Killer Cell Lectin-like Receptor G1 Inhibits NK Cell Function through Activation of Adenosine 5′-Monophosphate–Activated Protein Kinase

The increase in human life expectancy is associated with a greater incidence and severity of many infections and malignancy in older subjects (1–4). The identification of mechanisms that are responsible for the immune decline is therefore essential for the rationalization of ways to improve health during later life. The expansion of T lymphocytes after immune activation takes a finite time, resulting in a lag phase before sufficient numbers of T cells are generated (5). During this period, the host is vulnerable to infections that spread rapidly and/or cause severe pathology, and this is prevented by NK cells (6). NK cells are a first line of defense against viral infections through their cytotoxic activity and ability to secrete cytokines without prior activation (7, 8). Individuals with rare genetic NK cell defects are susceptible to lethal herpes virus infections early postinfection but recover when optimal T cell responses are mobilized 1–2 wk later (9). Therefore, the efficient function of both T and NK cells is required to combat infection at different phases of the immune response. Although NK cell numbers generally increase during ageing, with a shift from an undifferentiated CD56^bright to a differentiated CD56^dim phenotype (10–13), these cells have reduced cytotoxicity, as well as a decreased capacity to secrete cytokines, such as IFN-γ, MIP-1α, and IL-8 (14). This reduction in NK cell function may lead to decreased immunity, especially during the early stages of infection in older subjects (6), as well as to an increased susceptibility and greater severity of viral infections in this population.

NK cell function is determined by a balance of activating and inhibitory signals delivered by activating receptors, which recognize stress-induced ligands, and inhibitory receptors, which engage MHC class I or MHC class–like molecules on healthy cells, respectively (15, 16). Killer cell lectin-like receptor G1 (KLRG1) is a C-type lectin-like inhibitory receptor with an immune receptor tyrosine-based inhibitory motif in its cytoplasmic domain (17). It binds to the ubiquitously expressed cell adhesion molecules E-cadherin, N-cadherin, and R-cadherin (18), and binding of E-cadherin to KLRG1 prevents lysis of E-cadherin–expressing target cells (18). In humans, KLRG1 is expressed by 50–80% of NK cells and 20–40% of T cells (19). Expression of the receptor is found on mature (CD56^dim) NK cells (19, 20) and terminally differentiated T cells (18, 21, 22). Although KLRG1 expression on T cells was shown to dramatically increase with age (21–25), data on KLRG1 expression on NK cells in elderly individuals is scarce and controversial (26, 27). In a study by Hayhoe et al. (27), KLRG1 expression was found to be decreased, instead of increased, in old subjects. However, unlike in the present investigation, in which all of the samples were freshly isolated, the study by Hayhoe et al. used a mixture of fresh and frozen samples. Recent data from our group suggest that KLRG1 expression is reduced considerably during cryopreservation, which would explain the discrepancy between the data (S.M. Henson, unpublished observations). High KLRG1 expression correlates with low proliferative capacity (28, 29), impaired IFN-γ secretion (29, 30), and increased apoptosis (29) in NK cells. Moreover, in patients chronically infected with hepatitis C, blockade of KLRG1 signaling restored defective protein kinase B (Akt) phosphorylation and IFN-γ secretion (29), suggesting that KLRG1 actively contributes to functional defects observed in KLRG1⁺ cells.

