<scp>OMIP</scp>‐<scp>063</scp>: 28‐Color Flow Cytometry Panel for Broad Human Immunophenotyping

Abstract

A 28-color panel was developed to screen for a range of lymphocyte subsets in human peripheral blood mononuclear cells (PBMCs), particularly in patients with primary immunodeficiency (PID). Using this panel, we are able to avoid running the sample over multiple screening panels while still deeply phenotyping a diverse range of lymphocyte subsets including innate like lymphocytes (γδ, mucosal-associated invariant T [MAIT], natural killer [NK], and NKT cells), as well as multiple subsets of naïve and memory CD4+ and CD8+ T cells, and B cells. Specifically, naïve, central memory (cmem) and effector memory (emem) CD4+ T cells, naïve, cmem, emem and CD45RA+ revertant memory (TEMRA) CD8+ T cells, regulatory (Tregs), T follicular helper (Tfh) and T helper (Th) 1, and Th17 CD4+ T cells, and transitional, naïve, memory, and CD21lo B cells. Primary immunodeficiencies (PIDs) are caused by monogenic mutations that compromise the development, maturation, differentiation and/or function of immune cells 1, 2. Consequently, individuals with PIDs are highly susceptible to infection by a wide range of pathogens. One of the first steps to understanding the underlying cause of the clinical presentation of patients with PID is to identify any developmental or functional lymphocyte defects that may be present. This is important because several instances have been described where, despite the genetic lesion being determined, the mechanism by which defects in the affected gene result in disease pathogenesis are unknown. Thus, the establishment of “immune cell signatures” for different PID cohorts by flow cytometric-based profiling of immune cell populations in the peripheral blood of PID patients will aid in the diagnosis of these patients and reveal potential cellular defects that may contribute to disease in affected individuals 3. By way of example, patients with heterozygous dominant-negative mutations in STAT3 present with an autosomal dominant form of hyper IgE syndrome (AD-HIES) characterized by recurrent opportunistic Candida (fungal) and staphylococcal (bacterial) infections and compromised antibody responses. Over the years, we and others have established that patients with AD-HIES display a distinctive lymphocyte signature including reduced MAIT and NKT, but normal γδ T cells 4, reduced CD4+CCR6+ T helper (Th)17 and CD4+CXCR5+ circulating T follicular helper (cTfh) cells 5-7, and reduced CD20+CD27+ memory B cells 3, 8. Moreover, the failure to generate Th17 cells in AD-HIES directly accounts for increased susceptibility to fungal infections, while decreased memory B cells and Tfh cells explain the poor antibody responses in these individuals. Previously, this type of screening process would be performed using five different 7–10-color flow cytometric panels (Online Table 3). However, this can be time-consuming and requires more of each sample for a reliable readout. This is especially true when dealing with infrequent cell populations such as the innate like lymphocytes NKT, MAIT, and γδ T cells. Furthermore, access to a large number of PBMCs is not always feasible when dealing with rare patient samples. Hence, there is a currently unmet need for the establishment of a multi-parameter panel to enable detection of different lymphocyte populations in human peripheral blood using a single panel. In this OMIP, a flow cytometric panel that incorporates 28 different markers has been developed and optimized for the efficient screening of patients presenting with PIDs. The panel focuses on identifying the main subsets of B, CD4+ and CD8+ T cells and NK cells as well as NKT, γδ and MAIT cells present in peripheral blood (Tables 1-3). In such a way, this panel can detect and quantify the major lymphocyte subsets in addition to determining their maturation status. For initial broad characterization, we have separated the B and T cell lineages by CD20 and CD3, respectively. Within the CD3 positive compartment, the T cells are then further divided into CD4+ and CD8+ subsets to allow for recognition of the distinct subsets of helper and cytotoxic T cells, respectively. Within the CD4+ and CD8+ T cell populations, the panel includes monoclonal antibodies to numerous markers to enable determining the maturation, differentiation and activation status of these T cells (Figure 1). CCR7 and CD45RA are included to delineate