An IL-2 Paradox: Blocking CD25 on T Cells Induces IL-2–Driven Activation of CD56^bright NK Cells

Natural killer cells are an important element of the innate immune system and act against virally infected cells and tumor cells through a complex set of activating and inhibitory signals, the processing of which may lead to cytotoxicity or release of cytokines (1). In humans, there are two main classes of NK cells: CD56^dim and CD56^bright NK cells. CD56^dim NK cells, which constitute the majority (∼90%) of NK cells in peripheral blood, produce large amounts of perforin and mediate robust cytotoxicity toward MHC class I-deficient targets. Alternatively, CD56^bright NK cells, representing only a small fraction (∼10%) of NK cells in the blood, are found in higher proportions in the lymph nodes and inflammatory lesions. They are thought to have immunoregulatory properties, primarily through their rapid production of cytokines after activation (2). Because of their lower perforin and granzyme B production, CD56^bright NK cells were originally considered not to be cytotoxic. However, recent data indicate that they constitutively express granzymes A and K (3) and in addition to killing MHC class I-deficient targets, they also kill very different targets than those killed by CD56^dim NK cells, namely, autologous MHC class I-expressing immature dendritic cells (iDCs) (4) and activated T cells (5).

Multiple sclerosis (MS) is a debilitating immune-mediated disease of the CNS. Although the cause of the disease is not known, it is believed that the aberrant activation of the adaptive immune system underlies focal CNS tissue destruction visible by magnetic resonance imaging as MS lesions. Dysfunctional NK cells were described in relation to MS as early as 1980 when Benczur et al. (6) found decreased NK cytotoxicity toward MHC class I-deficient targets in MS patients. More recently, a correlation was found between periods of low NK cell functional activity and MS disease activity identified as new lesions on magneic resonance imaging scans in a time-course study of patients with relapsing-remitting MS (7). In the mouse model of MS, experimental autoimmune encephalomyelitis, NK cell-depleting Abs administered to mice prior to immunization with myelin oligodendrocyte glycoprotein_35–55 caused earlier onset and increased severity of experimental autoimmune encephalomyelitis (8). These, and many other human and animal studies (reviewed in Ref. 9), suggest that NK cells may play an important role in limiting neuroinflammation and perhaps autoimmunity in general (10).

CD25, the IL-2R α-chain, has been genetically linked to MS disease susceptibility (11). When CD25 associates with the intermediate-affinity IL-2R (K_d = 1 nM), composed of a β-chain (CD122), and the common γ-chain (γ_c; CD132), it increases the affinity of the signaling complex for IL-2 10- to 100-fold; thus forming the high-affinity IL-2R (K_d = 10 pM) (12). Because the high-affinity IL-2R is expressed on T cells upon activation and is believed to facilitate T cell entry into the proliferation cycle, CD25 blocking Abs (e.g., daclizumab [Dac] and basiliximab) were developed as immunomodulatory agents that could selectively inhibit activated T cells (13). Indeed, these agents inhibit the rejection of solid organ transplantation and Dac also limits immune-mediated pathology associated with inflammatory uveitis (14) and MS (15–18). We have previously reported that the inhibitory effect of Dac on MS disease activity is linked to the expansion of CD56^bright NK cells in treated patients (5). We observed that CD56^bright NK cells expanded during Dac therapy exhibited higher expression of IL-2R β-chain and proliferated more vigorously in response to IL-2 and IL-15 in vitro than their counterparts isolated before Dac administration. In addition, the expanded CD56^bright NK cells had higher surface expression of IL-2 regulated genes, such as CD44, CD127 (IL-7Rα), and KIR2DL4. Therefore, we hypothesized that Dac expands CD56^bright NK cells in an IL-2–dependent manner; by enhancing the availability of IL-2 for NK cell signaling via the intermediate-affinity IL-2R due to decreased consumption of IL-2 by activated T cells.

However, the evidence for this hypothesis was only indirect. Furthermore, a similar hypothesis for the IL-2–driven proliferation of CD56^bright NK cells was tested previously during therapeutic administration of IL-2 to cancer patients and was believed to be incorrect (19). Specifically, based on cell cycle analysis, Fehniger et al. (19) concluded that the robust expansion of CD56^bright NK cells in IL-2–treated cancer patients previously treated with chemotherapy resulted from enhanced NK cell differentiation from bone marrow progenitors combined with an IL-2–induced delay in NK cell death, rather than from NK cell proliferation. Because of the apparent importance of CD56^bright NK cells in regulation of neuroinflammation and the genetic link of CD25 to MS, we sought to characterize the mechanism of action of Dac in MS, including the mechanism of expansion of CD56^bright NK cells, in greater detail. To this end, we investigated all components of our hypothesis directly by analyzing matched cryopreserved samples collected from 25 MS patients who participated in two previously published clinical trials of Dac in MS (17, 20). First, we observed the expansion of CD56^bright NK cells under Dac therapy and examined their in vivo proliferation by quantification of Ki67 staining, then we searched for direct ex vivo evidence that Dac inhibits IL-2 signaling in T cells by quantifying the numbers of FoxP3⁺CD4⁺ T regulatory cells (Tregs), which are known to be dependent on IL-2 signal for their survival (21). We then directly analyzed STAT5 phosphorylation by T cells and NK cells in response to exogenous IL-2, measured NK cell functional activity directly ex vivo by using a flow cytometry based killing assay of GFP-tagged MHC class I-deficient target cells combined with CD107a staining that identifies degranulating effector cells, and finally, we performed in vitro functional studies, including transwell experiments and an IL-2 consumption assay to further support our conclusions.