AMP-responsive protein kinase (AMPK) is a protein sensor that integrates intracellular cues, such as low ATP levels and DNA damage signals, to regulate cell function (31, 32). We previously identified a novel AMPK-dependent pathway in highly differentiated CD4⁺ T cells that is activated by senescence and low nutrient sensing signals to inhibit proliferation and telomerase activity (32). Once active, AMPK triggers p38 MAPK activation, and the inhibition of either molecule restores proliferation and telomerase activity (32). Because terminally differentiated NK cells with impaired immune function accumulate during ageing (14), we hypothesized that the decline in NK cell function during ageing may also be regulated by an analogous AMPK-dependent mechanism. We found that, independently of its canonical inhibitory target Akt (22, 29), KLRG1 stimulated AMPK activity in NK cells with high KLRG1 expression (KLRG1^bright), and signaling through this pathway inhibited NK cell cytotoxicity, IFN-γ production, proliferation, and telomerase activity. Furthermore, KLRG1 was internalized upon ligation, bound to AMPK directly, and prevented its inhibitory dephosphorylation by protein phosphatase-2C (PP2C) phosphatases. This is a hitherto unrecognized mechanism of cell surface inhibitory receptor function in lymphocytes in which AMPK activation is amplified through protection from dephosphorylation, rather than de novo kinase activation. The central implication of these observations is that inhibition of KLRG1/AMPK signaling may restore NK cell function that is relevant for immune enhancement during ageing and in patients with malignancy.

After written informed consent was obtained, heparinized peripheral blood samples were taken from healthy volunteers without previous record of an autoimmune or malignant disease. Where donors are stratified by age, young is defined as ≤35 y old (age range: 20–34 y, median: 27 y, mean: 25.8 y) and old is defined as ≥70 y of age (age range: 70–86 y, median: 75 y, mean: 76.9 y).

PBMCs were isolated from heparinized peripheral blood using Ficoll-Paque (Amersham Biosciences). NK cells were isolated from PBMCs with the NK Cell Isolation Kit (Miltenyi Biotec), according to the manufacturer’s instructions. To obtain NK cell subsets with differential KLRG1 expression, NK cells were labeled with a biotinylated anti-KLRG1 Ab (BioLegend), washed, and labeled with Anti-Biotin MicroBeads (Miltenyi Biotec), according to the manufacturer’s instructions. KLRG1^bright NK cells were obtained by passing KLRG1-labeled NK cells through a magnetic column (Miltenyi Biotec) where KLRG1^bright NK cells were retained (positive fraction). Flow-through was run over two additional magnetic columns sequentially to obtain KLRG1^dim NK cells (positive fraction). All remaining cells were considered KLRG1^neg (negative fraction). Cell surface KLRG1 expression after cell isolation was determined with Streptavidin-Cy5 (BioLegend) or a directly labeled anti-KLRG1 Ab (BioLegend) (Supplemental Fig. 1B).

NK cells were cultured in complete medium (RPMI 1640 supplemented with 10% heat-inactivated FCS, 100 U/ml Penicillin, 100 mg/ml Streptomycin, and 2 mM l-glutamine; all from Invitrogen) unless otherwise stated. Glucose-free RPMI 1640 (Life Technologies) was used for glucose-deprivation experiments (6 h or overnight). MCF-10A cells were cultured in DMEM/F12 medium (Invitrogen) containing 5% horse serum (Life Technologies), 20 ng/ml epidermal growth factor (PeproTech), 0.5 mg/ml Hydrocortisone, 100 ng/ml Cholera Toxin, 10 μg/ml Insulin (all from Sigma-Aldrich), 100 U/ml Penicillin, and 100 mg/ml Streptomycin (both from Invitrogen).

The following Abs (all from BioLegend, unless otherwise indicated) were used: anti-KLRG1–biotin (2F1), anti-KLRG1–PE (2F1), anti-CD3 (UCHT1; BD Biosciences), anti-CD56 (HCD56), anti-CD16 (3G8; BD Biosciences), and anti-CD7 (M-T701). Biotin-conjugated Abs were detected using Cy5- or Cy3-conjugated Streptavidin (BioLegend). Annexin V staining was performed using Annexin V Binding Buffer and Annexin V (both from BioLegend). Ki67 staining was performed (B56; BD Biosciences) with the Foxp3 Staining Set (Miltenyi Biotec), according to the manufacturer’s instructions. When using total PBMCs, NK cells were identified as CD3⁻CD7⁺ lymphocytes (33) (Supplemental Fig. 1A). All samples were acquired on an LSR II flow cytometer (BD Biosciences). Data were analyzed using FlowJo v10 software (TreeStar, Ashland, OR).

