Human NK Cells Downregulate Zap70 and Syk in Response to Prolonged Activation or DNA Damage

Jason L. Pugh, Neda Nemat-Gorgani, Paul J. Norman, Lisbeth A. Guethlein and Peter Parham
J Immunol February 1, 2018, 200 (3) 1146-1158; DOI: https://doi.org/10.4049/jimmunol.1700542

Abstract

The extent of NK cell activity during the innate immune response affects downstream immune functions and, ultimately, the outcome of infectious or malignant disease. However, the mechanisms that terminate human NK cell responses have yet to be defined. When activation receptors expressed on NK cell surfaces bind to ligands on diseased cells, they initiate a signal that is propagated by a number of intracellular kinases, including Zap70 and Syk, eventually leading to NK cell activation. We assayed Zap70 and Syk content in NK cells from healthy human donors and identified a subset of NK cells with unusually low levels of these two kinases. We found that this Zap70lowSyklow subset consisted of NK cells expressing a range of surface markers, including CD56hi and CD56low NK cells. Upon in vitro stimulation with target cells, Zap70lowSyklow NK cells failed to produce IFN-γ and lysed target cells at one third the capacity of Zap70hiSykhi NK cells. We determined two independent in vitro conditions that induce the Zap70lowSyklow phenotype in NK cells: continuous stimulation with activation beads and DNA damage. The expression of inhibitory receptors, including NKG2A and inhibitory killer Ig-like receptors (KIRs), was negatively correlated with the Zap70lowSyklow phenotype. Moreover, expression of multiple KIRs reduced the likelihood of Zap70 downregulation during continuous activation, regardless of whether NK cells had been educated through KIR–HLA interactions in vivo. Our findings show that human NK cells are able to terminate their functional activity without the aid of other immune cells through the downregulation of activation kinases.

Introduction

Natural killer cells are lymphocytes of innate immunity that respond rapidly to infection. In this capacity, NK cells influence the outcome of acute infections (e.g., influenza virus) (1). In chronic infections [e.g., hepatitis C virus (2, 3) or HIV (4)], long-term disease progression is influenced by the NK cell response that occurred during the acute phase of infection.

In the early stages of the immune response, NK cells respond to virus-infected cells with proliferation, cytokine production, and the killing of infected cells (3, 5, 6). Studies using mouse models of disease show how NK cells can steer the overall immune response to a pathogen by limiting Ag availability (7) and selectively killing subsets of dendritic cells (8) and T cells (9). The extent and duration of the NK response can also affect long-term immunity, such as T cell memory (10). Following their initial proliferation and population expansion, NK cells decline in number, and their functional capacity is diminished (3, 5, 6).

This contraction and cessation of NK cell–mediated immune responses is influenced by the cytokine environment (11). However, the contributions of other factors are unclear. For instance, a variety of inhibitory receptors help to prevent NK cells from attacking healthy cells (12), but it is not known whether these inhibitory signals also participate in attenuating the effector cytokine and cytolytic responses of human NK cell populations once they are no longer needed. In contrast, the contraction and memory phases of B and T cell responses have been studied extensively (13), as has the contraction of mouse NK cell populations in response to murine CMV (14, 15).

NK cells detect damaged, infected, and malignant cells with an array of activating cell surface receptors (16–18). Upon engaging ligands expressed by target cells, NK cell surface receptors associate with transmembrane adaptor molecules, such as DAP-12 and CD3ζ, which, in turn, associate with Src kinases (19). These kinases initiate an intracellular phosphorylation cascade that leads to NK cell activation. Among the intracellular enzymes that propagate the cascade are other kinases, Zap70 and Syk (20, 21), as well as phospholipase C-γ2 (PLC-γ2) (22).

Strongly influencing the development and response of human NK cells are inhibitory receptors that recognize HLA class I molecules. These comprise the conserved NKG2A receptor, which has a lectin-like binding site that recognizes conserved HLA-E (23), as well as diverse killer cell Ig-like receptors (KIRs) that recognize polymorphic epitopes of HLA-A, HLA-B, and HLA-C (24). During the development of an NK cell, interaction between self–HLA class I and an inhibitory receptor educates the NK cell to be tolerant of healthy cells expressing a normal amount of HLA class I but intolerant of unhealthy cells expressing abnormally low amounts of HLA class I (25).

When bound to HLA class I ligands on healthy cells, NKG2A and inhibitory KIRs recruit tyrosine phosphatases, which inhibit signals coming from activating NK cell receptors (26), likely by direct dephosphorylation of the intracellular proteins involved in the activating pathway (27). The reduced amount of HLA class I on unhealthy cells cannot block the activating signals that instruct the NK cell to attack and kill the unhealthy target cell to which it is bound.

The signaling cascade of NK cell activation and its various components are well studied for the initiation of an NK cell response (19), but the role of these mechanisms in winding down the response of an NK cell population, once it is no longer needed, is unknown. Likewise, although the contribution of inhibitory HLA class I receptors to NK cell activation is defined (28), their role, if any, in terminating NK cell responses is also unknown. We explored these relationships in the current study. In human NK cells obtained from healthy CMV-free donors, we identified a subset of NK cells that expresses unusually low amounts of the Zap70 and Syk kinases, which are key components in the signaling cascade of T cells and B cells (29, 30). We replicate this phenotype in vitro and make connections between the emergence of these Zap70lowSyklow NK cells and the chronic stimulation of activating NK cell receptors.

Materials and Methods

HLA and KIR genotyping

Whole human blood was purchased from the Stanford Blood Center (Stanford, CA). DNA was extracted from whole blood using a QIAamp DNA Blood Mini Kit (QIAGEN, Valencia, CA), following the manufacturer’s instructions. Isolated DNA was HLA and KIR genotyped, as described previously (24). In summary, genomic DNA was sheared, and those fragments comprising complete HLA-A, HLA-B, HLA-C, and KIR genes were isolated using oligonucleotide probes (24). The DNA fragments were subjected to sequencing using an Illumina MiSeq System and v3 technology, with 300-bp paired-end reads (Illumina, San Diego, CA). KIR gene copy number and the identity of KIR and HLA-A, HLA-B, and HLA-C alleles were defined as described (24, 31).

Blood acquisition and processing

Leukoreduction and separation chambers containing 109 PBMCs were obtained from healthy CMV− donors at the Stanford Blood Center. PBMCs were separated from other cells on a Ficoll-Paque gradient (GE Healthcare, Chicago, IL), pelleted, and suspended at 107 per milliliter in FBS (Gemini Bio Products, Sacramento, CA) containing 10% DMSO (EMD Millipore, Billerica, MA). Aliquots were frozen using a Mr. Frosty device (Thermo Fisher Scientific, Waltham, MA) and kept in a −80°C freezer for >24 h before being stored in liquid nitrogen.

For use in experiments, frozen aliquots of PBMCs were thawed at 37°C in a water bath and suspended in 10 ml of RPMI 1640 medium (Corning, Manassas, VA) containing 2 mM l-glutamine (Thermo Fisher Scientific), 100 U/ml penicillin and streptomycin (Thermo Fisher Scientific), and 10% FBS (RPMI 10%-C). DNase I was added to a final concentration of 0.1 mg/ml to prevent cell clumping (Sigma-Aldrich, St. Louis, MO), and the cells were incubated for 30 min at 37°C. To ensure that cell surface markers were expressed normally, the cells were washed and transferred to 12-well plates, at 1.0 × 107 cells per well, and kept in a 37°C incubator with 5% CO2 for ∼12 h before any further manipulation was performed.

NK cell isolation

NK cells were isolated from PBMCs using the Untouched NK Isolation Kit with LS columns, as described by the manufacturer (Miltenyi Biotec, San Diego, CA). With this procedure, other types of PBMCs are depleted using specific Abs. In summary, PBMCs were first resuspended in PBS with 0.5% BSA and 2 mM EDTA (MACS buffer). The Miltenyi Biotec mixture of biotinylated Abs was added, allowing the Abs to bind to their target Ags on PBMCs. On the addition of paramagnetic streptavidin-coated beads, streptavidin on the beads bound to the biotin conjugated to the Abs. The cell and bead mixture was passed through a column in the presence of a magnet, which trapped all PBMCs, with the exception of NK cells, in the column. The flow-through, containing NK cells, was centrifuged to pellet the cells, which were washed and suspended in 1 ml of RPMI 10%-C.