To conduct these experiments, we used cryopreserved PBMC samples isolated from lymphocytapheresis taken from patients in two separate trials: one as described in (17) where patients, who were unresponsive to IFN-β therapy at baseline, were treated with Dac and IFN-β combination therapy and taken off IFN-β at late time points, and another trial where Dac was used as monotherapy (20). No significant differences were found between the two cohorts unless otherwise noted.

Cryopreserved PBMCs were thawed and 1 × 10⁶ cells were stained for FoxP3 and Ki67 (FoxP3 staining buffer set; eBioscience; San Diego, CA). Samples were acquired by flow cytometry (LSR II, with HTS; BD Biosciences; San Jose, CA) and analyzed with FACS Diva software (BD Biosciences). Abs (and clones) used include: Ki-67 (B56), CD127 (HIL-7R-M21), CD56 (B159), CD8 (RPA-T8), CD4 (SK3), CD3 (SK7; all BD Biosciences), FoxP3 (236A/E7) and CD25 (BC96; eBioscience). Staining was performed in duplicates and gating was set on appropriate isotype controls.

Cryopreserved PBMCs were thawed and plated in X-vivo media (Lonza, Walkersville, MD) at 1 × 10⁵ cells/well in a 96-well plate. They were rested for 1 h at 37°C and pulsed with 50–100 IU/ml of IL-2 (National Cancer Institute Biological Resources Branch Preclinical Repository; Frederick, MD; all patients were tested with 100 IU/ml, a subgroup was tested also with 50 IU/ml) or 10 ng/ml IL-7 (PeproTech, Rocky Hill, NJ) for 10 min; following which, they were fixed, permeabilized (Cytofix buffer and PhosFlow Perm buffer II, BD Biosciences), and stained with Abs acceptable for phospho-specific staining. Abs used include CD56 (MY31), CD3 (UCHT1), phosphorylated STAT5 (pSTAT5; 47, pY694-STAT5; all BD Biosciences), and CD4 (RPA-T4; eBioscience). Previous studies have determined the validity of the PhosFlow signaling methodology applied to cryopreserved PBMCs (22).

Cryopreserved PBMCs were thawed and NK cells were depleted by CD56 iMag microbeads (BD Biosciences). Alternatively, T cells were isolated from PBMCs from lymphocytaphereses by negative selection (T cell isolation kit II, Miltenyi Biotec, Auburn, CA). A total of 1 × 10⁶/ml T cells or NK-depleted PBMCs were plated in IMDM media (Lonza) containing 10% human serum and either M-A251 control Ab against CD25 that does not block the IL-2–binding tac epitope (BD Biosciences) or Dac (Zenapax, Hoffman-La Roche, Nutley, NJ; 10 μg/ml each). After incubating with the Abs at 37°C for 1 h, T cells were polyclonally stimulated with CD3/CD28 Dynabeads (Invitrogen; Carlsbad, CA) at a 0.3:1 bead/cell ratio. After IL-2 production tapered off (72 h later), cells were washed thoroughly, recounted, and reseeded at 1 × 10⁶/ml in 10% human serum media with 20 IU/ml IL-2, maintaining original Ab concentrations. After 48 h, supernatants were collected and measured for IL-2 by ELISA (IL-2 Ready-Set-Go ELISA kit, eBioscience).

Cryopreserved PBMCs from MS patients that have continued on long-term Dac therapy were thawed and NK cells and T cells were isolated by negative selection (MACS Human NK Cell Isolation Kit and MACS Pan T cell Isolation Kit II, Miltenyi Biotec). These populations were determined to be of 89.1 and 96.5% purity, respectively, by FACS. T cells and NK cells were rested overnight in IMDM media (Lonza) with 10% AB human serum (Gemini Bio-Products, West Sacramento, CA). The NK cells were then CFSE stained (Molecular Probes, Invitrogen) as described previously (5) and cocultured with T cells. The T cells were seeded at 5 × 10⁴ cells/condition (200 μl 10% serum in IMDM) and preincubated with Dac for 1 h to prevent T cell consumption of IL-2, washed twice, and stimulated with CD3/CD28 Dynabeads (Invitrogen) before coculture with increasing numbers (1 × 10⁴, 2.5 × 10⁴, or 5 × 10⁴) of NK cells. In transwell experiments, PBMCs were CFSE stained and then NK cells were separated/depleted by CD56 microbeads (Miltenyi Biotec). The NK-depleted PBMCs were polyclonally activated with plate-bound anti-CD3 (20 ng/ml) and anti-CD28 (10 μg/ml) Abs (PeproTech) and the NK cells were added into transwells (3-μm pore size, Polycarbonated Membrane; Corning Costar, Lowell, MA) at a 1:10 NK/T cell ratio. Alternatively, the transwell experiment was repeated with purified, negatively selected T cells and NK cells. T cells were polyclonally activated for 24–72 h, extensively washed, and reseeded to the lower compartment of transwells at 1 × 10⁶ activated T cells in media containing different blocking agents as indicated. Autologous NK cells were isolated from fresh or cryopreserved apheresis samples at the day of coculture with previously activated T cells. To differentiate between NK cells added to the upper compartment and activated T cells added to the lower compartment of transwell, only purified NK cells were CFSE stained before coculture. Proliferation assays were analyzed by FACS at day 3–5 for CFSE dilution. Absolute numbers of proliferating cells were evaluated as a ratio of CFSE-diluted NK cells to FITC beads that were added in equal number to each well before acquisition.