After cell surface staining on ice, cells were fixed with Cytofix Buffer (BD Biosciences) for 10 min at 37°C and permeabilized with ice-cold Perm Buffer III (BD Biosciences), followed by staining with the following Abs for 30 min at room temperature: anti–p-AMPKα (Thr¹⁷²) rabbit mAb (40H9), anti–p-ATM (Ser¹⁹⁸¹) rabbit mAb (D6H9), anti–γH2Ax-PE (Ser¹³⁹) mouse mAb (20E3) (all from Cell Signaling Technology), and anti–p-Akt (Ser⁴⁷³) mouse mAb (M89-61; BD Biosciences). Primary Abs were detected with goat anti-rabbit, goat anti-mouse, or donkey anti-rabbit IgG (all from Life Technologies) for 30 min at room temperature. All samples were acquired immediately after the staining on an LSR II flow cytometer (BD Biosciences). Data were analyzed using FlowJo v10 software (TreeStar).

The cytotoxic capacity of NK cells was assessed using the E-cadherin–expressing breast cancer epithelial cell line MCF-10A as target. Briefly, 20,000 MCF-10A cells were plated/well of a 96-well flat-bottom plate 24 h prior to the cytotoxicity assay. On the day of the assay, MCF-10A cells were labeled with Calcein-AM (Sigma-Aldrich) at 10 μM for 1 h before coculture with NK cells in complete medium containing 500 IU/ml recombinant human (rh)IL-2 (Miltenyi Biotec) and as described (34). NK cells transfected with small interfering RNA (siRNA) specific for KLRG1 or scrambled control siRNA for 36 h were pretreated with the AMPK agonist A769662 (Tocris Bioscience) at 150 μM or equivalent DMSO as negative control for 2 h before being added to the target cells. Effector and target cells were combined at a ratio of 40:1 in triplicate and cultured in complete medium containing 500 IU/ml rhIL-2 (Miltenyi Biotec) for 4 h. After 4 h of coculture, fluorescence was measured in 75 μl of cell culture supernatant using a SpectraMax Gemini spectrofluorometer. Specific lysis was calculated as percentage killing: (test release − spontaneous release)/(max release − spontaneous release) × 100.

For quantification of granzyme B and IFN-γ in cell culture supernatants, a Cytometric Bead Array Assay (Human Granzyme B and Human IFN-γ Flex Set; BD Biosciences) was used. Measurements were performed according to the manufacturer’s instructions. The lower limit of detection was 4 pg/ml for granzyme B and 0.8 pg/ml for IFN-γ.

siRNA was introduced into freshly isolated KLRG1^bright NK cells by electroporation using the Amaxa Human NK Cell Nucleofector Kit and Nucleofector technology (Lonza), according to the manufacturer’s instructions. Briefly, 3 × 10⁶ freshly isolated KLRG1^bright NK cells were resolved in Nucleofector solution provided by the manufacturer and 200 nmol PP2C (PP2Cα siRNA; sc-42937) or 300 nmol KLRG1 siRNA (sc-42937; both from Santa Cruz Biotechnology). Nucleofection was performed with the Nucleofector I Device Program U-01 (Lonza). Cells were transferred into 12-well cell culture plates containing pre-equilibrated complete medium and incubated for 36 h. NK cells transfected with KLRG1 siRNA were cultured in complete medium containing 500 IU/ml rhIL-2 (Miltenyi Biotec). Knockdown efficiency of KLRG1 siRNA was monitored by measuring cell surface expression of KLRG1 by flow cytometry (Supplemental Fig. 3A). Knockdown of PP2Cα was confirmed by immunoblot analysis (rabbit anti-PP2Cα mAb, clone D18C10, XP; Cell Signaling Technology) (Supplemental Fig. 3B). A scrambled control siRNA (sc-37007; Santa Cruz Biotechnology) was used in all experiments.