We adapted the manufacturer’s protocol to isolate KIR− NK cells. To do this, the mixture of MACS buffer and biotinylated Abs applied to PBMCs was supplemented with four biotinylated Abs: anti-KIR2D (NKVFS1; Miltenyi Biotec), anti-KIR3DL1 (DX9; BioLegend; San Diego, CA), anti-KIR3DL (5.133; Miltenyi Biotec), and anti-KIR3DL2 (clone 539304; R&D Systems/Thermo Fisher Scientific). Because this Ab mixture depleted all PBMCs, with the exception of KIR− NK cells (and the possible exception of NK cells expressing only KIR2DL4 and/or KIR2DL5 and no other KIR), it resulted in the purification of KIR3DL1− and KIR3DL2− NK cells. In making this modification, we found that it was critical to keep the 107 PBMCs per 50 μl cell/volume ratio of the staining reaction prescribed in the Miltenyi Biotec protocol. Although this volume consists of 20% Miltenyi Biotec Ab mixture and 80% MACS buffer in the Miltenyi Biotec protocol, in our modified protocol it consists of 20% Miltenyi Biotec Ab mixture, 5% each of the four additional Abs, and 60% MACS buffer. No other aspect of the protocol was changed.

NK cell culture and functional assays

NK cells were activated using NK activation beads (Miltenyi Biotec) in flat-bottom 96-well plates (BD Labware, Franklin Lakes, NJ). These beads are coated with streptavidin bound to biotinylated anti-CD2 and anti-NKP46 Abs (Miltenyi Biotec). A total of 100,000 or 200,000 NK cells was placed in each well with an equal number of NK activation beads and 200 μl of RPMI 10%-C medium containing 500 U/ml recombinant human IL-2. IL-2 was obtained from Dr. M. Gately (Hoffmann-La Roche) through the National Institutes of Health AIDS Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases, National Institutes of Health. For assays lasting >24 h, 100 μl of the culture medium was gently removed after 24 h and replaced with 100 μl of fresh RPMI 10%-C medium containing 1000 U/ml IL-2. In some experiments, cell division was tracked using BrdU (Sigma-Aldrich), which was included in the culture medium at a concentration of 10 μM. BrdU was kept at 10 μM in cultures over several days by replacing medium daily, as described above.

In experiments to study DNA damage and apoptosis in NK cells, VP-16 etoposide (Sigma-Aldrich) and/or Z-VAD FMK (BD Biosciences, San Jose, CA) was included in the culture medium at concentrations of 0.25 mg/ml and 15 μM, respectively.

In assays measuring NK cytolysis, NK cells were cultured overnight with 500 U/ml IL-2 in the absence of target cells. K562 target cells were then added to NK cells at a 10:1 NK/target ratio. The mixture of NK cells and target cells was transferred to a U-bottom 96-well plate and pelleted at ∼700 × g. To detect the increased expression of CD107a on the surface of NK cells that are killing targets, the cells were suspended in 200 μl of fresh RPMI 10%-C media containing 500 U/ml IL-2 and 2 μg/ml anti-CD107a Ab (eBioscience, San Diego, CA). To facilitate contact between NK cells and K562 targets, the cocultures were centrifuged at 100 × g for 2 min and placed in a 37°C and 5% CO2 incubator for 3.5 h. Cytolysis was measured as the percentage of NK cells that upregulated CD107a during the incubation period (32).

Assay of IFN-γ production was performed in the same way as the cytolytic assay, with the exception that NK cells were cocultured with K562 target cells for 16 h in flat-bottom 96-well plates. During the final 6 h of this assay, Brefeldin A (Thermo Fisher Scientific) was added to a final concentration of 10 μg/ml to prevent NK cell secretion of IFN-γ.

In assays to test the effect of cytokines on Zap70 loss, NK cells were incubated in RPMI 10%-C with 200 ng/ml IL-12 (eBioscience), 100 ng/ml IL-18 (Thermo Fisher Scientific), or 200 ng/ml IFN-γ (eBioscience). To induce KIR expression, KIR− NK cells were incubated in RPMI 10%-C with 10 ng/ml IL-15 (BioLegend).

Abs

Fourteen fluorescently conjugated mAbs specific for cell surface proteins were used in the isolation and analysis of NK cells: anti-CD3 (HIT3a; BioLegend), anti-CD3 (SP34-2; BD Horizon, BD Biosciences, San Jose, CA), anti-CD4 (A161A1; BioLegend), anti-CD8 (HIT3a; BioLegend), anti-CD14 (M5E2; BD Horizon), anti-CD16 (MHCD1617; Life Technologies), anti-CD19 (HIB19; eBioscience), anti-CD27 (M-T271; BioLegend), anti-CD56 (CMSSB; eBioscience), anti-CD107a (eBioH4A3; eBioscience); anti-KIR2DL1/S1 (A66899; Beckman Coulter, Brea, CA), anti-KIR3DL1 (DX9; BioLegend), anti-KIR3DL2 (clone 539304; R&D Systems), and anti-NKG2A (REA110; Miltenyi Biotec). In addition, unconjugated anti-KIR3DL2 (DX31, a gift from L. Lanier) was identified by fluorescently conjugated anti-mouse IgG (A-11001; Thermo Fisher Scientific). Unconjugated anti-KIR3DL2 (MBB2-2994; Novus Biologicals, Littleton, CO) was detected with fluorescently conjugated anti-rabbit IgG (Poly4046; BioLegend).

Six fluorescently conjugated mAbs were used to detect the following intracellular markers: anti-Zap70 (1E7.2; BioLegend); anti-Syk (4D10.2; BioLegend); anti–PLC-γ2 (K86-1161; BD Phosflow); anti–IFN-γ (45.B3; BioLegend); anti-BrdU (IC7225A; R&D Systems); and anti-BrdU (51-23619L; BD Pharmingen). In addition, biotinylated anti-Zap70 (SBZAP; Abcam) was detected with fluorescently conjugated streptavidin (12-4317; eBioscience).

Cell staining, fixation, and flow cytometry

Staining of PBMCs, with Abs specific for cell surface proteins, was done in U-bottom 96-well plates. Staining took place in a total volume of 50 μl of PBS containing 5% FBS. The cells and fluorescently conjugated Abs were incubated at 4°C in the dark (0.5–12 h). The Ab panels contained a maximum of 9 of the 16 Abs described above. The amount of each Ab used for staining was determined empirically by titration. Typically, 2 μl of the solution provided by the manufacturer was sufficient for one test. During subsequent steps, the reagents and plates were kept on ice. The stained cells in each well were washed two times with 200 μl of PBS. After each wash, the cells were centrifuged at ∼700 × g for 3 min at 4°C. In panels including a secondary anti-mouse IgG Ab, cells were stained in a three-step process to avoid off-target binding. Cells were first stained with the unconjugated primary Ab alone. After two washes, cells were stained with the secondary Ab alone. Following three more washes, cells were stained with a mixture of the other Abs specific for cell surface markers.

To identify and exclude dead cells from analysis, NK cells were stained with LIVE/DEAD Yellow (Thermo Fisher Scientific), according to the manufacturer’s instructions. For each well, the cells were stained for 30 min in 50 μl of a 500-fold dilution of LIVE/DEAD Yellow in PBS. The cells were washed twice and then fixed with 100 μl of BD Cytofix solution (BD Biosciences) for 10 min at 4°C. Next, 100 μl of perm buffer (1× BD Perm; BD Biosciences) was added to the cells, while still in BD Cytofix solution, to permeabilize them. The cells were washed twice with 200 μl of perm buffer. To ensure permeabilization of intracellular compartments, cells were incubated with 200 μl of perm buffer containing 10% DMSO at 4°C for 20 min. Cells were then washed three times with perm buffer. To ensure fixation of intracellular compartments, cells were subjected to a second fixation with BD Cytofix, followed by washing with permeabilization (perm) buffer. Prior to anti-BrdU staining, cells were additionally incubated with perm buffer containing 30 μg/ml DNase I (Sigma-Aldrich) for 30 min at room temperature in the dark. For intracellular Ab staining, cells were stained in perm buffer in a 50-μl volume for 30 min at 4°C, followed by two washes with perm buffer and a final wash in PBS containing 5% FBS.