To fluorescently tag K562 MHC class I-deficient tumor cells, they were transduced with GFP. Briefly, pMSGV1-eGFP, a retroviral vector carrying the GFP gene, was cotransfected with pRD114, providing env for efficient packaging into the packaging cell line, 293-GP. Pseudotype virus expressing eGFP was harvested and used for K562 transduction.

To make the GFP viral supernatant, 293GP cells (Clontech; Mountain View, CA) were seeded at 6 × 10⁶ cells/well in a 10-cm plate coated with Poly-D-Lysine (BD Biosciences). A total of 9 μg pMSGV1-GFP and 4.5 μg pRD114 were transfected into 293-GP using Lipofectamine 2000 (Invitrogen) according to the manufacturer’s protocol. At 24–48 h posttransfection, supernatants were collected and spun at 1000 × g, at room temperature for 10 min to remove cell debris. For the transduction of the GFP gene into K562 cells, 4 × 10⁶ K562 cells/well, prewashed with PBS, were mixed with 3 ml viral supernatant, 1 ml fresh media, 10 μg/ml protamine sulfate (APP Pharmaceuticals, Schaumburg, IL), and plated in a 6-well plate. The plate was centrifuged at 1000 × g, at 32°C for 2 h and then cultured in a 37°C, 5% CO₂ incubator. GFP expression was monitored using fluorescence microscopy and FACS.

Single K562 GFP clones were generated after a 0.3 cell/well limiting dilution of the eGFP transduced K562 cell mixture. Cells in eGFP positive wells were expanded and the purity of the clone and intensity of eGFP expression were confirmed by FACS analysis.

NK cell cytotoxicity was measured in a similar fashion to a protocol previously described (23). Cryopreserved PBMCs were thawed and plated at 1 × 10⁶ cells/well and rested for 30 min at 37°C before adding 1 × 10⁵ GFP-transduced MHC class I-deficient K562 target cells and CD107a Ab (H4A3; BD Biosciences). Control wells included effectors (NK cells only with CD107a Ab) without targets and targets (K562 cells only) without effectors. Cultures were incubated overnight, and subsequently stained for surface expression of CD3 (UCHT1), CD8 (RPA-T8), and CD56 (MY31; BD Biosciences). They were acquired by flow cytometry, additionally assessing for the presence of GFP and CD107a. Live GFP⁺ cells were gated on forward and side scatter and gating for CD107a (i.e., degranulation) was set from control wells of effectors without targets. Cells were proportionally enumerated between different conditions using a reference number of fluorescent beads that were added in equal numbers to all conditions.

Fresh PBMCs from normal donors were isolated by Ficoll separation (Lymphocyte Separation Medium; Lonza) and NK cells were isolated by negative selection (MACS Human NK cell Isolation Kit; Miltenyi). NK cells were then incubated overnight at 37°C and 5% CO₂ in IMDM media (Lonza) containing 10% human serum (Gemini Bio-Products) with either no additional cytokine, 10 or 100 IU/ml IL-2, or 2 or 20 ng/ml IL-15 (PeproTech). They were then washed thoroughly and some aliquots were taken for intracellular staining of perforin as described (5). After washing, NK cells were seeded in X-vivo media at 5 × 10⁴ cells with 5 × 10⁴ GFP⁺ K562 cells with 4 μl CD107a PE-Cy5 Ab per pretreatment condition. Control wells were identical to those used in the ex vivo cytotoxicity assay described previously. After 4 h, the coculture assay was stained for CD107a incorporation and intracellular perforin and analyzed for death of GFP⁺ K562 cells. Abs used included CD56 (B159), CD3 (UCHT1), perforin (δG9), granzyme A (CB9), and CD107a (H4A3) (all BD Biosciences). Gating for perforin was set on the appropriate isotype control and gating for CD107a (i.e., degranulation) was set from control wells of effectors without targets. Perforin expression in cultures with targets was measured from CD107a⁺CD56^bright NK cells that had released their granules in culture. Absolute numbers were again acquired using fluorescent beads for normalization.