Freshly isolated KLRG1^bright NK cells were preactivated with K562 target cells (E:T ratio of 10:1) and 200 IU rhIL-2 (Miltenyi Biotec) for 48 h. On day 2, cells were transduced with pHIV1-Siren lentiviral particles, as previously described (32). On day 6 of culture, cells were transferred to a 96-well plate coated with an anti-KLRG1 mAb or an IgG control Ab (both at 10 μg/ml; both from BioLegend), and K562 target cells (E:T ratio of 10:1) and fresh IL-2 (200 IU/ml) were added. On day 9 of culture, apoptosis was assessed by Annexin V staining, and proliferation was measured by Ki67 staining. Human telomerase reverse transcriptase (hTERT) expression was measured by intranuclear staining with the anti-TERT mAb (ab68781; Abcam). On day 9, NK cells were restimulated with K562 target cells at an E:T ratio of 10:1 and fresh IL-2 (200 IU/ml) and cultured for a total of 12 d (for experimental design see Supplemental Fig. 4A).

For coimmunoprecipitation of AMPK, PP2C, and KLRG1, cell lysate from 10 × 10⁶ isolated KLRG1^bright NK cells was prepared as previously described (32). Cell lysate was split into two and incubated with a polyclonal anti-AMPKα Ab or an isotype-control Ab (both from Cell Signaling Technology). Immunoprecipitated proteins were detected by immunoblot analysis with an anti-KLRG1 (ab170959; Abcam), anti-PP2C and anti-AMPK Ab (both from Cell Signaling Technology), and confirmation-specific mouse anti-rabbit IgG Ab (L27A9; Cell Signaling Technology), a mouse Ab to rabbit IgG L chain (L57A3; Cell Signaling Technology), or a secondary anti-mouse IgG Ab (7076; Cell Signaling), and developed with the ECL Prime Western detection kit (GE Healthcare).

AMPK immunoprecipitates were obtained as described above. After extensive washing, PP2C-like (PP2ase) activity was measured using the ProFluor Ser/Thr PPase Assay, according to the manufacturer’s instructions (Promega), and normalized against total levels of immunoprecipitated AMPK. Immunoprecipitated AMPK was detected by ELISA-based assays (ab181422; Abcam), according to the manufacturer’s instructions.

Purified KLRG1^dim and KLRG1^bright NK cells were incubated with AMPK agonist A-769662 (150 μM) or DMSO as a negative control for 12 h, followed by KLRG1 ligation or an isotype-control Ab (both from BioLegend) for 2 h at 37°C. Subsequently, cells were fixed with 2% paraformaldehyde at 37°C for 10 min and permeabilized with ice-cold Perm Buffer III (BD Biosciences) for 30 min on ice. KLRG1 was stained with a biotinylated anti-KLRG1 mAb, followed by Streptavidin-Cy5 staining (both from BioLegend). p-AMPKα was stained with an anti–p-AMPKα (Thr¹⁷²) rabbit mAb and goat anti-rabbit IgG (both from Cell Signaling Technology). Samples were run on an Amnis ImageStream cytometer using INSPIRE software (magnification ×60). Data were compensated and analyzed using IDEAS v.6.1 software (Amnis). Colocalization of KLRG1 and p-AMPK was determined on a single-cell basis using Bright Detail Similarity (BDS) score analysis. Colocalization was considered as BDS ≥ 2.0.

GraphPad Prism software was used to perform all statistical analyses. For parametric data, the Student t test or repeated-measures ANOVA test with Greenhouse–Geisser correction was used. For nonparametric data, the Wilcoxon matched-pairs signed rank test or Friedman test was used. The p values <0.05 were considered significant.

The present study was approved by the Ethical Committee of the Royal Free and University College London Medical School. Written informed consent was obtained from all study participants.

