Cytometry Part A最新文献

Spectral flow cytometry and high-parameter panels provide a powerful tool to meet the growing demands for deep immunophenotyping. This study aimed to adapt a published 40-color spectral flow cytometry panel for whole blood analysis, evaluate the impact of cryopreservation, and explore the potential benefits of overnight staining on data quality and cost-effectiveness. The optimization process revealed several artifacts, including loss of IgG signal and spectral profile changes in PerCP-eFluor710, which were addressed through protocol and reagent modifications. Cryopreservation resulted in significant alterations to granulocyte morphology and viability, limiting direct comparisons with fresh blood samples. Preservation also led to altered distributions in 30% of gated populations, particularly affecting T cell, dendritic cell, and monocyte subsets. Overnight staining with reduced antibody concentrations enhanced resolution for key markers while allowing for 5- to 20-fold reductions in antibody usage, substantially lowering panel costs. Contrary, extended incubation times also decreased the detection of markers critical for identification of NKT-like and regulatory T cell subsets, including CD2 and CD39. This underscores the importance of fluorochrome interference, preservation effects, and extended incubation times when optimizing high-dimensional panels, suggesting opportunities to improve staining protocols in clinical and epidemiological research, particularly where resolution or cost-effectiveness are primary concerns.

Over the past 6 decades, cytometry has evolved from a niche experimental method into a cornerstone of modern biomedical research. The development of the first cell sorter by Mack Fulwyler laid the groundwork for technologies that now define single-cell analysis. From the early challenges of instrument operation and laser alignment to today's era of automated, high-parameter systems, the field has undergone continual transformation. We now enter what may be termed the “diamond age” of cytometry, an era of exceptional sensitivity, resolution, and analytical depth, driven by innovations such as spectral flow cytometry. Here, we reflect on the historical milestones that shaped the discipline, the cultural and educational shifts within the community, and the future challenges of standardization and quantitative rigor.

Proliferating cells have a sustained high demand for regeneration of electron acceptors as nicotinamide dinucleotide (phosphate) (NAD(P)⁺/NAD(P)H) is involved in a number of critical redox reactions within cells. However, their analysis in living cells is still challenging. We propose that combining label-free NADH and NADPH fluorescence lifetime imaging (NAD(P)H-FLIM) and signal-enhanced nuclear magnetic resonance (NMR) spectroscopy allows new, deeper insights into changes in specific metabolic pathways in living cells. For proof of principle, NAD⁺-metabolism was perturbed by specific inhibition of the rate-limiting enzyme of the NAD⁺ “Salvage pathway” Nicotinamide phosphoribosyltransferase (NAMPT) by FK866 in RAMOS human lymphoma cells. FK866 treatment leads to NAD(H) reduction, followed by reduced RAMOS cell proliferation. The NAD(P)H-FLIM analysis revealed increased general NAD(P)H-dependent metabolic activity indicated by increased ratios of enzyme-bound to total NAD(P)H concentration upon NAMPT inhibition. More importantly, a marked reduction in lactate dehydrogenase (LDH) activity accompanied by NADPH oxidase activity increase is observed. Using signal-enhanced NMR spectroscopy a reduced flux of pyruvate to lactate catalyzed by LDH is detectable in real time in living cells. This strongly supports NAD(P)H-FLIM analysis and demonstrates that intervening in the NAD⁺ “Salvage pathway” can have specific and global consequences for cells. Our principle study shows how spatially-resolved metabolic imaging techniques, that is, NAD(P)H-FLIM, are complemented by real-time NMR, paving the way toward a comprehensive spatiotemporal understanding of metabolic pathways in living cells.

The use of flow cytometry to investigate phytoplankton functional groups is rapidly expanding worldwide, using lab- or ship-based instruments or autonomous environmental monitoring platforms. Automation, coupled with greater autonomy, allows for higher spatial and temporal resolution of phytoplankton groups, enhancing understanding of their dynamics and patterns, generating large datasets. The level of resolution is determined by both instrumental capabilities and optimization of its acquisition settings. Sharing these datasets with the scientific community, whether to improve global phytoplankton distribution resolution or facilitate the intercomparison of environmental indicators among monitoring laboratories, strongly relies on quality-controlled instruments and standardized data acquisition and analysis. This article focuses on CytoSense-type (CytoBuoy, NL) flow cytometers, which operate by recording the optical pulse shapes of particles as they pass through a laser beam. Different configurations such as laser wavelength and power, sheath fluid management, sample inlet design, and dataset output format were not considered, in order to focus on optimization and protocol standardization to resolve the whole phytoplankton size spectrum, from the smallest autofluorescing prokaryotes to colonies and chain-forming species. In this study, coincidence, PMT voltage, trigger threshold optimization, and regular quality control procedures are considered and discussed, using datasets from three types of instruments and two contrasted marine coastal waters as case studies. The primary goal of this study is to establish a framework to guide and support the exploration and application of this type of flow cytometer, ultimately achieving a reliable and optimal resolution for sample acquisition of natural waters.

Quantification of T-cells, B-cells, and NK-cells assay is crucial for diagnosing and monitoring immune diseases and evaluating lymphodepleting therapies. To standardize and validate an optimized workflow of the Beckman Coulter DuraClone IM Phenotyping Basic Kit for quantification of TBNK subsets in peripheral blood. Procedural changes included the use of an alternative lysis buffer, the addition of counting beads, the elimination of centrifugation steps, and an increase in acquisition volume. Validation followed CLSI H42-A2 and H62 guidelines, assessing accuracy, precision, linearity, and limit of quantification (LLOQ). Accuracy was evaluated by comparison with Immuno-Trol controls and by Bland–Altman analysis against standard immunophenotyping methods. External proficiency was assessed through participation in the College of American Pathologists (CAP) TBNK program in 2024. Procedural steps were reduced by 50%, and processing time by 38.6%. The modified method demonstrated high accuracy (−3 < bias < 35; cells/μL), a low bias based on Immuno-Trol targets, and strong agreement in the Bland–Altman analysis. The method successfully passed three CAP external proficiency tests in 2024, confirming interlaboratory reliability. Coefficients of variation for precision were below 10% for all subsets. Linearity exceeded R ² > 0.99 across clinically relevant ranges. Most subsets demonstrated an LLOQ below 10–50 cells/μL, which is suitable for clinical applications. The proposed modifications to the DuraClone IM kit protocol improved workflow efficiency and analytical performance without compromising accuracy or reproducibility. The validated method provides a standardized, reliable, and time-efficient alternative for lymphocyte subset quantification.