Intracellular staining of permeabilized cells was performed using the seven mAbs specific for intracellular markers described above. Staining concentrations were determined empirically by titration, but typically 2 μl per test was sufficient. Cells were analyzed using an LSR II flow cytometer (BD) at the Stanford Shared FACS Facility (Stanford, CA). The data obtained were analyzed using FlowJo v9.7.6 (TreeStar, Ashland, OR), and assessed for statistical significance using Prism 6 (GraphPad Software, La Jolla, CA).

Quantitative PCR analysis of NK cell mRNA

NK cells isolated from PBMCs were subjected to LIVE/DEAD staining and then sorted on a BD FACSAria II instrument at the Stanford shared FACS Facility. NK cells from groups of two to four donors were pooled and sorted based on Abs against KIR3DL1, pan-KIR2D, clone 539304, and/or NKG2A into separate tubes containing RPMI 10%-C. The cells were centrifuged, the supernatant was removed, and the cell pellets were processed using an RNeasy Protect Kit (QIAGEN), following the manufacturer’s instructions. RNA was eluted in 30 μl of UltraPure DNase/RNase-Free Distilled Water (Invitrogen). Three microliters of RiboLock RNase Inhibitor (Thermo Fisher Scientific), 3 μl of 50 mM MgCl2 solution (Invitrogen), and 3 U DNase I (Thermo Fisher Scientific) were added to each sample. Incubation of these samples in a 37°C water bath for 30 min digested any residual genomic DNA. To denature the DNase, 3 μl of 0.05 M EDTA (Hoefer) was added to the samples, which were then incubated for 10 min in a 65°C water bath. RNA integrity was assessed using an RNA 6000 Pico Kit and an Agilent Bioanalyzer 2100 (Agilent Technologies, Santa Clara, CA) at the Stanford Protein and Nucleic Acid (PAN) Facility (Stanford, CA). cDNA was synthesized from mRNA using SuperScript VILO Master Mix (Thermo Fisher Scientific). cDNA was subjected to unbiased preamplification using PreAmp Master Mix (Thermo Fisher Scientific) and examined by quantitative PCR (qPCR) using TaqMan Gene Expression Assays for Syk, Zap70, and β-actin (Hs00895377_m1, Hs00896345_m1, and Hs99999903_m1, respectively; Thermo Fisher Scientific) and a StepOnePlus Real-Time PCR System (Applied Biosystems, Foster City, CA) at the Stanford PAN Facility. For each preparation of cDNA, three replicate analyses were performed.

Results

A subset of peripheral blood NK cells has low Zap70, low Syk, and high PLC-γ2

NK cells from different human donors exhibit considerable functional variation that is not entirely due to HLA and KIR differences (33). One possible source of variation is the intracellular kinases that contribute to NK cell activation. To address this question, we used flow cytometry to quantify Zap70 in PBMCs from healthy donors. The CD14−CD19−CD3−CD56midhiCD16midhi cells defined by our NK cell gate exhibit bimodal expression of Zap70 (Fig. 1A). Averaging for 27 donors showed that 7.9 ± 4.9% of NK cells had <10% of the mean Zap70 present in the other ∼92% of NK cells (Fig. 1A, 1B). Less than 2% of NK cells expressed Zap70 at levels between those of Zap70low and Zap70hi NK cell populations (Fig. 1D). The fluorescent signal in the Zap70 channel observed for Zap70low NK cells was higher than that with the isotype control, indicating that Zap70low NK cells express some Zap70 (Fig. 1A, Supplemental Fig. 1A).

FIGURE 1.
FIGURE 1.

A subset of peripheral blood NK cells has low Zap70, low Syk, and high PLC-γ2. (A) Gating strategy to examine Zap70low NK cells in human blood. Upper panels (left to right): FSC-A and SSC-A define PBMCs, FSC-H and FSC-A define singlets, LIVE/DEAD stain and CD14low define leukocytes, and CD3low and CD19low exclude B and T cells. Lower panels (left to right): CD16midhi and CD56midhi broadly define NK cells, KIR2DL1/S1 defines a subset of KIR+ NK cells, and Zap70low cells make up a small portion of KIR2DL1/S1+ NK cells. Plots represent the data obtained from 36 donors. (B) Percentage of Zap70low NK cells from 27 donors, gated on total NK cells or on NK cells positive for any KIR (pan-2D, 3DL1, or 3DL2). ****p < 0.0001, paired t test. (C) Percentage of Zap70-deficient cells in NK cell subsets from 27 donors defined by various surface markers. ***p < 0.001, *p < 0.01, Tukey comparison of one-way ANOVA. (D) Representative NK cell plot of Zap70 versus Syk (left panel). Comparison of Zap70 and Syk phenotypes in NK cell populations obtained from 21 donors (right panel). ****p < 0.0001, paired t test. (E) Representative NK cell plot of PLC-γ2 versus Syk (left panel). Comparison of Zap70 and PLC-γ2 expression in the NK cell populations of 32 donors (right panel). All data are representative of at least five independent experiments; SEM is shown in all panels. **p < 0.001, paired t test.

Because we defined NK cells by negative gating, it was possible that Zap70low cells were not NK cells. Because NK cells are distinguished by expression of KIRs (34), we examined Zap70 expression by KIR+ NK cells. The proportion of KIR+ NK cells that were Zap70low (4.8 ± 3.4%) was comparable to the 7.9% Zap70low cells present in total NK cells (Fig. 1B), consistent with most Zap70low cells being NK cells.

A small subset of peripheral blood T cells with effector memory phenotype expresses KIR (35). These cells might also exhibit low CD3 expression as a consequence of activation. To determine whether these cells corresponded to Zap70low NK cells, PBMCs were stained with T cell–specific Abs. Those Abs were then bound to paramagnetic beads, and T cells were removed (>98%) from other PBMCs by MACS. The proportion of Zap70low cells within the KIR+ NK cell gate was not reduced by T cell depletion (Supplemental Fig. 1B), showing that KIR+Zap70low NK cells were not KIR+ T cells.

Zap70 expression was independently assessed with two Zap70-specific mAbs. Both Abs identified a similar proportion of Zap70low cells (7.1–7.6%) in isolated NK cell populations (Supplemental Fig. 1C). For a subset of donors, PBMC samples were divided equally into two subsamples: one was analyzed immediately for Zap70low NK cells, and the other was frozen and thawed prior to Zap70 analysis. No significant difference was observed in the frequency of Zap70low NK cells in these two preparations, although freeze-thawing decreased the variance between donors of Zap70low NK cells (Supplemental Fig. 1C). Zap70low NK cells were present in all eight NK cell subsets defined by combinations of high and low expression of CD56, CD16, CD27, NKG2A, and LILRB1 (Fig. 1C, Supplemental Fig. 1E). Of these markers, NKG2A expression was the best correlate of Zap70 expression. Among NKG2Alow NK cells, 8.2 ± 5.4% were Zap70low, whereas only 1.6 ± 1.4% of NKG2Ahi NK cells were Zap70low.

NK cell expression of Syk and PLC-γ2, other enzymes that contribute to NK cell activation, were similarly examined. A strong correlation (p < 0.0001) was observed between Syk and Zap70 expression. Thus, almost all Zap70low NK cells were also Syklow. Only small numbers of NK cells with SyklowZap70hi and SykhiZap70low phenotypes were detected (Fig. 1D). In contrast, PLC-γ2 expression was not positively correlated with Zap70 expression (p < 0.01) (Fig. 1D, 1E).

