The Aryl Hydrocarbon Receptor Antagonist StemRegenin1 Improves In Vitro Generation of Highly Functional Natural Killer Cells from CD34 + Hematopoietic Stem and Progenitor Cells

Abstract

Early natural killer (NK)-cell repopulation after allogeneic stem cell transplantation (allo-SCT) has been associated with reduced relapse rates without an increased risk of graft-versus-host disease, indicating that donor NK cells have specific antileukemic activity. Therefore, adoptive transfer of donor NK cells is an attractive strategy to reduce relapse rates after allo-SCT. Since NK cells of donor origin will not be rejected, multiple NK-cell infusions could be administered in this setting. However, isolation of high numbers of functional NK cells from transplant donors is challenging. Hence, we developed a cytokine-based ex vivo culture protocol to generate high numbers of functional NK cells from granulocyte colony-stimulating factor (G-CSF)-mobilized CD34⁺ hematopoietic stem and progenitor cells (HSPCs). In this study, we demonstrate that addition of aryl hydrocarbon receptor antagonist StemRegenin1 (SR1) to our culture protocol potently enhances expansion of CD34⁺ HSPCs and induces expression of NK-cell-associated transcription factors promoting NK-cell differentiation. As a result, high numbers of NK cells with an active phenotype can be generated using this culture protocol. These SR1-generated NK cells exert efficient cytolytic activity and interferon-γ production toward acute myeloid leukemia and multiple myeloma cells. Importantly, we observed that NK-cell proliferation and function are not inhibited by cyclosporin A, an immunosuppressive drug often used after allo-SCT. These findings demonstrate that SR1 can be exploited to generate high numbers of functional NK cells from G-CSF-mobilized CD34⁺ HSPCs, providing great promise for effective NK-cell-based immunotherapy after allo-SCT.

Introduction

Natural killer (NK) cells are CD3⁻CD56⁺ lymphocytes, which are part of the innate immune system and play an important role in the defense against virus-infected and transformed cells. NK-cell activation and subsequent killing of target cells are regulated by a balance in their expression levels of inhibitory receptors, including the killer immunoglobulin-like receptors (KIRs) and CD94/NKG2A heterodimer versus activating receptors, such as DNAX accessory molecule-1 (DNAM-1), natural cytotoxicity receptors (NCRs), and NKG2D. In homeostasis, NK cells are inhibited by their inhibitory receptors recognizing self human leukocyte antigen (HLA) class I molecules and/or HLA-E molecules presenting conserved HLA class I leader sequences. However, an NK-cell-mediated antitumor effect can be induced by upregulation of activating ligands or downregulation of HLA class I molecules on tumor cells. In addition, in the setting of haploidentical allogeneic stem cell transplantation (allo-SCT), donor NK cells expressing inhibitory KIRs may not be engaged by recipient HLA class I molecules and hence be activated. This phenomenon is called missing-self recognition and can contribute to the curative graft-versus-tumor (GVT) effect [1].

Because of their ability to kill tumor cells, NK cells are considered potent effectors for adoptive immunotherapy against cancer. So far, promising results have been obtained by infusion of haploidentical NK cells after immunosuppressive chemotherapy in adult and childhood acute myeloid leukemia (AML) [2 –4]. However, a limitation of these studies is the relatively low NK-cell numbers that can be enriched from apheresis products for multiple infusions. Furthermore, contaminating alloreactive T cells risk the induction of graft-versus-host disease (GVHD), especially when interleukin (IL)-2 or IL-15 is coadministered to boost NK-cell survival and expansion. To generate high numbers of allogeneic NK cells completely devoid of T-cell contamination, a good manufacturing practice (GMP)-compliant, cytokine-based ex vivo culture protocol has been developed by our group [5,6]. Using this procedure, CD34⁺ hematopoietic stem and progenitor cells (HSPCs) isolated from umbilical cord blood (UCB) can be expanded over 2,000-fold in large-scale bioreactors into a mixture of immature and mature NK cells with a purity >80%. Preclinical studies conducted in NOD/SCID-IL2Rγ^null mice demonstrated that these HSPC-NK cells have bone marrow (BM) homing capacity, display IL-15-driven in vivo expansion, and prolong survival of leukemia-bearing mice [7]. Currently, administration of this HSPC-NK-cell product after immunosuppressive chemotherapy is being investigated in a phase I clinical trial in older AML patients who are not eligible for allo-SCT (www.trialregister.nl and search for 2818).