On the basis of their relative expression of CD56 and CD16, we identified three subsets of human NK cells in peripheral blood: CD56^dim (CD56⁺CD16⁺⁺, 80.5 ± 7.2%), CD56^bright (CD56⁺⁺CD16⁻, 4.6 ± 2.8%), and CD56^brightCD16⁺ (CD56⁺⁺CD16⁺, 2.39 ± 1.2%) (Fig. 1A, Supplemental Fig. 1A). When comparing relative NK cell numbers between young (≤35 y) and old (≥70 y) donors, we found, consistent with reports from other groups (13, 35, 36), that the percentage of CD56^dim NK cells was significantly increased in older individuals, whereas the percentage of the more immature CD56^bright and CD56^brightCD16⁺ NK cell subsets was reduced compared with young donors (Fig. 1B). When comparing expression of KLRG1 on NK cell subsets in young and old donors, we found that, irrespective of age, KLRG1 expression was predominant within the more differentiated CD56^dim NK cell subset (Fig. 1C). Moreover, old donors had an increase in the frequency of KLRG1-expressing NK cells compared with young donors, and this was true for the CD56^brightCD16⁺ and CD56^dim NK cell subsets (Fig. 1C). In addition to an increase in KLRG1^pos NK cells, we found a higher expression level of KLRG1 in NK cells from old donors compared with young donors (Fig. 1D). Because the inhibitory capacity of KLRG1 is known to be directly proportional to the cell surface expression level of the receptor (37), we stratified CD56^dim NK cells, the main NK cell population expressing KLRG1, into cells with no (KLRG1^neg), intermediate (KLRG1^dim), or high (KLRG1^bright) KLRG1 expression levels (Fig. 1D). Using this approach, we found a significant increase in the percentage of KLRG1^bright NK cells in old donors compared with young donors (Fig. 1E). This was most pronounced within the CD56^dim NK cell compartment, whereas KLRG1 expression was low in CD56^bright NK cells of both age groups (Fig. 1C, 1E).

We showed previously that AMPK is spontaneously active (Thr¹⁷² phosphorylation in the AMPK-activation loop) in highly differentiated/senescent T cells (32), and we questioned whether this also occurred in CD56^dim NK cells that accumulate with age. We gated on the three NK cell populations, as defined in Fig. 1A, and found that AMPK was only active in the CD56^dim NK cell subset (Fig. 2A, 2B). Because these cells also express high levels of KLRG1 (Fig. 1C–E), we investigated whether both molecules were detectable within the same cell (Fig. 2C). When we isolated NK cells on the basis of negative, dim, or bright KLRG1 expression using magnetic beads (Supplemental Fig. 1B), we found that only KLRG1^bright NK cells exhibited spontaneous AMPK activation (Fig. 2C, 2D). Functional analysis of KLRG1^bright NK cells revealed low proliferative activity (Supplemental Fig. 2A), telomere shortening (Supplemental Fig. 2B), and low expression of the catalytic component of telomerase (hTERT; Supplemental Fig. 2C), as well as endogenous phosphorylation of the DNA damage response protein H2AX (γH2Ax; Supplemental Fig. 2D) and its upstream activator, ataxia telangiectasia mutated protein (p-ATM; Supplemental Fig. 2E), compared with KLRG1^dim and KLRG1^neg NK cells. These data indicate that KLRG1^bright cells express multiple characteristics of cellular senescence (38, 39) and suggest that activation of AMPK in KLRG1^bright NK cells may be derived from cell senescence signals (32).

We next examined the effects of KLRG1 ligation on AMPK activity in purified KLRG1^neg, KLRG1^dim, and KLRG1^bright NK cells. We used phosphorylation of Akt (Ser⁴⁷³), an established KLRG1-inhibitory target (22, 29), as a readout control. The addition of an agonistic KLRG1 Ab to freshly isolated KLRG1^neg, KLRG1^dim, or KLRG1^bright NK cells robustly enhanced AMPK activity in KLRG1^bright NK cells but had no effect on the KLRG1^dim or KLRG1^neg population (Fig. 2E). In contrast, Akt phosphorylation was not altered in response to KLRG1 ligation in KLRG1^bright or KLRG1^neg NK cells but was inhibited in KLRG1^dim cells (Fig. 2F). No change in AMPK or Akt activity could be detected when an irrelevant isotype-control Ab was used (Fig. 2E, 2F). Correspondingly, there was relatively low Akt expression in freshly purified KLRG1^bright NK cells that exhibit endogenous AMPK activity (Fig. 2G versus 2C) compared with KLRG1^dim and KLRG1^neg NK cells.

