Cytometry Part B: Clinical Cytometry最新文献

Platelet refractoriness is caused by antibodies against human leukocyte antigens or human platelet antigens. However, readily applicable assays that assist in selecting immunologically compatible platelet units remain limited. We established a flow cytometric platelet crossmatching assay and assessed its clinical utility by interpreting the results in conjunction with post-transfusion corrected count increments (CCI). Platelets were incubated with serum that may contain anti-platelet antibodies. Using flow cytometry, CD41-expressing platelets were gated, and the median fluorescence intensity (MFI) of fluorescein isothiocyanate (FITC)-conjugated anti-human IgG was measured. The MFI ratio was calculated as test sample MFI divided by baseline negative control MFI. The cutoff value and limit of detection (LoD) were estimated. Platelet crossmatching was performed using residual segments of transfused platelet units and patient serum, and the results were retrospectively interpreted in conjunction with CCIs and clinical findings. The MFI ratios were clearly distinguishable among the three groups: high-positive controls, low-positive controls, and known negative samples (p < 0.001). The cutoff value was calculated to be 1.35, and the LoD was 1.53. In total, eight platelet transfusion events in five patients were analyzed. Four cases were interpreted as non-immune refractoriness, and the remaining four showed adequate post-transfusion platelet increments. All corresponding platelet crossmatching results were negative, which was considered appropriate given that the assay is designed to reflect immune refractoriness. The flow cytometric platelet crossmatching assay was established and demonstrated to be applicable. The assay can help predict transfusion outcomes in alloimmunized patients and contribute to the selection of compatible blood units.

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.

The International Harmonisation Protocol (IHP) for Reticulocyte Percentage (%Retic) in CLSI-H44-A2 is a microscopic procedure performed on a new-methylene-blue-stained blood film (NMB-IHP). However, IHP-compliant Measurement Procedures (MPs) based on flow cytometry (FCM) are recommended for practical and accurately assigned reference values. Thiazole orange (TO) for nucleic acid staining and CD235a immunostaining for erythrocyte gating are currently widespread for MP using FCM (TO/CD235a-MP); however they have not been validated using NMB-IHP. We recently developed MP using FCM with anti-CD45/CD41a/CD61 antibodies, that are utilized for excluding platelets from the un-nucleated blood cell fraction to select erythrocytes (TO/CD41a/CD61-MP). The aim of this study is to validate TO/CD41a/CD61-MP for measurement of reticulocytes as an IHP-compliant MP. First, TO/CD235a-MP was validated as an IHP-compliant MP by comparing it with NMB-IHP. Then TO/CD41a/CD61-MP was compared with TO/CD235a-MP. The practical utility of TO/CD41a/CD61-MP was evaluated using XN-2000™(Sysmex) and Celltac G + ™(Nihon Kohden) hematology analyzers to assess the accuracy of different nucleic acid staining methods. We first confirmed that TO/CD235a-MP was consistent with NMB-IHP, then evaluated TO/CD41a/CD61-MP using TO/CD235a-MP. Regression analysis demonstrated consistency between TO/CD41a/CD61-MP and TO/CD235a-MP, thereby establishing TO/CD41a/CD61-MP as IHP-compliant MP. In the accuracy assessment, the correlation coefficients to TO/CD41a/CD61-MP were observed to be 0.97 for both hematology analyzers with %Retic ranging from 0.0 to 8.2. TO/CD41a/CD61-MP serves effectively as an IHP-compliant MP. This study provides the first empirical validation of FCM-based MPs.

Measurable residual disease (MRD) testing for chronic lymphocytic leukemia (CLL) is often done on peripheral blood (PB) since the concordance of results with bone marrow is high and testing is less invasive. When analyzing CLL MRD data, one must be aware of small, normal populations in the PB that may be mistaken for residual CLL cells. As part of our CLL MRD assay validation, PB samples were collected from 10 healthy donors and a 2-tube CLL MRD flow cytometry panel was stained for each donor using markers CD19, CD20, BAFF-R, kappa, lambda, CD5, CD200, CD23, CD38, CD81, ROR1, CD79b, CD43, and CD45. Additional markers were utilized to exclude T-cells, NK-cells, and myeloid cells from the analysis. Samples were acquired on the Navios EX flow cytometer, and the data were analyzed using FCS Express software. Once the test was implemented, CLL PB patient samples were monitored. All 10 PBs from healthy donors contained small populations of cells present in the lymphocyte gate which mimicked CLL cells in their expression of CD45, CD19, CD20, CD43 (both positive, although CLL cells showed dimmer positive expression), CD79b, and level of surface light chains, immunophenotypically compatible with plasmablasts. Clinical implementation of the CLL assay revealed 12 out of 77 (16%) CLL PB patient samples demonstrating a small population of plasmablasts. Plasmablasts normally exist in peripheral blood at levels detectable by flow cytometry MRD assays and may be a potential confounder in the identification of MRD in CLL.

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 (Assays_L&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 (Staff_Signout) or other laboratory operations (Staff_LabOps). 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 (Assays_Total) 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 Assays_L&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 Staff_LabOps. 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 Assays_Total per Staff_LabOps 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.