CD4+ and CD8+ T cells into naïve (CCR7+CD45RA+), central memory (CCR7+CD45RA−), effector memory (CCR7−CD45RA−) and effector memory revertant (CCR7−CD45RA+) populations 9. Naïve CD4+ T cells have the ability to differentiate into different helper subsets that have specific roles in immune responses to specific pathogens. For instance, Th1, Th2 and Th17 cells are essential for host defense against viruses and intracellular pathogens, extracellular pathogens, and fungi, respectively. Furthermore, CD4+ regulatory T cells (Tregs) are necessary to maintain immune homeostasis and to guard against autoimmunity, and Tfh cells are essential for the induction of antibody-mediated immunity and long-lived serological memory 10-12. To distinguish between these Th cell populations, we have used the surface receptors CD25, CD45RA, CD127, CXCR5, CXCR3 and CCR6 to identify Tregs (CD25hiCD127lo), Th1 (CD45RA−CXCR3+CCR6−), Th17 (CD45RA−CXCR3−CCR6+), Tfh (CD45RA−CXCR5+), and other Th cells including Th2 and Th9 cells (CD45RA−CXCR3−CCR6−) 11. In regards to CD8+ T cells, the inclusion of killer cell lectin like receptor G1 (KLRG1) has the added benefit of detecting T cell senescence in memory and effector CD8+ T cells 9. B cells (CD20+) develop in the bone marrow and have the important function of producing antibodies and are thus critical for intact humoral immune responses. Cell surface molecules have been used to identify distinct stages of B cell maturation and differentiation. As such, CD10, CD21, CD27, IgD, IgM, IgG and IgA can be used to resolve transitional (CD10+CD27−), naïve (CD10− CD27−), IgM+ memory (CD10−CD27+IgM+IgG−IgA−), and IgG (CD10−CD27+IgM−IgG+IgA−) and IgA (CD10−CD27+IgM−IgG−IgA+) isotype switched memory B cells 13-16. In a lot of instances, we can also detect a population of atypical or aged memory B cells (CD19highCD21low) that have been proposed to have multiple functions in health and disease, including being plasmablast precursors during anamnestic immune responses, harboring self-reactive specificities in the setting of Ab-mediated autoimmune disease, and being dysfunctional in the setting of chronic infection (e.g., HIV, malaria, Hepatitis B) and thus pathogenic due to an inability to clear these pathogens 17, 18. Finally, we have included additional markers to distinguish subsets of NK cells and innate-like T cells. These include γδ T (TCR Vαβ−Vγδ+), NK (CD3−CD56+) NKT (TCR Vα24JαQ+), and MAIT (TCR Vα7.2+CD161+) cells 19. We have also included the PD1 and KLRG1 as markers of activation and senescence, respectively. Following the outline above, we have defined the various immune cells subsets using this 28-color panel (Table 3). The gating strategy used to determine these cells is shown in Figure 1 following standard FSC/SSC, singlet, and live cell gating. Using this broad phenotyping panel, differences in the main lymphocyte subpopulations between healthy donors and patients with immune dysregulatory conditions are easily identified. Differences in a particular population can then be further studied using refined, more specific staining panels. This panel includes a few T cell subsets and differentiation markers used in OMIPs 009, 013, 017, 018, 025, 030, 033, 036, 042, 050; and B cell markers used in OMIP 003, 033, 043, 047, and 051 and innate like populations in OMIP 019, 021, and 046. While some panels such as OMIP 033 and 042 included a few different subsets of lymphocytes, and OMIP 044, OMIP 050 and OMIP 051 are previously published 30-parameter panels with broad phenotyping, there is currently no OMIP that allows for an extensive immunophenotyping of different human lymphocytes as described here. We thank Dr. Andrew Lim of BD Biosciences for providing custom conjugated test antibodies and advice on panel fluorochrome design, Drs John Wotherspoon and Bob Balderas for providing assistance and reagents to assist with instrument characterization and Prof. Stuart G. Tangye for critical review of this manuscript. R.S. is supported by the ISAC shared Resource Laboratory emerging leader program 2014–2018. C.S.M. is supported by grants and fellowships awarded by the National Health and Medical Research Council of Australia (1138359, 1139663, 1139865, 1127157, 1124681) and the Office of Health and Medical Research of the New South Wales State Government of Australia. Data S1 Online Materials. Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.

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