Robust multi-parameter single-platform quantification of myeloid and B-lymphoid CD34 progenitor cells in all clinical CD34 cell sources and in thawed PBSC.

As B-lymphoid progenitor cells do not give rise to in vitro colony formation and are unlikely to support myeloid engraftment, we validated a five-color extension of the single platform Stem Cell Enumeration (SCE) kit, to routinely quantify myeloid and B-lymphoid progenitor cells. Fresh samples (n > 20 each) of granulocyte colony stimulating factor mobilized blood (peripheral blood (PB)), cord blood (CB), bone marrow (BM), and apheresis products (APs) were stained in TruCOUNT™ tubes and the results were compared with those from the two-color CD45/CD34 reagent combination and the three-color SCE kit. To address repeatability, 10 samples from one AP were prepared by four technicians. Aliquots (n = 15) of four frozen AP were analyzed after thawing. Excellent correlations were observed between the three kits (R(2) > 0.99), for the quantification of white blood cells and total CD34. The extended kit showed considerable amounts of B-lymphoid progenitors in all CD34 sources (0-20% of all CD34 in PB, AP, and CB; 3-90% in BM). Very similar results were obtained when the same sample was prepared by different technicians. After thawing of frozen AP, the recovery of viable cells varied depending on the freezing medium employed, but the results from the different quantification methods were identical. Most non-viable cells were clearly identified with 7 Aminoactinomycin D (7AAD) but an additional gate in the forward scatter/side scatter was necessary to address dead cells negative for 7AAD. The extended SCE kit allows rapid and exact quantification of viable B-lymphoid and myeloid CD34(+) cells in all cell sources and in thawed stem cell harvests, and may thus improve the correlation between CD34 number and engraftment kinetics.

A flow cytometric method for platelet counting in platelet concentrates.

BACKGROUND: The platelets (PLTs) in PLT concentrates are counted with hematology analyzers, but varying results among different hematology analyzers are observed, making comparisons very difficult. Due to the absence of red blood cells in PLT concentrates, the International Council for Standardization in Hematology (ICSH) reference method was modified to be used for PLT concentrates and validated in an international comparative study. STUDY DESIGN AND METHODS: Five PLT samples were shipped to eight participating centers of the Biomedical Excellence for Safer Transfusion (BEST) Collaborative and counted on the same day. PLTs were stained with fluorescein isothiocyanate-labeled anti-CD41a in tubes (TruCount, BD Biosciences), measured on a flow cytometer, and analyzed with a uniform template. These samples were also counted on 15 hematology analyzers. RESULTS: The ICSH method and newly developed BEST method yielded PLT counting results with less than 1% difference (not significant). The intercenter coefficient of variation (CV) of the BEST method was on average 6.3% versus 7.6% on average for hematology analyzers. The CV of individual hematology analyzers was on average 0.9%, which was considerably lower than for the flow cytometers with a mean of 3.7%. CONCLUSION: The BEST flow cytometric method has a smaller intercenter CV and a smaller center-to-center deviation from the group mean compared to hematology analyzers. Conversely, individual hematology analyzers are more precise than the flow cytometric method. Thus, the flow cytometric method provides a calibration tool to allow comparisons between centers, but there is no need to replace routine counting with hematology analyzers.

A new one-platform flow cytometric method for residual cell counting in platelet concentrates.

BACKGROUND: According to German regulations and guidelines, residual red blood cells (rRBCs) and residual white blood cells (rWBCs) must number fewer than 3 x 10(9) cells/unit and 1 x 10(6) cells/unit in platelet concentrates (PCs), respectively. Due to low levels of residual cells in final products, there is still a need for fast, reliable, and sensitive methods of automated detection of these cell types. STUDY DESIGN AND METHODS: In Part A, 21 PCs were spiked with predetermined numbers of red blood cells (RBCs) and white blood cells (WBCs). The linearity, precision, and accuracy of the BD Thrombo Count assay (BD Biosciences Europe) were tested and validated according to international guidelines. Finally in Part B, 100 PCs prepared from pooled buffy coats were tested by the BD Thrombo Count assay and compared with other methods, including Nageotte (rWBCs) and Neubauer (rRBCs) counting chambers and the flow cytometric BD LeucoCOUNT (Becton Dickinson) assay (rWBCs). RESULTS: The unspecific background of blank PC samples was fewer than 0.02 cells/microL for WBCs and fewer than 34 cells/microL for RBCs (mean, 21). Linear regression and precision analyses of spiked PC samples were determined for both WBCs (r(2) = 0.992; range, 0.6-6.0 WBCs/microL) and RBCs (r(2) = 0.999; 800-8000 RBCs/microL). No carryover of cells or drift in results was detected in the automated sample acquisition mode. Analysis according to statistical methods of Bland and Altman demonstrated a high correlation between BD Thrombo Count and the Neubauer manual counting chamber. CONCLUSION: This novel flow cytometric test is a quick and reliable single-tube assay that has been demonstrated as a potential alternative for the existing manual microscopic counting procedures that are both time-consuming and laborious.