To corroborate these findings, we silenced KLRG1 by siRNA in ex vivo–isolated KLRG1^bright and KLRG1^dim NK cells (Supplemental Fig. 3A). Knockdown of KLRG1 reduced AMPK phosphorylation in KLRG1^bright NK cells but had no significant effect on KLRG1^dim cells that lack endogenous AMPK phosphorylation (Fig. 2H, 2I). Together, these findings show that KLRG1 activates AMPK signaling in highly differentiated primary human NK cells. Furthermore, Akt inhibition and AMPK activation constitute two independent pathways of KLRG1 function that may operate at different stages of NK cell differentiation.

Because KLRG1 ligation only enhanced AMPK phosphorylation in NK cells with pre-existing (endogenous) AMPK activity, we reasoned that KLRG1 engagement may activate AMPK through inhibition of dephosphorylation. In ex vivo–isolated KLRG1 subsets, ligation of KLRG1 induced AMPK activation only in KLRG1^bright NK cells (Fig. 3A). When we cultured purified KLRG1^neg, KLRG1^dim, and KLRG1^bright NK cells in glucose-free medium (40), AMPK phosphorylation was detected in all three cell subsets (Fig. 3A). In glucose-starved NK cells, KLRG1 engagement with the activating KLRG1 Ab enhanced AMPK activity not only in KLRG1^bright cells but also in KLRG1^dim cells by 2–3-fold (Fig. 3A). There were no KLRG1-related changes in AMPK activation in KLRG1^neg NK cells (Fig. 3A). This indicates that KLRG1 ligation enhances pre-existing AMPK activity. Therefore, we reasoned that KLRG1 might regulate AMPK function by preventing its inhibitory dephosphorylation by upstream phosphatases rather than directly inducing de novo AMPK activation.

Dephosphorylation of AMPK by upstream phosphatases, primarily PP2C, represents an important mechanism of AMPK regulation (41, 42). We tested whether KLRG1 enhanced AMPK activity through inhibition of PP2C. Immunoblot analysis revealed comparable PP2C expression levels in all three KLRG1 subsets (data not shown). We then silenced PP2C by siRNA (Supplemental Fig. 3B) and found that KLRG1 ligation failed to enhance AMPK activity in KLRG1^bright NK cells in the absence of PP2C (Fig. 3B). Of note, PP2C knockdown did not alter KLRG1 expression or cell viability in these cells (data not shown), indicating that the unresponsiveness of PP2C-silenced NK cells to KLRG1 ligation was not due to a reduction in KLRG1 cell surface expression. Thus, KLRG1 activates AMPK through a PP2C-dependent mechanism.

To assess whether KLRG1 signaling modulates activity of PP2C-like AMPK-associated phosphatases, we ligated KLRG1 on KLRG1^bright NK cells, immunoprecipitated AMPK, and performed ELISA-based in vitro phosphatase assays. We found that KLRG1 ligation significantly reduced phosphatase activity in AMPK immunoprecipitates compared with IgG control ligation (Fig. 3C). These data show that KLRG1 signaling impedes dephosphorylation of AMPK, a process mediated by the phosphatase PP2C.

We immunoprecipitated AMPK from freshly isolated KLRG1^bright NK cells and found that KLRG1, AMPK, and PP2C interacted endogenously (Fig. 4A). Because KLRG1 signaling inhibits phosphatase activity in the AMPK complex (Fig. 3C), we reasoned that KLRG1 ligation may disrupt AMPK–PP2C interaction. When we immunoprecipitated AMPK from KLRG1^dim NK cells treated with the AMPK agonist A-769662 or DMSO vehicle control, we found that KLRG1 ligation dissociated PP2C from AMPK, a process that was most evident on an AMPK-activation background (Fig. 4B, lanes 3 and 4).

