Cytometry Part B: Clinical Cytometry最新文献

CD38 and CD138 are important diagnostic markers in flow cytometric analysis of plasma cells (PC) in the context of multiple myeloma (MM). Anti-CD38 therapy, such as daratumumab, exacerbates CD38 detection. In addition, CD138 can be degraded and is then no longer easily detectable on the cell surface. Variable heavy domain heavy chain antibodies (VHH) are single variable domain antibody fragments. Clone JK36 consists of two anti-CD38 VHH fragments and allows targeting of a cryptic CD38 epitope that is not accessible to conventional antibodies (CA). Therefore, our aim was to test VHH in comparison to our conventional anti-CD38 antibody (LS198) in MM bone marrow samples after daratumumab therapy (d-t) compared to therapy-naïve (n) and samples with unknown therapy. A total of 111 samples were analyzed (n = 11 n, n = 81 d-t, n = 18 with unknown therapy). While CD38 was equally well detected by VHH and CA in therapy-naïve samples, CD38 could only be detected in 8% of d-t samples with CA but in 91% with VHH. This resulted in an overall significant reduction in the number of detectable PC, and three samples with undetectable PC by CA compared to VHH. Furthermore, CD138 was reduced/degraded in 52% of d-t samples of which 88% had undetectable CD38 by CA. In addition to proper detection of CD38, VHH is also able to determine a potential CD38 reduction of cell surface expression, as shown by a reduction in CD38 median fluorescence intensity (MFI) on d-t compared to n samples. One d-t sample revealed two distinct PC populations differing by dim and bright CD38 expression, only detectable by VHH. Interestingly, samples with unknown treatment history can be grouped into scenarios most likely treated with daratumumab, or rather treatment-naïve, respectively. In summary, VHH provides superior CD38 detection in d-t MM patients, which is vital for diagnostic samples, and it is capable of providing information about CD38 integrity on the cell surface.

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.

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.