Flow cytometric assay for the simultaneous determination of residual white blood cells, red blood cells, and platelets in fresh-frozen plasma: validation and two years' experience.

BACKGROUND: A fully automated single-tube assay with tubes (BD TruCOUNT, BD Biosciences) for absolute counting of residual cells in freshly prepared plasma by flow cytometry was developed (BD Plasma Count). STUDY DESIGN AND METHODS: The nucleic acid dye thiazole orange stains white blood cells (WBCs). The monoclonal antibodies anti-CD41a-peridinin chlorophyll protein-Cy5.5 and anti-glycophorin A-fluorescein isothiocyanate label platelets (PLTs) and red blood cells (RBCs), respectively. No fixation, permeabilization, or washing steps were required. Validation was done according to guidelines of the International Conference on Harmonization and the National Committee for Clinical Laboratory Standards. Cell-free plasma was spiked with each cell type for accuracy, reproducibility, and linearity measurements. RESULTS: Results showed no carryover or drift under automated sample acquisition conditions. Nonspecific background was fewer than 0.3 cells per microL for residual WBCs (rWBCs), fewer than 2.7 cells per microL for rRBCs, and fewer than 85 cells per microL for rPLTs. Determinations of rWBC and rPLT counts were linear with a coefficient of variation of less than 12 percent for the imprecision. Owing to cross-linking of the anti-glycophorin A antibody, linearity and precision for rRBCs diverged up to 21 percent at a count of 6000 rRBCs per microL. In a 2-year period, five operators investigated 2666 quality control (QC) samples of fresh-frozen plasma on 108 working days. Maximum cell numbers found were 196 for rWBCs, 3960 for rRBCs, and 28,952 for rPLTs per microL. In 31 cases (1.2%) rWBCs were out of specification. No outlier was observed for rRBCs and rPLTs. Residual RBC cell numbers determined were always within the acceptable concentration range of the assay. CONCLUSION: These data demonstrate that the single-tube test is suitable for routine QC assessment of the cellular contaminants of therapeutic plasma according to the European recommendations.

Bacteria detection by flow cytometry.

Since bacterial infection of the recipient has become the most frequent infection risk in transfusion medicine, suitable methods for bacteria detection in blood components are of great interest. Platelet concentrates are currently the focus of attention, as they are stored under temperature conditions, which enable the multiplication of most bacteria species contaminating blood donations. Rapid methods for bacteria detection allow testing immediately before transfusion in a bed-side like manner. This approach would overcome the sampling error observed in early sampling combined with culturing of bacteria and would, at least, prevent the transfusion of highly contaminated blood components leading to acute septic shock or even death of the patient. Flow cytometry has been demonstrated to be a rapid and feasible approach for detection of bacteria in platelet concentrates. The general aim of the current study was to develop protocols for the application of this technique under routine conditions. The effect of improved test reagents on practicability and sensitivity of the method is evaluated. Furthermore, the implementation of fluorescent absolute count beads as an internal standard is demonstrated. A simplified pre-incubation procedure has been undertaken to diminish the detection limit in a pragmatic manner. Additionally, the application of bacteria detection by flow cytometry as a culture method is shown, i.e., transfer of samples from platelet concentrates into a satellite bag, incubation of the latter at 37 degrees C, and measuring the contaminating bacteria in a flow cytometer.

A comparison of three rapid bacterial detection methods under simulated real-life conditions.