When we activated AMPK in purified KLRG1^bright NK cells by KLRG1 ligation, glucose-deprivation, or selective AMPK agonist stimulation (43), we observed a consistent reduction in cell surface KLRG1 expression (Fig. 4C). We reasoned that KLRG1 might have been internalized to facilitate interaction with AMPK. To test for internalization of KLRG1, we compared KLRG1^dim NK cells that lack endogenous AMPK activation and KLRG1^bright NK cells that spontaneously activate AMPK. To this end, we cultured isolated KLRG1^dim and KLRG1^bright NK cells overnight in the presence of the AMPK agonist A-769662 or DMSO vehicle control, followed by ligation of KLRG1 for 2 h and ImageStream analysis.

On a DMSO background, we found that KLRG1^bright, but not KLRG1^dim, NK cells showed endogenous AMPK activation and colocalization of KLRG1 and p-AMPK (Fig. 4D top panel, Fig. 4E). When KLRG1 was ligated, we found enhanced p-AMPK/KLRG1 colocalization in KLRG1^bright cells only (Fig. 4D, second panel, Fig. 4E). In contrast, AMPK was refractory to triggering of the receptor per se in KLRG1^dim NK cells (Fig. 4D second panel, Fig. 4E). Because pre-existing AMPK activity is essential for KLRG1 regulation of AMPK (see above), we reasoned that enforcing AMPK activation in KLRG1^dim cells would reconstitute p-AMPK/KLRG1 colocalization, even in this less-differentiated population. On an AMPK-activation background, we indeed observed KLRG1 and p-AMPK colocalization in KLRG1^bright NK cells, as well as in KLRG1^dim NK cells (Fig. 4D third panel, Fig. 4E). KLRG1 ligation now enhanced p-AMPK/KLRG1 colocalization further in both cell types (Fig. 4D fourth panel, Fig. 4E). Quantification of colocalization is shown in Fig. 4E for all four conditions, and quantification of KLRG1 and p-AMPK signals is shown in Supplemental Fig. 3C. We obtained comparable results when probing colocalization of KLRG1 and total AMPK by ImageStream (data not shown). Together, these data support a model whereby KLRG1 internalizes upon ligation, an event that requires pre-existing AMPK activation, and further enhances AMPK signaling.

We next investigated the functional implications of inhibiting NK cells via the KLRG1/AMPK axis. We probed KLRG1 function by coculturing KLRG1^bright NK cells with the breast cancer cell line MCF-10A, which expresses abundant levels of the KLRG1 ligand E-cadherin (44). Knockdown of KLRG1 by siRNA increased killing capacity (measured by direct tumor cell lysis), as well as release of granzyme B and IFN-γ secretion of KLRG1^bright NK cells, toward the MCF-10A cell line (Fig. 5A). Importantly, this process was AMPK dependent, because all three functions examined were abolished by addition of the AMPK agonist A-769662 to the KLRG1-silenced NK cells (Fig. 5A). These data identify an inhibitory pathway of KLRG1 function in primary human NK cells that requires AMPK signaling.

To further dissect the link between AMPK and KLRG1 function, we silenced AMPK in primary human NK cells, followed by KLRG1 ligation. We activated purified KLRG1^bright NK cells with the MHC class I–deficient NK target cell line K562 and rhIL-2 for 48 h and then transduced cells with lentiviral vectors encoding short hairpin RNA to AMPK (shAMPK) or an irrelevant scrambled short hairpin RNA (control RNA; shCtrl). Four days posttransduction, we transferred NK cells to plates coated with KLRG1 or an isotype-control Ab and reactivated them up to 6 d, as above (for experimental design and transfection efficiency see Supplemental Fig. 4). The knockdown of AMPK restored NK cell expansion (Fig. 5B), proliferation (Fig. 5C), and hTERT expression (Fig. 5D, 5E) but reduced DNA damage (Fig. 5F) and apoptosis (Fig. 5G) compared with KLRG1^bright NK cells transduced with shCtrl. For each function examined, the inhibitory effect induced by KLRG1 ligation was robust in NK cells transduced with shCtrl but was totally abolished in AMPK-silenced cells (Fig. 5B–G). Of note, AMPK knockdown did not alter KLRG1 cell surface expression in NK cells (data not shown). Thus, AMPK activation is a new inhibitory pathway of KLRG1 function in primary human NK cells, and blocking this pathway restores NK cell responsiveness.