To compare therapy samples to baseline samples and in vitro proliferation data, we analyzed our data using one-way RM ANOVA unless otherwise indicated. We used the Friedman one-way RM ANOVA on Ranks to analyze in vitro cytotoxicity data using increasing cytokine levels and paired t test for the IL-2 consumption assay. All differences listed in the text are changes in the median values. Values of p are as follows in figures: *p < 0.05; **p < 0.01; ***p < 0.001. Statistical analyses were performed with SigmaStat software version 3.5.

As previously reported (5, 17), CD56^bright NK cells expand on Dac therapy. Our results again illustrate the significant in vivo expansion of CD56^bright NK cells that continues between early and late therapy time points (B→T1: 115.6%; p < 0.01 and T1→T2: 78.9%; p < 0.001) (Fig. 1A). This expansion was at least partially driven by enhanced in vivo proliferation of CD56^bright NK cells as demonstrated by an early increase in cells positive for Ki67, a chromosomal marker of cells undergoing mitosis (B→T1: 34.1%; p < 0.01). However, the proliferation of CD56^bright NK cells plateaued as CD56^bright NK cells expanded, as the proportion of proliferating CD56^bright NK cells decreased slightly between T1 and T2 (T1→T2: −8.8%) (Fig. 1B). Nevertheless, the absolute numbers of proliferating CD56^bright NK cells remained significantly higher than at baseline during the entire Dac treatment (B→T1: +303.1%; B→T2: +591.5%; p < 0.05) (Fig. 1B, right panel).

In this case, we observed differences between the patients in the Dac monotherapy trial versus those who took Dac as an add-on therapy to IFN-β during the first 6 mo of treatment (i.e., combination therapy trial) (see 1Materials and Methods for a detailed explanation of the differences between the two trials). At baseline, patients from the combination therapy trial had significantly higher numbers of CD56^bright NK cells (55.4%, p < 0.05, Mann-Whitney rank sum test) than patients from the monotherapy trial. Accordingly, the decrease in the proportion of Ki67⁺ proliferating CD56^bright NK cells from T1 to T2 was almost entirely driven by patients from the combination therapy trial (−46.0% in combination therapy patients versus −5.7% in monotherapy patients; p < 0.05; t test). Dac treatment did not affect the number of CD56^dim NK cells (−6.6% from B to T2; ns).

We hypothesized that the observed expansion and enhanced proliferation of CD56^bright NK cells was due to higher in vivo availability of IL-2, because Dac inhibited IL-2 consumption by T cells. However, in vivo IL-2 production generally occurs at sites of immune activation, such as in lymphoid organs or inflamed tissues, which are not amenable to ex vivo testing. Therefore, we used FoxP3⁺ Tregs as a surrogate marker for in vivo IL-2 consumption by T cells, because FoxP3⁺ Tregs are known to constitutively express CD25 and are dependent on IL-2 for their proliferation and survival. Intranuclear staining for transcription factor FoxP3 (example of raw data in Supplemental Fig. 1) revealed that there was a statistically significant decrease in Treg populations induced by Dac therapy (B→T1: −39.1%; B→T2: −40.4%; p < 0.001) (Fig. 2A). Staining for Ki67 identified a proportion of proliferating Tregs, which decreased between B and T1 (B→T1: −24%; p < 0.05), but apparently returned to baseline at T2 (B→T2: −1.5%; ns) (Fig. 2B). However, when analyzing the number of proliferating Tregs per 1000 lymphocytes (Fig. 2C), it became clear that the absolute numbers of proliferating Tregs remained significantly below baseline values for both T1 and T2 (B→T1: −62.2%; B→T2: −58.4%; p < 0.001) (Fig. 2C).

The establishment of a new balance between absolute numbers of Tregs and their in vivo cycling indicated some type of functional adaptation of Tregs to the lack of high-affinity IL-2 signaling induced by the blockade of CD25 by Dac (Fig. 2D). Because Tregs can use other γ_c-signaling cytokines (especially IL-7) for their development and survival (24), we investigated the changes in IL-7Rα expression (CD127) on Tregs. We observed a progressive increase in the level of expression of CD127 induced by Dac treatment (B→T1: +16%; p < 0.01 and B→T2: +32.9%; p < 0.01) (Fig. 2E). The increased expression of CD127 in FoxP3⁺CD4⁺ T cells may be seen in recently activated T cells, which lack suppressive function (25). However, it is unlikely that our gating included activated T cells, because the level of CD127 expression on the FoxP3⁺CD4⁺ T cells remained significantly below the levels of CD127 expression on FoxP3⁻CD4⁺ T cells (mean fluorescence intensity [MFI] 198.5 versus 803.8; p < 0.001; rank sum test) during the entire treatment period. Because another in vivo marker of the lack of IL-2 signaling on Tregs is their decreased expression of FoxP3 (24), we analyzed the level of FoxP3 expression on Tregs under Dac therapy (Fig. 2F) and observed that Dac induced a sustained decrease in FoxP3 MFI (B→T1: −12.6%; B→T2: −14.5%; p < 0.001).