BACKGROUND: Bacterial screening of all produced platelet concentrates (PCs) is implemented in many countries to reduce the risk of transfusion-transmitted sepsis. This study compares three rapid bacterial detection methods by imitating real-life conditions. STUDY DESIGN AND METHODS: The sensitivity of a solid-phase scanning cytometer (optimized Scansystem, Hemosystem), fluorescence-activated cell sorting (FACS) analysis, and 16S RNA in-house nucleic acid testing (NAT) was evaluated by spiking PCs with four transfusion relevant bacteria (Staphylococcus aureus, Bacillus cereus, Klebsiella pneumoniae, and Escherichia coli ). Two different inocula (10 colony-forming units [CFUs]/mL and 10 CFUs/bag) were used to simulate real-life conditions. Samples were taken at 12, 16, 20, and 24 hours after spiking. RESULTS: With the high inoculum, NAT had a 100 percent rate of positive testing for all four types of bacteria (10/10 replicates) at each time point. With the exception of E. coli, the sensitivity of FACS and optimized Scansystem was comparable for the high inoculum. With the low inoculum, 60 percent of E. coli, 80 percent of B. cereus, 90 percent of K. pneumoniae, and 100 percent of S. aureus were NAT-positive 12 hours after spiking. In contrast, only 20 percent of E. coli, 10 percent of B. cereus, and 70 percent of K. pneumoniae were FACS-positive with the low inoculum 12 hours after spiking. CONCLUSIONS: In summary, the preliminary data revealed a higher sensitivity for NAT in comparison to FACS and optimized Scansystem under the defined study conditions. To imitate real-life conditions, further spiking studies with a low inoculum (10 CFUs/bag) and slower growing organisms should be conducted to examine the sensitivity of available detection systems.

Sterility testing of platelet concentrates prepared from deliberately infected blood donations.

BACKGROUND: In general the bacterial count in freshly donated blood is low and even lower in the corresponding platelet concentrates (PCs). By use of flow cytometry (FACS) for sterility testing, the reliability of early versus later sampling times was evaluated. STUDY DESIGN AND METHODS: Blood donations were spiked with various numbers of Staphylococcus epidermidis, Staphylococcus aureus, Bacillus cereus, and Klebsiella pneumoniae. The corresponding PCs were prepared by the buffy-coat method and stored at 22 degrees C. A 20-mL sample was collected from each PC directly after preparation and after 8 hours. Samples were stored at 35 degrees C. Sterility testing of both PCs and samples was by FACS analysis at different time points. RESULTS: All stored PCs were found positive by FACS analysis, with detection times ranging between 8 and 24 hours (K. pneumoniae, B. cereus), 8 and 91 hours (S. aureus), and 144 hours (S. epidermidis). In the samples incubated at 35 degrees C, bacteria were detected after 8 to 19 hours (K. pneumoniae, B. cereus), 8 to 67 hours (S. aureus), and 19 to 43 hours (S. epidermidis). Some of the samples did not contain bacteria. CONCLUSION: Detection times for slow-growing bacteria are significantly shortened when PC samples are incubated at 35 degrees C: the numbers of bacteria in freshly prepared PCs may, however, be so low that the samples drawn for sterility testing do not contain a single bacterium. Our results do not support a shortening of the 24-hour or greater sampling time recommended by the manufacturers of established test systems, because also for consistent detection by FACS, bacteria need to grow in the PCs to sufficient numbers.

Basics of flow cytometry-based sterility testing of platelet concentrates.

BACKGROUND: Flow cytometry (FACS) is a common technique in blood banking. It is used, for example, for the enumeration of residual white blood cells in plasma and in cellular blood products. It was investigated whether it can also be applied for sterility testing of buffy coat-derived platelet concentrates (PCs). STUDY DESIGN AND METHODS: Plasma-reduced PCs were spiked with bacteria and stored at 20 to 24 or 37 degrees C for various times. The following 10 species were used: Bacillus cereus, Enterobacter cloacae, Escherichia coli, Klebsiella pneumoniae, Propionibacterium acnes, Pseudomonas aeruginosa, Staphylococcus aureus, Staphylococcus epidermidis, Serratia marcescens, and Yersinia enterocolitica. Bacterial DNA was stained with thiazole orange. After the platelets were lysed, bacteria were enumerated by FACS. RESULTS: All bacteria species used were detectable by FACS. The lower detection limit was approximately 100 bacteria per microL, that is, 10(5) per mL. In general, the titers measured were 1.2- to 3-fold higher than those determined by colony forming assay. In one case (K. pneumoniae) in which the dot plot of the bacteria cloud overlapped with that of bacteria debris, they were consistently lower. When PC samples were inoculated with approximately 1 colony-forming unit per mL of bacteria and kept at 37 degrees C, most species were detected within 21 hours or less. Exceptions were E. cloacae and P. acnes, which were detected after 24 to 40 and 64 hours, respectively. At 20 to 24 degrees C, the detection times were strongly prolonged. CONCLUSION: Sterility testing of PCs by FACS is a feasible approach. The present data suggest incubating PC samples for 20 to 24 hours at 37 degrees C before testing. For slow-growing bacteria, the incubation period must be prolonged by 1 to 2 days.