In HLA-matched nonmyeloablative and T-cell-depleted allo-SCT, early NK-cell repopulation has been associated with decreased relapse rates, without increasing GVHD incidence [8,9]. Moreover, high NK-cell numbers in stem cell grafts have been associated with a decreased incidence of GVHD [10]. In addition, transplants from donors with KIR-B haplotypes, containing several activating KIRs, led to lower rates of relapse and improved survival [11 –13]. For these reasons, it would be highly valuable to exploit HSPC-NK-cell products for adoptive immunotherapy after allo-SCT. Since NK cells of donor origin will not be rejected, multiple NK-cell infusions, without the need for immunosuppressive chemotherapy to prevent rejection, could be administered after allo-SCT. Consequently, these cells may potentially induce long-term GVT effects. However, to obtain large numbers of NK cells from donor origin, peripheral blood (PB)-derived or BM-derived CD34⁺ HSPCs, which have a lower expansion potential compared with UCB-derived CD34⁺ HSPCs, should be expanded and differentiated into NK cells.

Recently, it was described that expansion of CD34⁺ HSPCs can be enhanced by inhibition of the aryl hydrocarbon receptor (AhR) using the antagonist StemRegenin1 (SR1) [14]. AhR is a ligand-inducible transcription factor, which plays an important role in biological responses toward xenobiotic agents, such as digoxin [15,16]. However, it has become clear that AhR also has multiple naturally occurring ligands, like tryptophan metabolites and dietary compounds [15]. Furthermore, AhR turned out to regulate differentiation of multiple immune cells, including dendritic cells [17,18], regulatory T cells [19,20], γδ T cells [21], T helper 17 (Th17) cells [22], and notably NK cells [23]. Based on these findings, we hypothesized that addition of SR1 to our culture system might improve expansion of CD34⁺ HSPCs and differentiation of these cells into NK cells. We found that SR1 not only enhances expansion of CD34⁺ HSPCs but also upregulates the expression of early and late NK-cell-specific transcription factors, thereby potentiating differentiation of SR1-expanded CD34⁺ cells into NK cells. These SR1-induced NK cells have a high purity, express high levels of activating receptors, and efficiently target AML and multiple myeloma (MM) cells. Importantly, proliferation and cytolytic functions of SR1-induced HSPC-NK cells are not inhibited by cyclosporin A (CsA), in contrast to mycophenolic acid (MPA), which is used only shortly after allo-SCT, facilitating multiple infusions relatively shortly after allo-SCT. Therefore, our SR1 culture system holds great promise for future donor HSPC-NK-cell adoptive immunotherapy after allo-SCT to boost antitumor and antiviral immunity, leading to prolonged relapse-free survival.

Materials and Methods

Cell lines

Cell lines (K562, THP1, HL-60, U266, UM9, and RPMI8226) were cultured in the Iscove's modified Dulbecco's medium (IMDM; Invitrogen, Carlsbad, CA) containing 50 U/mL penicillin, 50 μg/mL streptomycin, and 10% fetal calf serum (FCS; Integro, Zaandam, The Netherlands).

Isolation of CD34⁺ stem and progenitor cells

BM samples were collected from healthy donors after obtaining written informed consent. Bone marrow-derived mononuclear cells were isolated by the Ficoll–Hypaque (1.077 g/mL; GE Healthcare, Uppsala, Sweden) density-gradient centrifugation. Peripheral blood-derived mononuclear cells were obtained from apheresis material of stem cell donors who were treated with granulocyte colony-stimulating factor (G-CSF; Neupogen^®) 10 μg/kg/day subcutaneously for 5–6 days after obtaining written informed consent. CD34⁺ HSPCs were isolated using anti-CD34 immunomagnetic beads (Miltenyi Biotec, Bergisch Gladbach, Germany) according to the manufacturer's instructions. CD34⁺ HSPCs were directly used for NK-cell generation.

