirmed significantly increased workload since the prior ICCS 2013 workload survey with a concurrent decrease in StaffLabOps. Faster introduction of new assays correlated with other successes, including quicker validation of and going live with new cytometers. Among all laboratories, those that performed myeloid MRD assays versus those that did not were also found to be faster to introduce new assays. The need for sufficient staffing has been emphasized because laboratories with both higher annual volumes of myeloma MRD assays and higher ratios of AssaysTotal per StaffLabOps were slower to introduce new assays. "Lack of staff and/or time dedicated or protected for assay development" and, more generally, "staff number" were the most commonly identified major barriers for new assay development, with the former specifically linked to slower introduction of new assays among all laboratories.
Multiparametric flow cytometry is a highly valuable method for the assessment of measurable residual disease (MRD) in multiple myeloma patients. The aim of the study was to evaluate a one-tube MRD panel on a DxFlex flow cytometer including all EuroFlow recommended immunophenotypic markers (i.e., cytoplasmic light chain kappa and lambda, CD19, CD27, CD38, CD45, CD56, CD81, CD117, CD138) extended by CD200 to a total of 11 fluorochromes in one tube. Bone marrow aspirates from clinical routine underwent an ammonium chloride-based bulk lysis, followed by staining the surface antibodies and, after permeabilization, kappa, and lambda light-chain intracellular staining. We acquired 1 × 107 cells per sample with a DxFlex flow cytometer. We determined the limit of detection (LOD) and lower limit of quantification (LLOQ) as recommended by the International Myeloma Working Group. For the clinical evaluation, 68 samples from 53 patients with multiple myeloma under or after treatment were analyzed with the one-tube MRD panel, and the results were compared with our routine plasma cell panel (RPCP). Six of the 68 samples were additionally sent to another laboratory to confirm our results with the contemporary EuroFlow next-generation flow (NGF) assay. For our novel one-tube MRD panel we determined a LOD of 0.00016% (1.6 × 10-6) and a LLOQ of 0.00059% (5.9 × 10-6). Out of 68 specimens, 55 (80.9%) showed concordant results between the MRD- and the RPC-panel. Thirteen (19.1%) specimens showed a distinct population of abnormal plasma cells with the MRD panel not detectable with the RPC panel. The six samples simultaneously measured with our novel one-tube MRD panel and the EuroFlow NGF assay showed concordant results. The novel one-tube MRD panel meets the quality specifications of the International Myeloma Working Group. Our clinical evaluation found higher sensitivity for the one-tube MRD panel when compared to our RPC panel and concordant results with the contemporary EuroFlow NGF assay. Therefore, our novel one-tube MRD panel is well-suited for detecting MRD in multiple myeloma patients.
Quantifying malarial parasite density is crucial for diagnosis and treatment in endemic areas. While Malaria-derived particles (MDPs) have been linked to malaria pathology, a direct quantification method for routine laboratory use remains unestablished. To address this, our study optimized a flow cytometry approach to enumerate MDPs per microliter of blood. Specimens were incubated with propidium iodide and red blood cell (RBC) lysis solution. The number of MDPs was quantified using a CytoFlex flow cytometer, size-standard beads, and counting beads. Electron microscopy was used to study the ultrastructures of the malarial parasites in the lysed RBC specimens. A significant increase in MDP levels was detected in blood samples from P. falciparum and P. vivax infections, but fewer than 1 particle/μL of MDPs were detected in the controls. The number of MDPs correlated with the percentage of infected red blood cells (iRBCs) obtained by manual counting (R2 = 0.94). The dilution assay demonstrated a strong correlation between the measured and expected values of the MDPs. An electron microscopic study demonstrated that different stages of malarial parasites exist in lysed RBCs in the form of membrane-bound spherical cells. A positive association was established between parasite density and MDPs across both P. falciparum (R2 = 0.94) and P. vivax (R2 = 0.91) infections. We demonstrated the potential use of flow cytometry for determining the MDP concentration. The developed approach is reliable and straightforward for the diagnosis and treatment of patients with malarial parasite infection in routine laboratory settings.
Antibody titration is essential for optimizing platelet flow cytometry, a technique widely used to evaluate platelet phenotype, activation status, and function. This manuscript outlines practical approaches for platelet antibody titration in whole blood, with tailored strategies for constitutive and inducible markers. It emphasizes the use of appropriate controls, consideration of marker coexpression, and selection of subsaturating antibody concentrations to maximize signal resolution while minimizing background and artifactual activation. Quantitative metrics such as the stain index and separation index are introduced as tools for evaluating staining performance. The discussion also addresses key technical variables, including combinatorial titration, spillover spreading, lot variability, and antibody-induced activation. Titration under final assay conditions is recommended to ensure reproducibility and biological relevance. These strategies provide a foundation for developing robust, high-resolution platelet assays that support both research and clinical applications, particularly as flow cytometry evolves toward greater automation and standardization.