To examine how Dac therapy altered IL-2 signaling, we analyzed the phosphorylation of STAT5 on T cell and NK cell subsets in PBMCs by flow cytometry in response to exogenously added IL-2 (50–100 IU/ml).

IL-2-induced STAT5 phosphorylation in T cells (CD4⁺ T cells are depicted as a representative T cell subset) (Fig. 3) was significantly inhibited by Dac therapy in comparison with baseline samples (B→T1: −40.2%; p < 0.001, B→T2: −28%; p < 0.001) (Fig. 3A). We observed no changes in total STAT5 protein induced by Dac therapy (Supplemental Fig 2). There was additionally an early decrease by CD8⁺ T cells in STAT5 phosphorylation to IL-2 (B→T1: −36.8%; p < 0.001; data not shown), but phosphorylation rebounded at the later time point (B→T2: 5.3%; p < 0.05) (data not shown). If cells were further incubated in vitro with Dac (10 μg/ml) prior to IL-2 pulsing, signaling was completely abrogated at all time points (data not shown). As a signaling control we selected IL-7, because it also phosphorylates STAT5 and because we observed an upregulation of CD127 (the α-chain of the IL-7R) on FoxP3⁺ Tregs. Surprisingly, we observed an increase in signaling to IL-7 by CD4⁺ T cells (B→T2: 35.6%; p < 0.05) (Fig. 3B) and CD8⁺ T cells (B→T2: 36.7%; p < 0.001) (data not shown).

In contrast to the effect of Dac therapy on IL-2 signaling on T cells, we observed no inhibition of IL-2 signaling on CD56^bright NK cells. In fact, there was an upward, nonsignificant trend in the proportion of CD56^bright NK cells that phosphorylated STAT5 in response to IL-2 throughout the treatment course (Fig 3C). Due to Dac-induced expansion of CD56^bright NK cells, this resulted in a highly significant increase in the absolute numbers of IL-2–induced pSTAT5⁺ CD56^bright NK cells during Dac therapy (B→T1: +165.7%, B→T2: +407.0%; p < 0.001) (Fig 3C, right panel). We observed no significant changes in IL-7–induced STAT5 phosphorylation on CD56^bright NK cells (data not shown).

So far our data indicate that Dac therapy inhibits IL-2–driven STAT5 phosphorylation by T cells, but does not affect IL-2 signaling on NK cells and that CD56^bright NK cells proliferate more vigorously in vivo after initiation of Dac treatment. Activated T cells are the main producers of IL-2 in vivo and we have previously reported (5) that Dac treatment does not inhibit IL-2 secretion by activated T cells. Therefore, we wanted to examine if Dac limits the consumption of IL-2 by T cells and as a consequence, if the resultant excess of T cell-derived IL-2 can drive proliferation of CD56^bright NK cells.

To this end, we performed an IL-2 consumption assay (Fig. 4A) where T cells were polyclonally activated for 3 d, then extensively washed and reseeded in the presence of Dac or M-A251 control anti-CD25 Ab (10 μg/ml each) and exogenously added IL-2 (20 IU/ml). Because the IL-2 production by T cells lasts 24–48 h poststimulation, and the CD25 expression peaks 72 h poststimulation (data not shown), the 72-h time point was selected as optimal to measure IL-2 consumption. As demonstrated in Fig. 4A, activated T cells treated with M-A251 had almost completely consumed all exogenously added IL-2 (90.7%; p < 0.05; paired t test) in 24 h. However, Dac-treated activated T cells had consumed only 17% (ns; paired t test) of exogenously added IL-2.

Next, we evaluated the proliferation of CD56^bright NK cells in coculture with increasing numbers of T cells polyclonally activated in the presence of Dac (Fig. 4B). We observed that CD56^bright NK cells in culture with larger numbers of T cells proliferated more vigorously compared with CD56^bright NK cells cultured with fewer T cells (1:1→2.5:1: 66%; p < 0.001, 2.5:1→5:1: 31.4%; p < 0.001and 1:1→5:1: 116.6%; p < 0.001).

To investigate whether T cell to NK cell contact is necessary for the observed enhanced proliferation, we used transwells to separate CFSE-stained NK cells from polyclonally activated T cells (Fig. 4C, 4D). We observed significantly enhanced survival and proliferation of CD56^bright NK cells in transwell inserts if T cells were activated in the presence of Dac in comparison with control Ab M-A251 (Fig. 4C; compare first and second panels). Furthermore, proliferation of CD56^bright NK cells was almost completely abrogated in the presence of IL-2 neutralizing Ab (Fig. 4C, 4D). Addition of a low dose of IL-2 (20 IU/ml) enhanced proliferation of CD56^bright NK cells above the M-A251 condition, but did not reach levels observed in the Dac condition. We also observed that activated NK cells were able to migrate through 3-μm pores of the transwell to the lower compartment, likely due to a strong chemotactic gradient produced by activated T cells. Our setup purity checks demonstrated >95% NK cell purity in the upper compartment and >95% T cell purity in the lower compartment (Supplemental Fig. 3) at the beginning of transwell coculture. Yet, consistently in every experiment we observed transmigration of activated NK cells through the 3 μm semipermeable membrane and, in fact, significant enrichment of proliferating CD56^bright NK cells in the lower compartment. Therefore, we provide both raw and analyzed data depicting CFSE-stained NK cells and CFSE-negative T cells in both compartments.