Ex vivo expansion of CD34⁺HSPCs and differentiation into NK cells

CD34⁺ HSPCs were plated into 24-well or 6-well tissue culture plates (Corning, Inc., Corning, NY). Cells expanded during 9 or 10 days with a high-dose cytokine cocktail (expansion cocktail I) consisting of 25 ng/mL IL-7 (ImmunoTools, Friesoythe, Germany), 25 ng/mL stem cell factor (SCF; ImmunoTools), 25 ng/mL thrombopoietin (TPO; CellGenix, Freiburg, Germany), and 25 ng/mL Flt3L (ImmunoTools). From day 9 or 10 to day 14 or 15, TPO was replaced by 20 ng/mL IL-15 (Miltenyi Biotec). After day 14 or 15, cell differentiation was initiated by replacing expansion cocktail I by a new high-dose cytokine cocktail (differentiation cocktail) consisting of 20 ng/mL IL-7, 20 ng/mL SCF, 20 ng/mL IL-15, and 1,000 U/mL IL-2 (Proleukin^®; Chiron, München, Germany). Where mentioned, 2 μM SR1 (Cellagen Technology, San Diego, CA) was added to the culture medium (Fig. 1A). Cells were cultured in the CellGro GMP DC medium (CellGenix) supplemented with 10% human serum (Sanquin Bloodbank, Nijmegen, The Netherlands) and a low-dose cytokine cocktail consisting of 250 pg/mL G-CSF (filgrastim, “Neupogen”; Amgen, Inc., Thousand Oaks, CA), 10 pg/mL GM-CSF (ImmunoTools), and 50 pg/mL IL-6 (ImmunoTools). During the first 14 or 15 days of culture, low-molecular-weight heparin (Clivarin^®; Abbott, Wiesbaden, Germany) was added to the medium at a final concentration of 20 μg/mL. Freshly isolated CD34⁺ cells were plated at a concentration of 1–4 × 10⁵/mL. After 3 days of culture, cells were transferred to a new plate to deplete stromal cells. From day 14 or 15 onward, cell counts were kept above 2 × 10⁶ cells/mL. Cell cultures were refreshed with at least 30% new medium every 2–3 days. Cultures were maintained at 37°C in 95% humidity and 5% CO₂. NK cells were used in experiments after 5 weeks of culture.

FIG. 1.

StemRegenin1 (SR1) enhances expansion of peripheral blood-derived CD34⁺ hematopoietic stem and progenitor cells (HSPCs) and improves natural killer (NK)-cell differentiation. (A) Representation of the cytokine-based ex vivo culture protocol used for the generation of NK cells. (B–D) Kinetics of differentiation (B) and expansion (C) of CD34⁺ HSPCs in our ex vivo culture protocol in the presence or absence of 2 μM SR1 in one representative donor, determined by flow cytometry (FCM). (D) Summary of total expansion, NK-cell yield [calculated by (total expansion × percentage of CD56⁺ cells)/100], and NK-cell differentiation after 5 weeks of culture in our ex vivo culture protocol in the presence or absence of 2 μM SR1 in 10 donors, determined by FCM. **P < 0.01, ***P < 0.001, Wilcoxon matched-pairs signed-rank test, paired one-tailed Student's t-test.

RNA isolation and real-time quantitative reverse transcriptase polymerase chain reaction

Total RNA from 0.5 to 2 × 10⁵ cells, collected weekly from HSPC-NK-cell cultures, was isolated using the Quick-RNA^™ MicroPrep Kit (Zymo Research, Irvine, CA). Next, cDNA was synthesized using M-MLV reverse transcriptase (Invitrogen) in a standard reaction as described earlier [24], after which real-time polymerase chain reaction (PCR) was performed using the following TaqMan gene expression assays (Applied Biosystems, Forster City, CA): AhRR (Hs01005075_m1), TOX (Hs01055573_m1), ID2 (Hs04187239_m1), EOMES (Hs00172872_m1), GATA3 (Hs00231122_m1), SH2D1B (Hs01592483_m1), IFN-γ (Hs00989291_m1), GZMB (Hs01554355_m1), and PRF1 (Hs99999108_m1). For all genes, Ct values were normalized to GAPDH (Hs02758991_g1) by calculating ΔCt = Ct_{target gene} − Ct_GAPDH per sample. Finally, gene expression levels were quantified relative to GAPDH as follows: 2^(−[ΔCt]).

Flow cytometry

Cell numbers and expression of cell surface markers were determined by flow cytometry (FCM). Anti-human CD45-ECD (J.33; Beckman Coulter, Woerden, The Netherlands) and anti-CD56-PC7 (HCD56; BioLegend, San Diego, CA) antibodies were used to follow cell number and NK-cell differentiation during culture using the Coulter FC500 flow cytometer (Beckman Coulter). The population of viable CD45⁺ cells was determined by exclusion of 7-AAD-positive cells (Sigma, St. Louis, MO). For phenotypical analysis, cells were incubated with antibodies in the FCM buffer (phosphate-buffered saline/0.5% bovine serum albumin; Sigma) for 30 min at 4°C. After washing, cells were resuspended in the FCM buffer and analyzed. The following conjugated monoclonal antibodies were used for NK-cell phenotyping: anti-NKG2A-PE (Z199; Beckman Coulter), anti-DNAM-1-FITC (DX11; BD Biosciences Pharmingen, Breda, The Netherlands), anti-CD16-FITC (3G8), anti-CD3-FITC (UCHT1), anti-NKG2D-PE (1D11), anti-NKp30-PE (P30-15), anti-NKp44-PE (P44-8), anti-NKp46-PD (9E5), anti-CD158b-PE (Dx27), anti-CD158e-PE (Dx9), anti-CD158a/h-PE (HP-MA4), anti-CD62L-PE (DREG56), anti-CD253-PE (RIK-2), anti-CXCR3-PE (G025H7), anti-CXCR4-PE (12G5), anti-IgG1-PE (MOPC-21), anti-IgG2a-PE (MOPC-173), anti-IgG2b-PE (MCP-11), and anti-IgG1-FITC (MOPC-21; all from BioLegend).