Accurate quantification of chimeric antigen receptor (CAR) T cells is essential for monitoring post-infusion CART expansion and persistence and for real-time clinical decision-making. Multiparameter flow cytometry (MFC) enables rapid, live-cell detection with absolute quantification and concurrent immunophenotypic characterization. This review focuses on the practical and technical aspects of flow cytometry-based CAR T-cell monitoring, including selection of CAR detection reagents (target-specific, construct-specific, and target-agnostic strategies), assay optimization, purpose-driven panel design, and matrix-appropriate validation for peripheral blood and other clinically relevant specimens. We also address assay considerations unique to gene-edited allogeneic CAR T-cell products, including the use of surrogate immunophenotypic approaches when construct-specific reagents are unavailable. Finally, we discuss the role of MFC in identifying CAR T-cell clonal expansions and in evaluating suspected secondary hematolymphoid neoplasms in the post-CAR T setting.
The objective of this 12-color/13-antibody single-tube panel is to assist in the diagnosis of T/NK-cell leukemias and lymphomas. In the clinical setting, the absence of a standardized T/NK-cell panel limits inter-laboratory data comparability, contributes to diagnostic variability, and results in redundant efforts across laboratories to design and validate panels for flow cytometry. Developing and promoting a panel that allows for rapid and fairly comprehensive labeling of surface T/NK-cell markers, including the anti-T-cell receptor β-chain constant region 1 (TRBC1) antibody, will streamline the workflow by establishing a standard immunophenotyping T/NK-cell panel for detecting neoplastic T/NK cells and lay the foundation for future COMIPs.
X-linked agammaglobulinemia (XLA) is most often caused by impaired Bruton's tyrosine kinase (BTK) function. Rapid laboratory support is needed when genetic findings are inconclusive or pending. We developed a flow-cytometric assay to quantify intracellular BTK protein in monocytes, aiding the diagnosis of XLA. A stain index (SI) reference range was established using healthy donors (n = 25; SI median 1.14, range 1.01-1.66). ROC analysis defined a diagnostic cutoff of SI = 1.79, providing 95% sensitivity and 100% specificity compared to patients with pathogenic BTK variants (n = 40). Using this cutoff, abnormal BTK expression was detected in 38/40 genetically confirmed XLA patients. All patients with variants of uncertain/likely pathogenic significance (n = 17) had SI values above the cutoff. Bone marrow analysis in a patient subset showed a developmental arrest at the Pre-BI stage. The assay offers a rapid, clinically applicable readout to support XLA diagnosis and carrier evaluation in females, effectively complementing genetic testing.
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The diagnosis of T-cell neoplasms remains one of the most challenging areas in hematopathology due to the immunophenotypic heterogeneity and subtle aberrancies often present in these entities. This "Best Practice" manuscript provides a practical framework for laboratories to design, validate, and interpret immunophenotyping studies of immature and mature T-cell neoplasms. We outline the utility of key antigens in the screening and classification of T-cell lymphomas/leukemia including TRBC1 and TRBC2. Analytical strategies using the "difference from normal" method and template-based gating are discussed, along with validation considerations aligned with CLSI H62 guidelines. By integrating these principles into laboratory workflows, this manuscript aims to standardize and improve the assessment of T-cell neoplasms across diverse clinical settings.
CD300e is a marker of mature monocytes in flow cytometry; however, there is limited detailed information on staining patterns in conjunction with other monocyte markers. We evaluated the flow cytometric staining patterns of CD64, CD14, and CD300e in 12 negative and 33 positive peripheral blood specimens and 16 negative and 56 positive bone marrow specimens. The positive cases were involved by myeloid neoplasms (increased blasts and/or abnormal monocytes). Flow cytometry plots were reviewed for each case, the monocyte population was identified by bright CD64 expression, and the monocyte maturation pattern was visualized by CD14 versus CD300e plots. Peripheral blood and bone marrow differential counts were collected. A total of 39% (22/56) of the positive bone marrow cases showed a different maturation pattern from the negative bone marrow cases. Of the positive peripheral blood cases, 28/33 (85%) showed a CD14 by CD300e pattern different from that observed in the negative peripheral bloods. When the subset of bone marrow cases involved by monocytic neoplasms was evaluated, there was no significant difference between monocyte percentage by flow cytometry versus morphology and between blast plus promonocyte percentage by flow cytometry versus morphology. We conclude that isolation of monocytes by bright CD64 expression and low side-scatter and subsequent evaluation of the CD14/CD300e maturation pattern may help identify myeloid neoplasms. Quantification of CD64 + CD14- and/or CD64 + CD300e- cells by flow cytometry may aid blast/blast equivalent identification/quantification.