Finally, to confirm that activated T cells were driving the proliferation of CD56^bright NK cells through soluble IL-2, we performed supernatant transfer experiments, where we collected supernatants from activated T cells cultured for 24–48 h in the presence of M-A251 control Ab, Dac, or IL-2–blocking Ab. We isolated NK cells by negative selection from apheresis samples; CFSE stained them and cultured them in supernatants collected from activated T cells for 3 d (Fig. 4E, 4F). Again, we observed that supernatants from activated T cells cultured in the presence of Dac-induced proliferation of CD56^bright NK cells that was at least 2- to 3-fold higher than proliferation of CD56^bright NK cells cultured with the media from M-A251 cultures. Blockade of IL-2 resulted in complete abrogation of survival and proliferation of CD56^bright NK cells.

In addition to describing the expansion of the CD56^bright cell population, we sought to define functional changes of CD56^bright NK cells induced by Dac therapy. To do this, we used a flow cytometric killing assay with GFP transduced MHC class I-deficient cancer cell line, K562, as a target cell and used the detection of transient surface expression of LAMP protein CD107a, normally expressed only intracellularly in cytotoxic granules, as identification of effector cells in complex PBMC cultures. Although both CD56^dim and CD56^bright NK cells (but not CD8⁺ and CD4⁺ T cells) are cytotoxic in this assay, the proportion of CD56^dim NK cells in PBMC cultures and their activation status was not affected by Dac therapy (5); therefore, we expected that the observed changes could be directly attributed to changes in CD56^bright NK cell subsets. We confirmed the validity of this assumption by evaluating changes in the degranulation of both NK cell subsets.

In baseline samples, we observed on average 14.5% killing of K562 cells (Fig. 5A). This baseline cytotoxicity increased by 139.4% (p < 0.01) to 34.7% killing at T1 and further by 55.1% (p < 0.001) between T1 and T2 (T2; 53.9% killing). This enhanced cytotoxicity was paralleled by a significant increase in the proportion of degranulating CD56^bright NK cells as visualized by CD107a incorporation (B→T1 + 44.9%; p < 0.001, B→T2 + 50.4%; p = 0.001) (Fig. 5B). Analysis of the absolute numbers of degranulating NK cell subsets demonstrated a highly significant increase in degranulating CD56^bright NK cells (B→T1 +207.9%; p < 0.001, B→T2 +530.2%; p < 0.001) (Fig. 5C, left panel) which correlated with the observed enhanced killing (R_Spearman = 0.38, p = 0.001) (Fig. 5D). No significant change in the numbers of degranulating CD56^dim NK cells was observed (Fig. 5C, right panel).

Although our data point to IL-2 as a central factor in the expansion and activation of CD56^bright NK cells observed during Dac therapy, they do not rule out contributions of other factors that may be important in vivo, such as IL-15. Therefore, we next investigated whether IL-2 or IL-15 could induce changes in the function of CD56^bright NK cells in vitro analogous to those observed after Dac therapy in vivo. Fresh NK cells were isolated by negative selection from healthy donor apheresis. These NK cells were stimulated overnight with varying amounts of IL-2 and IL-15, and subsequently examined for their ability to kill MHC class I-negative targets. Intracellular cytokine staining after overnight activation revealed that all CD56^bright NK cells expressed perforin, but increasing concentrations of IL-2 (Fig. 6A) and IL-15 (Fig. 6D) induced higher perforin expression (MFI) in NK cells that were cultured without targets. IL-2 was more effective in this regard, especially at lower concentrations (10 IU/ml IL-2 = 6.1 ng/ml increased perforin MFI by 61.3%; p = 0.001, in comparison with 2 ng/ml IL-15, which increased perforin MFI by 26.9%; p < 0.05). When NK cells were cocultured with targets, they degranulated vigorously (Fig. 6B, 6E), losing their intracellularly preformed perforin (Fig. 6A, 6D; NK cells + target) as they killed GFP-tagged K562 cells (Fig. 6C, 6F). Although both degranulation and K562 killing was again enhanced by IL-2 and IL-15; IL-2, especially at low doses, was significantly more effective in functional activation of CD56^bright NK cells. Low-dose IL-2 increased degranulation of CD56^bright NK cells by 111.63% (Fig. 6B) in comparison with 35.94% induced by low-dose IL-15 (Fig. 6D). Similarly, killing efficiency was enhanced by 32.8% (p < 0.05; Fig. 6C) by low-dose IL-2 and only by 2.36% (ns; Fig. 6F) by low-dose IL-15. The loss of perforin MFI in CD56^bright NK cells on coculture with targets (Δ perforin MFI ^−/+ target) was significantly greater when NK cells were pretreated with IL-2 (+184.9% with IL-2 10 IU/ml and +209.3% with IL-2 100 IU/ml; p < 0.05 for both) (Fig. 6A, right panel) as compared with IL-15 (+134.9% with IL-15 2 ng/ml and +136.9% with IL-15 20 ng/ml, both ns) (Fig. 6D, right panel). This cytokine-enhanced loss of perforin by CD56^bright NK cells on their coculture with targets correlated strongly with enhanced efficacy of K562 killing (R_Spearman = 0.718; p = 0.00141) (Supplemental Fig 4).