Fluorescence-activated cell sorting of HSCP-NK cells

After 5 weeks of culture, CD56⁺ cells were isolated from the total cultured cells. For this purpose, cells were stained for 15 min at 4°C using the appropriate concentration of anti-CD56-PEcy7 (CD56-PC7, HCD56; BioLegend). Cells were washed and resuspended in the fluorescence-activated cell sorting (FACS) buffer at a concentration of 1.5 × 10⁶/mL and subsequently sorted at the FACSAria Cell Sorter (BD Biosciences Pharmingen).

FCM-based cytotoxicity assays

Cell lines were labeled with 1 μM carboxyfluorescein diacetate succinimidyl ester (CFSE; Molecular Probes; Invitrogen, Eugene, OR), and primary AML blasts were labeled with 1.5 μM CFSE, both at a concentration of 1 × 10⁷/mL for 10 min at 37°C. The reaction was terminated by adding an equal volume of FCS. After washing, cells were resuspended in IMDM/10% FCS to a final concentration of 3 × 10⁵/mL. Target cells (3 × 10⁴) were cocultured in triplicate with effector cells at different effector:target (E:T) ratios in a total volume of 200 μL IMDM/10% FCS in 96-well round-bottom plates (Corning, Inc.). Effector cells and target cells alone were plated out in triplicate as controls. In experiments with primary AML blasts, AML blasts were derived from BM samples from five patients at the time of diagnosis and were supplemented with IL-3 (50 ng/mL; CellGenix), SCF (25 ng/mL), Flt3L (20 ng/mL), GM-CSF (100 ng/mL), G-CSF (100 ng/mL), and IL-15 (5 ng/mL). To measure degranulation of NK cells, anti-CD107a-PC7 (H4A3; BioLegend) was added to the coculture. After overnight coculture at 37°C, 50 μL supernatant was discarded and 50 μL Coulter^® Isoton^® II Diluent containing 0.2 μL 7-AAD was added instead. Cells were harvested, and the number of viable target cells was quantified by FCM by gating on forward scatter and side scatter and exclusion of 7-AAD-positive cells. Target cell survival was calculated as follows: % survival = [(absolute number of viable CFSE⁺ target cells cocultured with NK cells)/(absolute number of viable CFSE⁺ target cells cultured in medium)] ×100%. The percentage of specific lysis was calculated as follows: % lysis = [100 − (% survival)], as described earlier by Jedema et al. [25,26]. Degranulation of NK cells during overnight coculture was determined as the percentage of CD107a expressing cells measured by FCM.

Enzyme-linked immunosorbent assays

The production capacity of interferon (IFN)-γ and Granzyme B by NK cells was evaluated by enzyme-linked immunosorbent assay according to the manufacturer's instructions (IFN-γ; Pierce Endogen, Rockford, IL, and Granzyme B; Mabtech, Nacka Strand, Sweden). To this end, NK cells (1 × 10⁵) were cocultured in triplicate with target cells (1 × 10⁵) in a total volume of 200 μL IMDM/10% FCS in 96-well round-bottom plates (Corning, Inc.). NK cells alone were plated in triplicate as controls. After incubation overnight at 37°C, 150 μL supernatant was collected and stored at −20°C until use.

Proliferation assays

HSPC-NK cells were cultured according to our culture protocol for 35 days. After 35 days, NK cells were labeled with 1 μM CFSE as described previously. After staining, cells were resuspended in a differentiation medium at a concentration of 2 × 10⁶/mL and plated in duplicate in a 96-well plate. CsA (BioVision, Inc., Milpitas, CA) or MPA (Sigma-Aldrich, Zwijndrecht, The Netherlands) was added to final concentrations of 0.01–1 μg/mL (CsA) or 0.1–10 μg/mL (MPA). Half of the medium containing MPA or CsA in the final concentration was refreshed every 2–3 days. After 7 days, cells were harvested. The number of viable target cells was quantified by FCM by gating on forward scatter and side scatter and exclusion of 7-AAD-positive cells. Proliferation was analyzed by determining the CFSE dilution within CD56⁺ cells.