Zap70lowSyklow NK cells have less functional potential than Zap70hiSykhi NK cells

Subsets of NK cells from the same donor, and varying in their expression of Zap70 and Syk, were compared for their capacity to respond to HLA class I–deficient K562 cells with accumulation of CD107a, a surrogate measure of the degranulation that occurs during target cell killing (36). Zap70hiSykhi NK cells produced a strong response, accumulating CD107a in 31.2% of the cells (Fig. 2A). In contrast, Zap70lowSyklow NK cells from the same donors exhibited a feeble response: only 2.4% of the cells accumulated CD107a. Thus, low levels of Zap70 and Syk correlated with severe functional impairment of NK cells. NK cells expressing only one of the two kinases had a functional capacity that was between that of Zap70lowSyklow and Zap70hiSykhi NK cells. On average, 14.7% of Zap70lowSykhi NK cells and 14.4% of Zap70hiSyklow NK cells responded to K562 targets. In summary, high expression of both kinases correlated with strong cytotoxic potential, low expression of both kinases correlated with weak cytotoxic potential, and high expression of one kinase correlated with intermediate cytotoxic potential.

FIGURE 2.
FIGURE 2.

Zap70lowSyklow NK cells have less functional potential than Zap70hiSykhi NK cells. (A) NK cells isolated from 11 donors were primed for 24 h with IL-2 and cocultured with K562 target cells, at a 10:1 E:T ratio, for 3.5 h in the presence of anti-CD107a Ab and IL-2. (B) NK cells from the same 11 donors were cocultured with K562 target cells at a 10:1 E:T ratio for 8 h in the presence of IL-2. For the last 6 h, brefeldin A was included in the culture to block secretion. Accumulation of IFN-γ was then measured by flow cytometry. (C) Isolated NK cells from 26 donors were primed for 24 h with IL-2 and IL-12, and cells were cocultured with K562 target cells at a 10:1 E:T ratio for 16 h. For the last 6 h, brefeldin A was included in the culture to block secretion. Accumulation of IFN-γ was measured by flow cytometry. All data represent at least three independent experiments, each with different donors; SEM is shown in all panels. ****p < 0.0001, ***p < 0.001, *p < 0.05, Tukey comparison from a paired ANOVA.

To compare the cytokine response of Zap70hiSykhi and Zap70lowSyklow NK cells, unprimed NK cells were cocultured with K562 targets for 8 h in the presence of IL-2, and cellular accumulation of IFN-γ was measured. On average, 17% of Zap70hiSykhi NK cells produced IFN-γ compared with 9.7% of Zap70lowSyklow NK cells (Fig. 2B). In the assay of cytokine production, the difference between Zap70hiSykhi and Zap70lowSyklow NK cells was less than a factor of 2. In contrast, the difference between the cytotoxic responses was >10-fold. NK cells expressing only one of the two kinases had weaker responses than Zap70lowSyklow NK cells: 4.8% of Zap70lowSykhi NK cells and 2.1% of Zap70hiSyklow NK cells responded to K562 with the production of IFN-γ (Fig. 2B, 2C).

To determine whether Zap70lowSyklow NK cells make IFN-γ in response to inflammatory conditions, NK cells were cultured with IL-2 and IL-12 for 24 h. Thus primed, the NK cells were cocultured with K562 targets for 16 h. For flow cytometry, the gate was set for IFN-γ+ NK cells based on control NK cells that were primed with cytokines but not exposed to K562 cells. Under these conditions, only 0.1% of Zap70lowSyklow NK cells produced IFN-γ. In contrast, an average of 23% of Zap70hiSykhi NK cells made IFN-γ in response to K562 cells (Fig. 2C). In this assay, the difference between the responses of Zap70lowSyklow and Zap70hiSykhi NK cells was >200-fold.

Zap70 and Syk loss is induced by extended NK cell activation

To determine whether NK cells downregulate Zap70 and Syk as a consequence of producing an immune response, we cocultured NK cells and K562 targets for 3.5 h, a period sufficient for NK cell activation and target cell cytolysis. NK/target cells were used at 2:1, 5:1, 10:1, and 20:1 ratios. No significant differences in the frequency of Zap70lowSyklow NK cells were observed between cultures with different E:T ratios or with cultures of NK cells alone (Fig. 3A). Thus, loss of Zap70 and Syk was not induced by a short-term culture of NK cell activation and target cell cytolysis.

FIGURE 3.
FIGURE 3.

Zap70 and Syk loss is induced by extended NK cell activation. (A) Frequency of Zap70lowSyklow NK cells among isolated NK cells that were cocultured for 24 h with K562 target cells at various NK/K562 ratios. Shown are the results from six donors. Representative of three independent experiments. No significant differences were found, as assessed by the Tukey posttest. (B) Frequency of Zap70low or Syklow NK cells from nine donors after isolated NK cells were cocultured for 3 d with activation beads coated with anti-CD2 and anti-NKp46 Abs or with anti-DNAM1 and anti-NKG2D Abs. Cocultures were performed at a 2:1 NK cell/bead ratio in RPMI-10%-C media with 500 U/ml IL-2. Representative data are shown from at least three independent experiments. (C) Kinetics of Zap70 loss following incubation with anti-NKp46 and anti-CD2 activation beads, as in (B). NK cells and activation beads were cocultured at a 2:1 ratio. Shown are the data combined from four experiments involving the NK cells of 24 donors. (D) Frequency of Zap70lowSyklow NK cells from eight donors, following coculture or not with K562 target cells at various target to effector (T/E) ratios. Cocultures occurred for 1 or 3 d. Additional K562 cells were added on days 1 and 2 for the 3 d 1:5 ratio group. Representative of three independent experiments. (E) Comparison of Zap70 loss following a 24-h incubation of isolated NK cells in RPMI-10%-C media supplemented with various cytokine combinations. Shown are the combined data from three experiments, in which NK cells of 16 donors were studied. (F) Comparison of the percentage of divided cells (BrdU+) in Zap70low versus Zap70hi NK cells on day 3 of bead stimulation, as in (C), for six donors. Representative data from at least three independent experiments. (G) Correlation of gMFI of Zap70 signal in KIR+ NK cells of six donors, as in (C), with gMFI of BrdU signal in KIR+ NK cells that have divided, as in (C). Representative of three independent experiments, each with different donors. SEM is shown in all panels. ****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05, Tukey posttest from paired one-way or two-way ANOVA, as appropriate (B, E, and G); paired t test (F).

To assess the effect of long-term chronic stimulation of NK cells, one set of beads was coated with anti-CD2 and anti-NKp46 Abs, and a second set of beads was coated with anti-DNAM1 and anti-NKG2D Abs. Each bead has the potential to engage two activating NK cell receptors and, thereby, activate NK cells (37). Flow cytometry showed that isolated NK cell preparations in this experiment had an average of 3.0 ± 4.0% Syklow NK cells and 1.3 ± 0.9% Zap70low NK cells prior to any manipulation (Fig. 3B). NK cells were cultured with each bead set separately for 3 d in the presence of IL-2. Of the NK cells stimulated through their CD2 and NKp46 receptors, 49.3 ± 15.6% were Zap70low, and 45.8 ± 15.7% were Syklow. Of the NK cells stimulated through their DNAM1 and NKG2D receptors, 41.7 ± 14.7% were Zap70low, and 34.2 ± 14.5% were Syklow. Following culture with either set of activation beads, the frequencies of Zap70low and Syklow NK cells were significantly greater (p < 0.0001) than those observed before culture (Fig. 3B). Statistical comparison also showed that CD2/NKp46 stimulation resulted in significantly more Zap70 and Syk loss than DNAM1/NKG2D stimulation (Fig. 3B).

To compare the kinetics of Zap70 and Syk downregulation, NK cells were cocultured with the two activation beads for different times (0, 1, 3, 5, and 7 d) and analyzed for Zap70 and Syk expression. By including BrdU in the culture medium, we could determine the extent of NK cell division that took place. To distinguish NK cells from beads, our analysis focused on KIR+ NK cells. During 1 d of coculture, the frequency of SyklowKIR+ NK cells increased from 3.5 ± 3.4 to 26.6 ± 4.7% (Fig. 3C). A similar downregulation of Zap70 required 3 d of coculture, at which point 26.6 ± 14.1% of KIR+ NK cells were Zap70low. That Syk loss occurs more quickly than Zap70 loss raises the possibility that, in each cell, Syk is lost first and Zap70 is lost second. After 5 d of coculture, the frequency of Syklow NK cells had increased to 35.4 ± 7.4%, whereas the percentage of Zap70low NK cells was 24.1 ± 10.8%, not significantly different from that on day 3 (Fig. 3C).

