Introduction: Why consider a flow cytometric differential?
Flow cytometry underlies the technology commonly used in many modern hematology analyzers; however, beyond confirming a suspected case of leukemia or lymphoma, immunophenotyping by flow cytometry does not play a major role in the day to day activities in most hematology laboratories. Several investigators have proposed expanding the role of flow cytometry immunophenotyping in the clinical hematology laboratory by using flow cytometry to perform a white blood cell (WBC) count and differential. The current reference method for white blood cell differential counts is manual microscopy(1). Although accurate and reproducible in most cases, manual counts are both time and labor intensive, and can be difficult in patients with low WBC counts. In addition, manual counts provide relatively poor enumeration of populations present at low numbers. Some of these difficulties are addressed using instruments such as CellaVision DM1200, which capture images of cells for technologists to classify. However, manual differentials may have additional problems not addressed by such instruments. For instance manual differentials may not be accurate in enumerating cells that are not uniformly distributed across a slide and cells with unusual morphology may be misclassified. Several of these problems would be addressed by using a WBC differential generated by flow cytometry immunophenotyping. Use of flow cytometry would allow evaluation of many more cells than possible by morphology, allowing for more accurate enumeration of populations present at low numbers. Additionally, flow cytometry allows for specific positive identification of cell types, thereby decreasing misclassification of cells with unusual morphology.
Several groups have investigated flow cytometric methods for performing a differential count (2-5) with good correlations reported with the manual differential. Ideal reagents for a flow cytometry differential would allow specific identification of all cell types of relevance and, to minimize specimen requirements, would be performed using a single tube. Our group recently investigated an 8-color, single-tube, lyse, no-wash flow cytometric method to perform an extended 8 part differential as a potential replacement reference method for WBC differential. This method has many potential applications including validation of new hematology automated analyzers and clarifying cases with difficult morphology. In addition, this method has the potential to supplement or perhaps ultimately replace either the manual differential or current methods used to perform instrument automated differential counts in the clinical laboratory. This article reviews our method for performing a flow cytometry based differential, discusses the performance of this assay, and brings up potential pitfalls of using such a method.
Methodology: How could one perform a flow cytometric differential?
To investigate the flow cytometry differential, we compared the flow cytometry WBC count and differential to a manual differential generated per CLSI H20-A2 (200 cell differential on each of two slides) and, when available, an automated WBC count and differential generated using a Sysmex XE2100 on a test group of 300 peripheral blood samples ranging from relatively normal to complex. Specifically, the study group included 102 random peripheral blood samples meeting criteria for acceptance of an automated instrument differential in the clinical hematology laboratory, 85 samples failing criteria for acceptance of an instrument differential (requiring technologist review or manual differential prior to reporting), and 113 samples collected at random from a hematology/oncology outpatient clinic. The methods for performing the flow cytometric differential are detailed in reference 6 and the gating strategy is outlined in Figure 1. Briefly, sample preparation was as follows: 100 ul of whole blood was stained with 75 ?l of a cocktail of monoclonal antibodies consisting of CD45 APC-Cy7, CD16+CD19 FITC, CD33+CD64 PE-Cy5, CD123 PE, HLA-DR PE-Cy7, CD34+CD117 APC and CD38 A594 and 25 ul of a 100 ug/ml solution of the membrane permeant DNA binding dye Hoechst 34580. Following incubation for 15 minutes at room temperature in the dark, 1.5 ml of buffered NH4Cl containing 0.25% ultrapure formaldehyde was added and incubated for an additional 15 minutes at room temperature in the dark. The samples were prepared in TruCount tubes (Becton Dickinson) with accurate reverse pipetting of the blood to allow generation of an absolute count for all WBC populations. The samples were acquired on an LSRII (BD, San Jose, CA) with a goal of 100,000 nucleated events collected per sample.
Figure 1. Methods: Generating the flow cytometric differential (Adapted from reference 6, Clinical Cytometry)
The flow cytometry differential was performed using the following 8 color panel in conjunction with TruCount beads to generate an 8 part differential as outlined below: CD45 APC-Cy7, CD16+CD19 FITC, CD33+CD64 PE-Cy5, CD123 PE, HLA-DR PE-Cy7, CD34+CD117 APC and CD38 A594
a. A gate was drawn around TruCount beads on the CD123 versus SSC histogram as shown. Then monocytes (pink) were isolated from the viable nucleated cells using CD33+CD64, HLA-DR, and CD16+CD19.
b. Monocytes were then excluded, and of the remaining cells, basophils (purple) and plasmacytoid dendritic cells (light blue) were identified and then excluded using expression of CD123 (bright on both populations) and HLA-DR, which is positive on plasmacytoid dendritic cells and negative on basophils.