We identified a previously unrecognized function of the inhibitory receptor KLRG1 in linking cell surface inhibitory receptor signaling to AMPK activation in human NK cells. NK cells integrate signals from inhibitory and activating receptors to detect and eliminate infected and transformed cells (45, 46). Although activating NK cell receptors recognize stress-induced ligands on target cells, inhibitory NK cell receptors recognize MHC class I and MHC class I–like molecules on healthy cells (8, 46) as a means of protecting healthy tissue from NK cell attack. It is conceivable that an imbalance toward inhibitory immune-receptor signaling, as we now demonstrate for KLRG1 in individuals >70 y, increases the activation threshold of NK cells and leads to attenuation of effector function. Although this mechanism potentially protects from autoreactivity and collateral damage during infection, it might render hosts more susceptible to infection and/or malignancy by attenuating NK cell function. Therefore, we suggest that blocking of the KLRG1/AMPK signaling axis may boost NK cell activity in the older population.

Our data support a model whereby persistent (endogenous) AMPK activity is regulated through protection from dephosphorylation rather than by upstream kinase activation. The phosphatase PP2C is known to dephosphorylate (and thus inactivate) AMPK in vitro and in vivo (41, 42), but the precise physiological mechanism by which PP2C itself is regulated in humans was not known. We now show that KLRG1 internalizes upon ligation, binds directly to AMPK, and stimulates its function by inhibiting PP2C-like phosphatase activity. Correspondingly, KLRG1 ligation strongly diminished PP2C binding to AMPK in human NK cells. This links AMPK activation with surface inhibitory receptor signaling, suggesting that immune-inhibitory receptors, which increase with age, chronic viral infections, and cancer (14, 47, 48), may orchestrate AMPK-dependent metabolic pathways to inhibit NK cell function.

We further demonstrate that KLRG1 requires pre-existing AMPK activity to stimulate the AMPK pathway. Thus, KLRG1 seems to function as a natural AMPK enhancer in NK cells that is incapable of triggering de novo kinase activation but induces potent signal amplification. This occurs endogenously in highly differentiated NK cells that exhibit senescence characteristics, such as pre-existing DNA damage and spontaneous AMPK activity (32). We showed recently that endogenous DDR signaling activates AMPK in senescent T cells, and we now extend this observation to highly differentiated human NK cells. This raises the paradox of why constitutive activation of a low-energy sensor molecule would require continuous energy consumption to fuel its own activation in senescent cells. Because KLRG1 impedes AMPK dephosphorylation, we propose a model in which endogenous AMPK activity is maintained by active surface inhibitory receptor signaling rather than continuous de novo upstream activation. Beyond biochemical considerations, the central implication for this is that endogenous AMPK activity may be controlled by means of surface inhibitory receptors in human NK cells, a mechanism that was previously unknown.

It remains to be determined whether KLRG1 has a similar role in highly differentiated human T cells that also express high levels of KLRG1 (19, 49) and spontaneously activate AMPK (32). Furthermore, it is not known whether this pathway is shared with other T cell inhibitory receptors, such as CTLA-4 and PD-1, for which blocking Abs have been approved or are in clinical development.

In summary, we demonstrate that there is an intimate relationship between senescence and energy-sensing pathways in NK cells that can be further modulated by inhibitory receptor engagement. Intervention in this signaling axis may enhance NK cell activity.

The authors have no financial conflicts of interest.