To complement the above experiment, in which NK cells were stimulated with bead-bound Abs, we stimulated NK cells with live K562 cells and assessed the loss of Syk and Zap70. NK cells were cocultured with K562 target cells at a 5:1 or 1:1 ratio. During 1 d of coculture, most target cells were killed, but no loss of Syk or Zap70 was observed in NK cells. To increase the stimulation, NK and K562 cells were cocultured for 3 d. To maintain stimulation, fresh K562 target cells were added each day. Under these conditions, there was significant loss of Syk and Zap70. With an E:T ratio of 5:1, the frequency of Zap70lowSyklow NK cells increased from 2 to 13.8% (Fig. 3D). With an E:T ratio of 1:1, the frequency of Zap70lowSyklow NK cells further increased to 41.8 ± 30.6%. In summary, chronic stimulation of NK cells by human cells lacking HLA class I or by Abs bound to activating NK cell receptors induces significant loss of Syk and Zap70 in the NK cell population.

Because a 3 d stimulation with bead-bound anti-receptor Abs was necessary to achieve Zap70 loss, we explored the possibility that inflammatory cytokines produced by NK cells accumulate over several days in media and cause Zap70 and Syk loss. In examining the effect of such cytokines, IL-18 and IFN-γ had no effect, and IL-12 slightly increased the frequency of Zap70low NK cells from 4.4 to 6.4% (Fig. 3E). These results indicate that inflammatory cytokines are not a significant cause of NK cell loss of Zap70.

We cocultured NK cells and beads coated with only one ligand, either anti-CD2 or anti-NKp46. This activation by single receptors did not increase the proportion of Zap70lowSyklow cells compared with activation by both receptors. Instead, it resulted in fewer Zap70lowSyklow cells: 10.2% from CD2-mediated activation and 20.7% from NKp46-mediated activation, compared with 23.7% from activation by both receptors (Supplemental Fig. 2A). This result, together with the results shown in Fig. 3B, supports a model in which stronger forms of chronic activation produce higher frequencies of NK cells that are deficient for Zap70 and Syk.

To determine whether activated NK cells could induce Zap70 loss in unstimulated NK cells, we measured the frequency of Zap70low NK cells, after 3 d of stimulation by beads coated with anti-CD2 and anti-NKp46, among NK cells exclusively expressing the NKp44, NKp80, or NKp46 activation receptor. NKp46+ NK cells made up 23.93% of Zap70low NK cells compared with a combined average of 16.95% Zap70low for NKp44+ and NKp80+ NK cells (Supplemental Fig. 2B). These results point to Zap70 loss being a direct consequence of ligand engagement by the activation receptors and not a bystander effect.

We assessed whether Zap70 downregulation induced by the stimulating beads correlates with NK cell division and proliferation. An analysis of BrdU and Zap70 expression showed that the Zap70low phenotype primarily occurred in NK cells that had divided during the period of stimulation (Fig. 3F). To determine whether the extent of Zap70 downregulation correlated with the number of NK cell divisions, we compared the geometric mean fluorescence intensity (gMFI) of BrdU in NK cells that had divided with the Zap70 gMFI in all NK cells for each donor. Higher BrdU gMFI was correlated with lower Zap70 gMFI across donors (Fig. 3G), showing that Zap70 loss correlates with increasing proliferation.

The results of these various analyses point to chronic engagement of activation receptors as one cause of the Zap70lowSyklow phenotype in NK cells. Additionally, our results imply that the Syklow phenotype occurs prior to the Zap70low phenotype and that both phenotypes can be passed from mother to daughter cells.

Zap70 and Syk loss is a consequence of DNA damage but not of caspase activation

To determine how DNA damage affects Zap70 and Syk expression, NK cells were treated with etoposide (VP-16). This drug causes DNA breaks, leading to caspase-mediated apoptosis (38). Treating NK cells with etoposide for 24 h activated caspase 3, a component of apoptotic signaling, in 22.6% of the cells (line graphs in Fig. 4A, averages in Fig. 4B). Etoposide also induced loss of Zap70 (Fig. 4A) and Syk (Fig. 4B); their expression was lost by 32.3 and 34.2% of NK cells, respectively (Fig. 4B). That the frequency of Zap70low and Syklow NK cells was greater than that of cells activating caspase 3 suggested that loss of Zap70 and Syk was not caused by apoptosis-induced proteolysis.

FIGURE 4.
FIGURE 4.

Zap70 and Syk loss is a consequence of DNA damage but not of caspase activation. (A) Representative flow cytometry plots showing the proportions of isolated NK cells positive for anti-cleaved caspase 3 Ab (upper panels) or NK cells that are Zap70low (lower panels). NK cells from a single donor treated with DMSO (left panels, control) or with 0.25 mg/ml VP-16 (right panels) in DMSO, for 24 h. Data are representative of 17 donors. (B) Percentage of Syklow, Zap70low, or cleaved caspase 3+ in isolated NK cells from nine donors following 24 h of treatment with 0.25 mg/ml VP-16 etoposide or DMSO control. (C) Percentage of cleaved caspase 3+ NK cells from six donors following 24 h of treatment with DMSO control, 15 μM Z-VAD-FMK caspase inhibitor alone, 0.25 mg/ml VP-16 etoposide, or Z-VAD-FMK inhibitor plus VP16. All cultures contained 500 U/ml IL-2. (D) Percentage of Zap70low NK cells following 24 h of treatment, as in (C), for seven donors. All data are representative of at least three independent experiments using NK cells from different donors; SEM is shown in all panels. ****p < 0.0001, ***p < 0.001, *p < 0.05, Tukey posttest from paired one-way or two-way ANOVA, as appropriate.

As another test for involvement of apoptosis-induced proteolysis, NK cells were treated with Z-VAD FMK, a universal caspase inhibitor (39). In the context of etoposide treatment, Z-VAD FMK effectively blocked caspase cleavage (Fig. 4C). However, treating NK cells with Z-VAD FMK alone had no effect on the frequency of Zap70low NK cells (Fig. 4D). Treating NK cells with etoposide and Z-VAD FMK caused a more variable Zap70 loss than that caused by etoposide alone, but the frequency at which NK cells lost Zap70 was not significantly different for treatment with etoposide alone or combined with Z-VAD FMK (Fig. 4D). Like Zap70, the frequency of Syk loss caused by etoposide was unaffected by caspase blockade (Supplemental Fig. 2C). These results suggest that Zap70 and Syk loss is caused by the DNA damage response but is independent of caspase-mediated apoptosis.

Inhibitory KIR and NKG2A protect NK cells from losing Zap70 and Syk

The frequency of Zap70low cells is higher for KIR− NK cells than for KIR+ NK cells (Figs. 1B, 5A). Moreover, NK cells defined by the binding of anti-KIR3DL1 or anti-KIR2DL1/S1 have a higher frequency of Zap70low NK cells than NK cells binding both Abs. Thus, NK cells expressing KIR3DL1 and KIR2DL1/S1 are less likely to downregulate Zap70 than NK cells expressing just one of these receptors.

FIGURE 5.
FIGURE 5.