In this study, we present evidence that Dac increases the in vivo proliferation of CD56^bright NK cells at the same time as it limits numbers and proliferation of IL-2–dependent FoxP3⁺ Tregs. Together with this increase in numbers, Dac therapy also enhances the ability of CD56^bright NK cells to kill MHC class I-deficient targets, a function that can be mimicked in vitro by physiologically achievable doses of IL-2, but much less effectively by IL-15. Presented in vitro models indicate that Dac inhibits the consumption of IL-2 by activated T cells, leading to greater availability of this cytokine for signaling by CD56^bright NK cells, which is paradoxically not inhibited, but rather enhanced by Dac treatment. Although all these data are fully supportive of our hypothesis that Dac therapy leads to the paradoxical IL-2–driven activation and expansion of CD56^bright NK cells, they do not exclude the possibility that other cytokines, such as IL-15, may also contribute to this effect in vivo. This is especially true because soluble IL-15, which we used in vitro, may not have the same effect as IL-15 transpresented in a complex bound to the IL-15R α-chain (26) in vivo. However, although the IL-15/IL-15Rα complex activates human NK cells in vivo, it also induces differentiation of CD56^bright NK cells into CD56^dim NK cells (26), which we did not observe in association with Dac treatment. Collectively, our in vitro data supported by stated in vivo observations provide strong support for the role of IL-2 in the activation of CD56^bright NK cells in Dac-treated subjects.

Our data do not contradict the possibility raised by Fehniger et al. (19) that IL-2 enhances differentiation of CD56^bright NK cells from its precursors and inhibits apoptosis of these regulatory NK cells in vivo. In fact, we believe that both of these hypotheses are fully compatible. Although Fehniger et al. did not observe increased proliferation of CD56^bright NK cells expanded by IL-2 administration to cancer patients, these individuals were also treated with chemotherapeutic agents that could have inhibited cellular proliferation in vivo. Our Ki67 data clearly demonstrate that CD56^bright NK cells proliferate more vigorously after initiation of Dac treatment in MS patients. The second difference between our study and the one performed by Fehniger et al. (19) lies in the source of IL-2; whereas, in one system IL-2 is provided exogenously, in our system, IL-2 is produced in the physiological environment (i.e., lymph node), most likely as a result of immune activation. As such, it is produced in a coordinated manner and likely in association with the production of other physiological factors (e.g., IL-15 by dendritic cells or activated monocytes/macrophages and IL-7 by stromal cells) that may further enhance the activation of CD56^bright NK cells and their successful entry into the proliferation cycle. Our observation that as absolute numbers of CD56^bright NK cells increase during Dac therapy, the proportion of proliferating CD56^bright NK cells decreases (between T1 and T2 of Dac therapy) suggests that expanded NK cells compete for a limited source of IL-2 in vivo, which ultimately leads to a plateau of CD56^bright NK cells expansion during long-term Dac therapy.

As would be expected from its efficient blockage of the IL-2–binding tac epitope on CD25, Dac therapy inhibits the ability of T cells to phosphorylate STAT5 in response to 50–100 IU of exogenous IL-2. However, Dac does not inhibit STAT5 phosphorylation to the same dose of exogenously added IL-2 on CD56^bright NK cells, despite the fact that a proportion of these cells (10–80% depending on the donor) express CD25 ex vivo. 50 IU/ml is a marginal dose (∼2 nM) for intermediate-affinity IL-2R; indeed we have observed that decreasing this dose further to 10–20 IU/ml (<1 nM) abrogates signaling also on NK cells (data not shown), consistent with published data that K_d for IL-2 binding to intermediate-affinity receptor is 1 nM (12). However, the fact that CD56^bright NK cells still phosphorylate STAT5 to 50–100 IU/ml of IL-2 in the presence of Dac, although T cells do not, indicates that there is a wider range of IL-2 concentrations that can trigger signaling through the intermediate-affinity IL-2R and that the final effective concentration likely depends on the expression levels of IL-2R β-chain (CD122) and -γ_c (CD132). We have previously demonstrated that CD56^bright NK cells have 10-fold higher expression of CD122 in comparison with CD56^dim NK cells and 100-fold higher expression of CD122 in comparison with T cells (5), a finding that likely underlies the differences we observed in the current study in the ability of CD56^bright NK cells versus T cells to use intermediate-affinity IL-2R for signaling.