c. Of the remaining cells, CD45 and side scatter were used to generate a rough lymphoid cell gate that was purified by removing CD33+CD64 positive events. The lymphocytes were then separated into B cells (CD19+, HLA-DR+ shown in green), NK cells (CD16+, HLA-DR- shown in dark blue) and T cells (lymphoid cells lacking CD19 and CD16, shown in red), which were combined to give the lymphocyte enumeration.
d. The lymphocytes were then excluded leaving mature neutrophils, immature granulocytes, eosinophils, blasts, and nucleated red blood cells. Mature neutrophils (green) were identified on the basis of CD16 expression. Eosinophils (orange) were separated from immature granulocytes and blasts on the basis of CD33+CD64 and CD45 expression in conjunction with side scatter. Eosinophils were then excluded and the immature granulocytes were isolated from the remaining cells on the basis of side scatter versus expression of CD33+CD64 with forward scatter low debris being excluded from the immature granulocyte cell gate (this latter plot is not shown).
e. Blasts were identified using CD45 versus side scatter characteristics in conjunction with expression of CD34+CD117 (blue). Finally, nucleated red cells (aqua) were isolated from the remaining nucleated cells on the basis of expression of DNA binding dye versus forward scatter.
Performance: What are the benefits and potential pitfalls of the flow cytometric differential?
The flow cytometric differential as described in this study, performs well when compared to the manual differential (6) and shows similar efficacy as has been reported in prior studies evaluating flow cytometry to generate a WBC differential, (3,5) confirming the utility of this approach. Best correlations with morphology were seen with enumeration of neutrophils, lymphocytes, mononcytes, eosinophils, and blasts. Poorest correlations were seen in enumeration of basophils and immature granulocytes. These latter populations were present in relatively low numbers in all studied groups, therefore, difficulties in identifying these two populations are likely due to bias by morphology as only 400 cells were counted to determine the morphologic differential. Similar difficulties in enumerating immature granulocyte(3,5) and basophil(3,5) populations by morphology have been reported in prior studies.
The ability to identify and accurately quantify populations present at low numbers and/or not readily identified by morphology could allow the flow cytometry differential as described in this assay to perform tasks that the morphologic differential cannot. For instance, the flow cytometry differential as it is currently described could assist in identification and quantification of very small abnormal plasma cell populations in the peripheral blood of patients with myeloma. Additionally, the flow cytometry differential could be used to identify and quantify plasmacytoid dendritic cells, CD16 positive monocytes (3), or CD64 positive neutrophils (7), three populations that are not distinguishable by morphology but have been noted to change number in various states of disease and health. It should be noted however, that although the potential exists, we have not specifically evaluated the performance of this assay for such hypothetical functions at this point.
A notable source of discordance highlighted in this study between flow cytometry and morphologic classification is misclassification of abnormal lymphoid cells in patients with B cell neoplasms as blasts by morphology. As neoplastic lymphoid cells may take on a blastoid morphology (in particular in cases including CLL with increased prolymphocytes, blastoid mantle cell lymphoma, or large cell lymphoma), such cells may easily be misclassified as blasts by morphology. Such problems of misclassification are easily overcome by antibody driven, specific cell identification methods characteristic of flow cytometry immunophenotyping. The utilization of specific markers, in this case CD34 and CD117, for blast identification provides a significant improvement over morphology; however, it should be noted that not all blast populations express these markers (for instance, monocytic leukemias, and some lymphoblastic leukemias may be negative), therefore refinement of an optimal strategy for blast identification is warranted.
One potential pitfall of using an antibody based method to calculate a WBC count and differential results as antibody expression may be altered in various states. For instance, several problems may be encountered when using CD16 to identify neutrophils. The flow cytometric differential method we employed utilized strong expression of CD16 to highlight mature neutrophils. CD16 expression may be decreased or absent in neutrophils of patients with paroxysymal nocturnal hemoglobinuria (PNH) as CD16 is a GPI linked protein (see this months CSI case for more details about PNH). In patients with PNH, the flow cytometric differential may misclassify CD16 negative neutrophils as eosinophils. Similarly, expression of some antigens we have incorporated into this assay have been described as altered in various clinical states. For instance, CD16 expression has been described as decreased on neutrophils in patients following trauma. The impact of such physiologic alteration of antigen expression on the assay is unclear. Future investigations will evaluate the use of the flow cytometric differential in various states of disease and health.
Sindhu Cherian, MD
Assistant Professor, Laboratory Medicine
Associate Director of the Hematopathology Laboratory
University of Washington, Seattle, WA, USA.
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