Inhibitory KIR and NKG2A protect NK cells from losing Zap70 and Syk. (A) Comparison of the percentage of Zap70low NK cells within various KIR+ subsets, as measured by flow cytometry following NK cell isolation in 11 donors. Data are representative of three independent experiments using NK cells from different donors. (B) Comparison of Zap70 expression in NK cells expressing KIR3DL1, KIR3DL2, or any KIR2D (KIR+) or not expressing any inhibitory KIR (KIR−) in relation to Zap70 and NKG2A in isolated NK cells from 25 donors. Representative of three independent experiments using NK cells from different donors. (C) Relative transcript levels of Zap70 and Syk measured by qPCR and normalized to β-actin transcript levels, derived from pools of sorted NK cells (NKG2Ahi/KIR+ or NKG2Alow/KIR−). Transcript amounts are represented as a percentage of the average normalized transcript amount present in the NKG2Ahi/KIR+ group. Representative from eight donors, acquired from three separate sorts and three technical replicates. SEM is shown in all panels. ****p < 0.0001, ***p < 0.001, **p < 0.01, Tukey posttest from paired one-way or two-way ANOVA, as appropriate.

A similar hierarchy was observed for the pan-KIR2D Ab and KIR3DL1 Abs. The frequency of Zap70low NK cells was lowest in the subpopulation of NK cells binding both Abs, highest in NK cells binding only anti-KIR3DL1, and intermediate in NK cells binding only anti–pan-KIR2D (Fig. 5A). These results initially suggested that educated NK cells were less likely to be Zap70low than uneducated NK cells.

We similarly examined Zap70 and Syk loss in NK cells expressing NKG2A and/or KIR. Of NK cells lacking expression of inhibitory KIR and expressing low amounts of NKG2A, 36.6 ± 20.8% were Zap70lowSyklow (Fig. 5B). The NKG2AhiKIR− subpopulation of NK cells contained 6.5 ± 4.8% Zap70lowSyklow cells, a frequency similar to the 7.5 ± 5.9% of Zap70low cells in the NKG2AlowKIR+ subpopulation of NK cells (Fig. 5B). Thus, the presence of NKG2A or an inhibitory KIR appeared sufficient to retain Zap70 and Syk expression. The subpopulation of NK cells that coexpress NKG2A with an inhibitory KIR had the lowest frequency of Zap70lowSyklow cells (3.5 ± 3.1%) (Fig. 5B).

One possible cause for the differences in kinase expression between Zap70hiSykhi and Zap70lowSyklow NK cells is transcriptional regulation. We tested this hypothesis for NKG2Alow/KIR− NK cells, because this subpopulation had the highest frequency of ex vivo Zap70lowSyklow cells (36.6 ± 20.8%). NK cells were sorted into those coexpressing NKG2A and KIR and those lacking KIR and having low levels of NKG2A (Fig. 5B). These two pools were adjusted to contain the same number of NK cells, and RNA was extracted. RNA from several donors was combined and used to make cDNA, which was subjected to real-time qPCR with primers amplifying Zap70, Syk, and β-actin, a housekeeping gene for normalization.

Zap70 transcript levels in NKG2AlowKIR− NK cells were 81% of those in NKG2AhiKIR+ NK cells (Fig. 5C). Syk transcripts in NKG2AlowKIR− NK cells were 113% of those in NKG2AhiKIR+ NK cells (Fig. 5C). If transcriptional regulation was the primary cause of Zap70 and Syk loss, we estimated that NKG2AlowKIR− NK cells should display ≤50% of the Zap70 and Syk transcripts of NKG2AhiKIR+ NK cells. This conservative estimate was based on the ≥27-fold difference in the amounts of Zap70 and Syk proteins in Zap70hiSykhi and Zap70lowSyklow cells (example graphs in Fig. 1A, 1D) and the 36.6% frequency of Zap70lowSyklow NK cells in the NKG2AlowKIR− NK cell population (Fig. 5B). Given that NKG2AhiKIR+ NK cells have a lower level of Syk transcripts than NKG2AlowKIR− NK cells, as well as a level of Zap70 transcripts that is only 19% higher than that of NKG2AlowKIR− NK cells, we conclude that the ex vivo differences in Zap70 and Syk protein levels between Zap70hiSykhi and Zap70lowSyklow NK cells are not solely due to transcriptional regulation.

KIR expression prevents loss of Zap70 in a ligand-independent manner

We studied the education of NK cells expressing KIR3DL1, by the cognate Bw4 ligand that is carried by subsets of HLA-A and HLA-B allotypes (33). Analysis focused on the subpopulation of NK cells for which KIR3DL1 is the only inhibitory KIR. We found no significant difference in the frequency of Zap70low NK cells between donors who have the Bw4 epitope and donors who lack it (Fig. 6A).

FIGURE 6.
FIGURE 6.

KIR expression prevents loss of Zap70 in a ligand-independent manner. (A) Comparison of the frequency of Zap70lowKIR3DL1+ single-positive NK cells from donors who carry the Bw4 epitope (n = 9) or who lack the Bw4 epitope (n = 13). “Weak Bw4” denotes a donor with KIR3DL1*001, KIR3DL1*015, HLA-B*52:01, and HLA-A*24:02, and “strong Bw4” denotes a donor with two copies of KIR3DL1*001, HLA-B*51:01, and HLA-A*32:01. Data represent at least three replicate experiments. NS, Student t test. (B) Representative plots of NK cells prior to the MACS depletion of KIR+ NK cells using anti-KIR3DL1, anti-KIR3DL2, and anti–pan-KIR2D Abs (left panel), following the depletion of KIR+ NK cells (middle panel), and following 24 h of treatment with IL-15 (right panel). Representative of the data from 18 donors and at least three replicate experiments. (C) Comparison of the frequency of Zap70low cells from eight donors within KIR3DL1 single-positive NKG2A− NK cells and KIR−NKG2A− NK cells, prior to MACS depletion of KIR+ NK cells (ex vivo) or following KIR induction by IL-15 on KIR− NK cells. p < 0.0001, two-way ANOVA of KIR3DL1 comparison. Data are representative of three independent experiments, each with different donors. (D) Isolated NK cells from four donors were cocultured with class I–deficient 221 target cells or 221 cells expressing transfected HLA-B*58:01 in the presence of 500 U/ml IL-2 at a 10:1 E:T ratio. Frequency of CD107a+ and Zap70low KIR3DL1+ NK cells after 3.5 h of coculture. Representative of three independent experiments. NS, Tukey posttest from two-way ANOVA. (E) Kinetics of different KIR+ NK subsets during extended stimulation with activation beads, as in Fig. 3C. Data were obtained from the NK cells of 17 donors. Representative data are from three combined experiments. SEM is shown in all panels. ***p < 0.001, Tukey posttest from two-way ANOVA.

Avidity of the interaction between KIR3DL1 and Bw4+ is modulated by the polymorphism of KIR3DL1, HLA-A, and HLA-B and affects NK cell education (40). We assessed its effect on the population of Zap70low NK cells by determining high-resolution HLA class I and KIR genotypes for our donor panel (Supplemental Figs. 3, 4) and comparing the frequency of Zap70low NK cells in the two donors predicted to have the strongest and weakest interactions of Bw4 with KIR3DL1, as predicted from published data (40). The donor with the weakest interactions had KIR3DL1*001, KIR3DL1*015, HLA-B*52:01, and HLA-A*24:02 (Fig. 6A, weak Bw4), whereas the donor with the strongest interactions had two copies of KIR3DL1*001, HLA-B*51:01, and HLA-A*32:01 (Fig. 6A, strong Bw4). That the weakest interaction is associated with 1.6% Zap70low NK cells and the strongest interaction is associated with 1.7% Zap70low NK cells showed that the strength of NK cell education does not contribute to the loss or retention of Zap70. This conclusion was consistent with our analysis of Bw4+ and Bw4− donors as a whole, which did not show any significant difference in Zap70 levels due to the presence or absence of the Bw4 epitope. In contrast, the expression of KIR by NK cells does favor retention of Zap70, irrespective of its participation, or not, in NK cell education.

To test whether KIR expression is sufficient to induce expression of Zap70, we isolated KIR− NK cells using a modified magnetic separation technique and cultured them with IL-15 for 24 h. This treatment induced KIR expression (41) (Fig. 6B). We then examined those NK cells with newly expressed KIR3DL1 but lacking other KIR and NKG2A. The proportion of Zap70low cells in this subpopulation of NK cells was similar to that observed in the same subpopulation of ex vivo NK cells, but it was significantly different from the proportion of Zap70low NK cells lacking KIR3DL1, other KIR, and NKG2A following IL-15 treatment (Fig. 6C). Although it is possible that other factors that influence Zap70 expression were induced by IL-15, only the cells induced to express KIR showed increased Zap70 expression.