When comparing baseline numbers and proliferation of CD56^bright NK cells in patients participating in the Dac monotherapy trial, versus those that were enrolled in the Dac add-on trial to IFN-β, we observed significantly higher numbers of CD56^bright NK cells in patients who were on IFN-β therapy in comparison with untreated patients. This observation is in agreement with the recent report of an expanded number of CD56^bright NK cells in IFN-β–treated patients (27) and supports our previously stated conclusion that IFN-β and Dac have an additive effect in the treatment of MS (17). Nevertheless, even in patients who participated in the Dac with IFN-β combination therapy trial, the numbers of CD56^bright NK cells and their cytotoxicity further increased after withdrawal of IFN-β (i.e., between T1 and T2), indicating that Dac is significantly more potent than IFN-β in expanding and activating CD56^bright NK cells in vivo.

Another unexpected finding in our study was the observation that the Dac-induced decrease in STAT5 phosphorylation to IL-2 was associated with a compensatory increase in IL-7 signaling on T cells. Both IL-2 and IL-7 (in addition to IL-4, -9, -15, and -21) use the γ_c (CD132) as a part of their signaling complex. Animal data suggest that a hierarchy exists in the preferential recruitment of γ_c to IL-15R (and likely also IL-2R) as opposed to the IL-7R (28). Furthermore, IL-2 signaling inhibits IL-7R α-chain (CD127) expression in activated T cells (29), indicating that although there is a partial overlap in the effects of these two cytokines on T cells (e.g., promoting T cell survival), the IL-2 signal is dominant under physiological conditions. A Dac-induced blockade of this physiological dominance of IL-2 signaling likely enhanced the availability of γ_c for IL-7 signaling. One apparent consequence of this compensatory increase in IL-7 signaling on CD4⁺ T cells is the stabilization of FoxP3⁺ Treg numbers and their in vivo proliferation during prolonged Dac treatment (i.e., between T1 and T2). Indeed, it was in this particular population of CD4⁺ T cells where we observed an increase in IL-7Rα (CD127) expression induced by Dac treatment. Animal studies indicate that some γ_c-signaling cytokines (specifically IL-7, and to a much lesser degree IL-15 but not IL-21) are able to enhance viability and upregulate FoxP3 expression in Tregs (24, 30), although not to the point that they can fully rescue the phenotype of IL-2–deficient mice. We observed stabilization of Treg numbers but also their levels of FoxP3 expression between T1 and T2 therapy time points, at levels that represented 86% (MFI) to 60% (number of Tregs) of baseline values. Our data represent to our knowledge the first in vivo evidence that such a compensatory function of IL-7 on Tregs survival and proliferation also occurs in humans. Furthermore, these data present evidence for in vivo cross-regulation between CD25 and CD127-mediated signaling, which, based on genetic data (11), may play an important role in the development of human autoimmunity.

CD56^bright NK cells are emerging as an important regulatory population in MS; however, it remains unclear whether they exert an immunoregulatory role on T cell responses predominantly in the lymphoid organs or whether they gain access to the CNS compartment. We believe that expansion of CD56^bright NK cells occurs predominantly in lymph nodes, where IL-2 is produced under physiological conditions in the healthy immune system (31). Indeed, several patients treated with Dac developed generalized lymphadenopathy with expansion of CD56^bright NK cells confirmed by biopsy (B. Bielekova and J. Perry, unpublished observations). Although the current study focused on determining the mechanism of Dac-induced expansion of CD56^bright NK cells, follow-up studies will be required to expand our understanding of how exactly these cells contribute to inhibition of CNS inflammation in MS patients.

In conclusion, this current study significantly expands our knowledge about the pleiotrophic effects of this CD25-targeting therapy on the human immune system. Perhaps the most striking results in this regard are the vastly different effects of Dac therapy on two populations of CD25-expressing immunoregulatory cells: FoxP3⁺ Tregs and CD56^bright NK cells. Proliferation of FoxP3⁺ Tregs is inhibited and their numbers are contracted; whereas, proliferation of CD56^bright NK cells is induced and their numbers are significantly expanded by CD25 blockade. We provide evidence that the main reason for this discrepancy lies in the differential ability of the intermediate-affinity IL-2R expressed on T cells versus CD56^bright NK cells to respond to physiological doses of IL-2. Thanks to a compensatory in vivo increase in IL-7 signaling, the quantitative decline in FoxP3⁺ Tregs under Dac therapy is significantly smaller (∼30–40%) than the prominent increase in CD56^bright NK cell numbers (>300%), proliferation (>400%) and their functional activity (∼180%). As it is likely that different immunoregulatory cell populations have overlapping in vivo functions, perhaps this is the main reason why Dac therapy is generally well tolerated and has not been associated so far with the induction of lymphoproliferation and autoimmunity that has been observed in CD25-deficient animals.

We thank Azita Kashani for expert blood processing and Dr. Richard Morgan of the Surgery Branch, National Cancer Institute, National Institutes of Health, for the generous donation of plasmids for the eGFP transduction of K562 cells.

Disclosures B.B. is a coinventor on patents owned by the National Institutes of Health on the use of Dac in MS and has received patent royalty payments.