Immunophenotyping by flow cytometry is a valuable test providing important information in a timely manner. In clinical laboratories, it is performed using validated antibody panels designed to ensure consistent and accurate results. However, unforeseen situations, such as unique or unusual immunophenotypes, or supply chain issues, may necessitate ad hoc modifications to these panels. This manuscript provides guidance for performing minor modifications, such as substituting or adding one or two antibodies, while maintaining the integrity of the assay. These modifications are intended for rare clinical situations and are not substitutes for the full validation protocols outlined in CLSI H62. An example of this would be a patient with a rare, but not uncommon, situation in which a B cell lymphoma lacks expression of CD19, CD20, and surface light chains, such that the lineage of the neoplastic cells cannot be determined without a straightforward addition or substitution of another marker into a laboratory's available panel. The recommendations and best practices herein aim to optimize patient care by allowing laboratories to adapt to unique clinical scenarios without compromising assay performance and are not a way to permanently modify the assay. Key considerations include assessing the impact on fluorescence compensation, antibody binding, assay sensitivity, and overall assay performance. The manuscript provides limitations for the extent of modifications, examples, and troubleshooting strategies to ensure reliable results when ad hoc changes are made. Proper documentation with review and approval by laboratory medical directors is recommended to mitigate risks associated with these modifications.
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Circulating monocyte partitioning refers to the relative quantification of the three main monocyte subsets in the peripheral blood, namely classical (cMo), intermediate (iMo), and non-classical (ncMo) monocytes, as assessed by flow cytometry, a new nomenclature described 15 years ago. This distribution is influenced by physiological variation as well as by certain therapeutic interventions. In addition, pathological alterations in monocyte partitioning are now well characterized in chronic hematological neoplasms. Most notably, a relative accumulation of cMo exceeding 94% of total circulating monocytes was identified more than a decade ago as a phenotypic hallmark of chronic myelomonocytic leukemia (CMML) and has since been incorporated into the most recent revision of the WHO classification. Altered monocyte partitioning has also been reported in patients with myelodysplastic syndromes (MDS) and myeloproliferative neoplasms (MPNs), highlighting its broader relevance across myeloid disorders.
Measurable residual disease (MRD) monitoring by multiparameter flow cytometry (MFC) is well established in bone marrow (BM) samples for acute myeloid leukemia (AML), but its use in peripheral blood (PB) remains less developed. We adapted a semi-automated and well validated MFC-MRD assay, originally developed for BM, to PB aiming to evaluate its applicability and concordance with established methods. To transfer the MFC-MRD assay from BM to PB, we incorporated population-specific reference values derived from healthy and chemotherapy-exposed non-AML controls. MRD detection was based on a combined aberrant leukemia associated immunophenotype (aLAIP) and different from normal (DfN) approach. We compared MFC-MRD detection in paired PB and BM samples and evaluated concordance with molecular MRD (Mol-MRD) diagnostics. Additional analyses included assessment of mast cells as markers for BM hemodilution as well as time requirements for analyses. Reference values for aberrant populations in PB were generally lower than in BM, particularly considering immature markers. At follow-up, MFC-MRD positivity was less common in PB than in BM, leading to a concordance rate of 73.7%. Both quantitative and qualitative differences in aberrant antigen expression between PB and BM were observed. Most cases of MRD positivity were characterized by newly emerging aberrant features rather than persistent ones. Concordance between MFC-MRD from PB and Mol-MRD from BM was 64.6%, similar to that of MFC-MRD from BM versus Mol-MRD from BM (58.5%). Most discordant cases (MFC-MRDpos/Mol-MRDneg) may reflect clonal hematopoiesis or limited spectrum of molecular panels. Mast cell quantification proved useful in identifying hemodiluted BM samples, which comprised ~15% of cases. Inter-rater agreement for MRD detection was strong (Kα > 0.8). The MFC-MRD gating and evaluation was rapid (~2 min/sample in both PB and BM). The adapted MFC-MRD assay is feasible and robust in PB, offering a fast and reproducible alternative to BM-based MRD monitoring. However, the prognostic relevance of the proposed method needs to be validated in a prospective cohort of patients.
Förster resonance energy transfer (FRET) serves as the fundamental mechanism underlying a wide array of technologies employed in biomedical research and clinical diagnostic applications. However, unintended FRET is often overlooked, despite its potential to introduce assay artifacts that may lead to misinterpretation. Here, we examine the impact of unwanted FRET on clinical flow cytometric testing, focusing on the T-cell assay incorporating TRBC1 and TRBC2, which has emerged as a powerful method for assessing T-cell clonality. We illustrated the effect using a representative case of spurious antigen expression associated with false TRBC restriction and demonstrate through control experiments that this artifact arises when target antigens in close proximity are labeled with fluorochromes having overlapping spectra. We quantified the spectral overlap integral J(λ), and generated a matrix for commonly used fluorochromes, providing a practical tool to identify fluorochrome pairs with a high FRET propensity that should be avoided when labeling closely situated antigens. Overall, recognizing the potential effects of unintended FRET and implementing strategies to prevent or minimize its impact are essential for ensuring accurate interpretation of clinical flow cytometric assays.