We examined whether binding of KIR3DL1 to Bw4+HLA-B could increase NK cell expression of Zap70. Isolated NK cells were cocultured for 3.5 h with 221 target cells, which do not express class I HLA, or with transfected 221 cells expressing Bw4+HLA-B*58:01. Assaying for CD107a accumulation showed that the presence of HLA-B*58:01 reduced the frequency of CD107a-producing NK cells gated on KIR3DL1+ from 28.1 to 4.3% (Fig. 6D). This result shows that NK cell KIR3DL1 bound to HLA-B*58:01 on the target cells and generated inhibitory signals. Such signals were not produced in NK cells cultured with 221 cells, which lack Bw4. In contrast, we observed no significant difference in the frequency of Zap70low cells in KIR3DL1+ NK cells cultured with 221 cells or 221 cells expressing HLA-B*58:01 (Fig. 6D).

To test whether KIR expression protected activated NK cells from loss of Zap70, we tracked Zap70 expression across a broad range of KIR+ NK cell subsets during extended culture with activation beads. In these cultures, the relationships between specific KIR subsets and the extent to which they downregulated Zap70 did not fully replicate those seen for ex vivo NK cells (compare Fig. 5A with Fig. 6E). However, coexpression of KIR3DL1 and KIR2D by NK cells was significantly more protective against stimulation-induced Zap70 loss than expression of only KIR3DL1 or KIR2D (Fig. 6E). This protection occurred even though NK cells coexpressing KIR3DL1 and KIR2D had undergone more division than the cells expressing KIR3DL1 or KIR2D (Supplemental Fig. 2D). These results are consistent with KIR expression providing protection against Zap70 loss.

A KIR3DL2-specific Ab marks Zap70lowSyklow NK cells and T cells

Analysis of three anti-KIR3DL2 Abs showed that mouse mAb 539304 and a rabbit polyclonal Ab have an overlapping specificity, with 539304 binding to many fewer NK cells than the polyclonal Ab (Supplemental Fig. 2E). The mouse mAb DX31 binds to a different subpopulation of NK cells than the other two Abs (Fig. 7A).

FIGURE 7.
FIGURE 7.

The KIR3DL2 target of Ab 539304 marks Zap70low NK cells and T cells. (A) Frequency of NK cells, from five donors, that are recognized by the DX31 and 539304 monoclonal anti-KIR3DL2 Abs and a polyclonal anti-KIR3DL2 Ab. Representative data from at least three independent experiments, each with different groups of donors. (B) Percentage of Zap70low NK cells in five donors identified by DX31 or 539304, as in (A). Representative of at least three independent experiments using different donors. (C) Comparison of the frequency of Zap70low cells in the NK cell subsets identified by anti-KIR3DL1, anti–pan-KIR2D, or 539304 Abs for NK cells of 34 donors. Representative of three independent experiments. (D) Frequency of NK cells binding the 539304 Ab, among various subpopulations of NK cells that express hi or low NKG2A. NK cells from 29 donors were studied. Representative of three independent experiments. (E) Binding of the 539304 Ab to NKG2A+, pan-KIR2D+, and KIR3DL1+ NK cells obtained from the 29 donors. Donors are grouped into those that have the A3/11 epitope of HLA-A and those that lack this epitope. Representative of three replicate experiments. (F) Comparison of the frequencies of Zap70low NK cells expressing KIR3DL1, but not KIR3DL2, and of NK cells that bind the 539304 Ab but do not express KIR3DL1, from five donors, prior to MACS depletion of KIR+ NK cells (ex vivo) or following MACS depletion of KIR+ NK cells and subsequent KIR induction by IL-15. Representative of three independent experiments, each with different donors. (G) Comparison of gMFI of Zap70 in CD4+ T cells, CD8+ T cells, and NK cells from six donors expressing KIR3DL1 or expressing the KIR3DL2 epitope recognized by 539304 as the only KIR. Result is representative of three independent experiments, each with different donors. (H) Levels of Zap70 and Syk transcripts measured by qPCR, normalized to the transcript levels of β-actin. Data were derived from pools of cells that were sorted to be 539304+DX9− or 539304−DX9+. Transcript amounts are represented as a percentage of the average normalized transcript in 539304−DX9+ NK cells. Data are representative of those obtained from the NK cells of 12 donors from three independent sorts and three technical replicates. (I) For all NK cells studied, gMFI of Zap70 is correlated with the square root of the frequency of NK cells binding the anti-KIR3DL2 Ab 539304 (n = 34). Data are representative of three technical replicates of subsets of donors. SEM is shown in all panels. Test of linear fit = p < 0.0001. ***p < 0.001, Sidak comparison of two-way ANOVA.

Nearly all NK cells bound by Ab 539304 are Zap70low (Fig. 7B, 7C). NK cell subpopulations expressing NKG2A, KIR3DL1, or the combination of KIR3DL1 and KIR2D have higher frequencies of 539304+ NK cells than other NK cells (Fig. 7D). To determine whether NK cell education affected the proportion of 539304+ NK cells, we divided the donor cohort into two groups: donors having the HLA-A3/11 epitope recognized by KIR3DL2 and donors lacking the A3/11 epitope. Both groups of donors were found to have similar frequencies (Fig. 7E), showing there is no effect of KIR3DL2-mediated education on expression of the epitope recognized by the 539304 Ab.

Culturing KIR− NK cells with IL-15 induced de novo expression of KIR and of the epitope recognized by Ab 539304. 539304+ NK cells were nearly all Zap70low (Fig. 7F). Similar analysis of T cells (35) identified small mutually exclusive subsets expressing either the KIR3DL2 epitope recognized by 539304 or the KIR3DL1 epitope recognized by the DX9 Ab (Supplemental Fig. 2F). CD4 and CD8 T cells bound by 539304 were also noted to have lower Zap70 expression when analyzed for gMFI (Fig. 7G).

Because the epitope recognized by 539304 appears to mark cells lacking Zap70, we investigated whether 539304+ NK cells express Zap70 or Syk transcripts. Groups of 539304+DX9− NK cells, as well as 539304−DX9+ NK cells, were obtained from several donors and pooled. RNA was extracted from the two pools and reverse transcribed to give cDNA, which was analyzed by real-time qPCR for Zap70 and Syk transcripts, as described above. That 539304+DX9− NK cells contained far fewer Zap70 and Syk transcripts than 539304−DX9+ NK cells (Fig. 7H) indicates that the Zap70 and Syk genes are not expressed in 539304+ NK cells, which, in this regard, are different from other ex vivo Zap70lowSyklow NK cells (compare Fig. 5C with Fig. 7H).

Expression of the 539304 epitope and Zap70 content were assessed in a cohort of 34 donors. The best linear fit for an x–y plot of these two parameters showed an inverse correlation between the square root of the percentage of NK cells bound by the 539304 Ab and the median NK cell expression of Zap70 (Fig. 6F). Thus, NK cell populations with higher levels of Zap70 expression tend to have lower numbers of clone 539304+ NK cells.

Discussion

We describe a subset of human NK cells having low levels of the Zap70 and Syk kinases, critical factors for NK cell activation. Consistent with this role, Zap70low/Syklow NK cells are functionally deficient. Although Zap70low/Syklow cells make up a small fraction of NK cells ex vivo, the Zap70low/Syklow phenotype can be induced in vitro by subjecting NK cells to prolonged stimulation with beads coated with Abs specific for activating NK cell receptors. Characterization of Zap70low/Syklow NK cells was facilitated by a mouse mAb 539304, which was raised against KIR3DL2 and binds specifically to the surface of Zap70low/Syklow NK cells and T cells.

Populations of ex vivo NK cells consist of cells expressing Zap70 and Syk at various levels. NK cells having high levels of both kinases were strongly cytolytic, whereas NK cells having high levels of one kinase achieved only half this response, and NK cells having high levels of neither kinase were barely cytolytic. Thus, Zap70 and Syk are noted to make additive contributions to NK cell cytotoxicity. Downregulation of Zap70 and Syk is predominantly a property of NK cells for which an activating receptor has engaged its cognate ligand. Cell division is also highly associated with Zap70 loss, but it is unlikely to be its cause, because only some cells that lose Zap70 during prolonged stimulation have divided. Our observations favor a mechanism in which sustained activation of receptor signaling is the primary cause of Zap70 and Syk downregulation.

Paolini et al. (42) demonstrated that human NK cells downregulate Zap70 and Syk following CD16 engagement through ubiquitination and proteolysis. We now note that this is a more general phenomenon, induced by sustained engagement of multiple activating NK cell receptors. Syk loss by ex vivo NK cells caused by CD16 binding to obinutuzumab correlated with a functional shift toward IFN-γ production (43). In our study, we found that loss of Syk and Zap70 in vitro correlates with a reduction in IFN-γ production.

Our observations suggest that the degree to which Zap70 and Syk are downregulated in an NK cell population correlates directly with the extent of NK cell activation. Previous analysis of the reduction in Zap70 and Syk expression caused by CD16 engagement fits with this trend (42), because CD16 is the one receptor that can activate NK cells without involvement of another activating receptor (37). Chronic stimulation of mouse NK cells by NKG2D causes downregulation of the associated DAP-10 and DAP-12 signaling proteins and impedes other unrelated activation pathways (44, 45). Such studies point to chronic engagement affecting the abundance of proteins that contribute to various stages in the activation pathway. Our findings also suggest that Zap70 and Syk do not need to participate in signaling for their downregulation to occur following stimulation. This is reflected by the fact that engagement of DNAM1 and NKG2D induced Zap70 and Syk loss, but neither activation receptor is known to propagate signals through Zap70 or Syk in human NK cells (46, 47). The connections among activating receptor signaling, kinase loss, and functional cessation raise the possibility that these events combine to form a physiological mechanism for limiting the extent and duration of the NK cell response. Because an enhanced NK response can limit T cell memory (7), targets and mechanisms for limiting NK responses are of increasing relevance to vaccine design (48).

Our analysis shows that expression of an inhibitory NK cell receptor acts to prevent NK cells from losing Zap70 and Syk, an effect observed for ex vivo NK cells and NK cells stimulated in vitro. NK cells expressing NKG2A, inhibitory KIR, or LILRB1 are all protected from losing Zap70 and Syk. Such protection does not require the presence of the inhibitory receptor’s cognate HLA class I ligand and, thus, is independent of NK cell education. We also observed that NK cells expressing more types of inhibitory KIR had correspondingly increased retention of Zap70 and Syk, suggesting a direct and quantitative relationship between inhibitory receptor expression and Zap70/Syk retention. However, Zap70 and Syk are unlikely to interact directly with the ITIMs of these inhibitory receptors, particularly when those receptors are not engaged (49, 50). Further work will be necessary to determine how inhibitory receptors influence the recycling of proteins in the activation pathway.

Various models have been proposed to explain how NK cells become self-tolerant (51–53). That prolonged NK cell activation causes loss of Syk and Zap70, and that this can be prevented by the presence of an inhibitory receptor, seems compatible with the model in which NK cells lacking inhibitory receptors become “disarmed” as a consequence of chronic stimulation of the activating receptors (25). However, our finding that inhibitory receptor engagement does not affect Zap70 and Syk levels does not support this model. Our observations suggest a new model in which human NK cells use dual mechanisms to achieve self-tolerance. In the first mechanism, NK cells that do not express sufficient inhibitory receptors have low amounts of Zap70 and Syk. In the second mechanism, NK cells that express an inhibitory receptor may be enhanced through education, which takes place when inhibitory receptors bind their cognate ligands. This educational effect likely occurs through a mechanism that is independent of Zap70 and Syk but might involve the redistribution of activation receptors at microdomains on the cell surface (54). Educated or not, mature peripheral NK cells increase their chances of losing Zap70 and Syk after sustained engagement with activation ligands.

In allogeneic NK cancer immunotherapies, the transfer of educated donor NK cells into a host with uneducated NK cells mediates a graft-versus-leukemia response, causing a significant reduction in mortality (55). Our findings suggest that selective transplantation of NK cells expressing more KIR, even ones irrelevant to the educational context, may help to retain NK cell functions over days of stimulation. When cancer patients are treated with radiotherapy, their NK cells are more likely than their T cells to die from radiation damage (56). Moreover, the population of NK cells that survives irradiation suffers a dramatic loss of function (57). Results from our study raise the possibility that this functional deficiency is caused by NK cell downregulation of Syk and Zap70.

Although it is well established that Zap70 and Syk are essential for T cell activation (30), their precise contributions to NK cell function are less clearly defined. Mutation in human Zap70 can cause SCID (30, 58) that is characterized by the absence of CD8 T cells, functionally defective CD4 T cells, and normal numbers of NK cells (59, 60). Study of NK cells from one such patient showed that they lyse K562 target cells effectively (60). The NK cells of mice deficient for both Zap70 and Syk can also lyse various tumor cells, despite their inability to signal through ITAM-bearing receptors (61). Several lines of evidence point to Zap70 and Syk as having distinctive functions in humans and mice (58, 62). In this study, we have uncovered a natural and coordinated downregulation of Zap70 and Syk in human NK cells that profoundly affects their function.

Syklow NK cells are associated with latent human CMV (HCMV) infection (63, 64). In this context, NK cells with the Syklow phenotype had normal Zap70 expression. Our analysis of HCMV− donors shows that dynamic control of Syk and Zap70 is not limited to latent HCMV infection but appears to be part of normal NK cell regulation.

Patients with hepatitis B (65), melanoma (66), or lung adenocarcinoma (67) have functionally deficient NK cells, which are characterized by high surface expression of Tim-3. In contrast, the functionally deficient Zap70lowSyklow NK cells that we describe have chronically low Tim-3 expression (Supplemental Fig. 1F), as was also seen for Syklow NK cells of HCMV-infected individuals (63, 64). Further investigation should define connections between NK cell exhaustion and the regulation of Zap70 and Syk in specific disease contexts.

KIR3DL2 is distinguished from other KIRs in that it is expressed predominantly as a dimer at the cell surface (68). The observed differential reactivity of anti-KIR3DL2 Abs could be explained if Ab 539304 is specific for the monomeric form of KIR3DL2. One function of KIR3DL2 is to bind microbial CpG-oligodeoxynucleotides at the NK cell surface and transport them to endosomes, where they are recognized by TLRs (69). If dimeric, but not monomeric, KIR3DL2 binds CpG-oligodeoxynucleotides, chronic activation of NK cells by CpG-oligodeoxynucleotides could cause endocytosis of dimeric KIR3DL2, as well as downregulation of Zap70 and Syk, leaving 539304-reactive monomers as the only form of KIR3DL2 at the NK cell surface.

Enhancement and inhibition of NK cell activity are increasingly likely to prove valuable as immunotherapies in different disease contexts. Realizing these goals requires knowledge of mechanisms that activate NK cell populations in the immune response and of mechanisms that contract and terminate their activity when they are no longer needed. In this article, we have shown that NK cells can reduce their functional role through downregulation of Syk and Zap70 kinases.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank Zakia Djaoud, Amir Horowitz, Emily Wroblewski, and Hugo Hilton for technical and scientific input, Cathy Crumpton and Ometa Herman (Stanford Shared FACS Facility) for flow cytometry and sorting support, and Alberto Lovell (Stanford PAN Facility) for qPCR support and services.

Footnotes

  • This work was supported by National Institutes of Health Grant R01 AI22039 (to P.P.).

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    gMFI
    geometric mean fluorescence intensity
    HCMV
    human CMV
    KIR
    killer Ig-like receptor
    PAN
    Protein and Nucleic Acid
    PLC-γ2
    phospholipase C-γ2
    qPCR
    quantitative PCR.

  • Received April 17, 2017.
  • Accepted November 15, 2017.

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