Statistical analysis

Results from different experiments are described as mean ± standard error of the mean. Statistical analysis was performed using a one-tailed paired Student's t-test, a two-tailed unpaired Student's t-test, or a one-way analysis of variance if values had a normal distribution (normality was determined using the Kolmogorov–Smirnov test). For values without normal distribution, we used the Wilcoxon matched-pairs signed-rank test. Differences were considered to be significant for P values <0.05.

Results

SR1 enhances expansion of PB- and BM-derived CD34⁺ HSPCs and improves NK-cell differentiation

In this study, we investigated whether NK cells could be generated in vitro from PB- or BM-derived CD34⁺ HSPCs. We used the feeder-free culture protocol described in Fig. 1A, which was reported previously to generate high numbers of functional NK cells from UCB-derived CD34⁺ HSPCs [5,6]. However, in our initial cultures using PB- or BM-derived HSPCs, expansion and differentiation were low, which was associated with high numbers of stroma-like cells observed in culture plates (data not shown). Therefore, we investigated transfer of nonadherent cells to a new culture plate after 3 days of culture. Although this resulted in much lower outgrowth of stromal cell layers in the culture plates, expansion and NK-cell differentiation were still low. For this reason, we investigated whether addition of the AhR antagonist SR1 (2 μM), which is known to improve expansion of CD34⁺ HSPCs and to influence NK-cell differentiation, could improve NK-cell generation from HSPCs. Importantly, addition of SR1 strongly enhanced expansion of CD34⁺ cells from both PB-derived HSPCs (Fig. 1) and BM-derived HSPCs (Supplementary Fig. S1A, B; Supplementary Data are available online at www.liebertpub.com/scd). Mean NK-cell yield after 5 weeks of culture was 235-fold (range, 115- to 904-fold; n = 10; Fig. 1D) for G-CSF-mobilized CD34⁺ cells and 129-fold (range, 33- to 301-fold; n = 4; Supplementary Fig. S1B) for BM-derived CD34⁺ HSPCs. Interestingly, differentiation also improved by addition of SR1 to our cultures, resulting in a purity of 83% ± 9% for G-CSF-mobilized CD34⁺ HSPCs and 84% ± 18% for BM-derived CD34⁺ cells. The remaining non-NK cells in the cultures represented mainly CD14⁺ and/or CD15⁺ mature monocytic and myelocytic cells (11% ± 6%). A small frequency of CD34⁺ cells could also be detected at the end of the culture process (0.7% ± 0.4%). Most importantly, the amount of CD3⁺ T cells in the SR1-based NK-cell cultures investigated was very low (0.1% ± 0.1%). Altogether, these data demonstrate that the AhR pathway negatively influences NK-cell development and that by combining SR1 with cytokine mixtures, high numbers of CD56⁺ NK cells can be generated from G-CSF-mobilized and BM-derived CD34⁺ HSPCs ex vivo. Since G-CSF-mobilized CD34⁺ HSPCs are the main stem cell source for allo-SCT, we concentrated on these HSPC-NK cells for further investigations.

SR1 influences expression of transcription factors important for NK-cell differentiation and maturation

To gain insight into the molecular processes behind the SR1-enhanced differentiation of CD34⁺ HSPCs into NK cells, we analyzed the gene expression profile of several transcription factors that are described to be important for NK-cell differentiation and maturation [27 –31]. To analyze the culture composition, we concomitantly determined the percentage of CD56⁺ cells in our cultures at different time points (Fig. 2A). Next, we compared expression levels of several transcription factors in the presence or absence of SR1 in total cells at these time points in our ex vivo culture system (Fig. 2B). We observed efficient downregulation of aryl hydrocarbon receptor repressor (ie, direct target gene of AhR signaling [14]) in the presence of SR1, indicating that a concentration of 2 μM SR1 is sufficient for AhR inhibition. Interestingly, we observed higher expression of thymocyte selection-associated HMG box factor (TOX), which is important in early NK-cell differentiation [27], from day 7 onward (Fig. 2B, C). This suggests that SR1 increased the number of NK-cell precursors, even before the induction of NK-cell differentiation was initiated by addition of IL-15 to the culture and before CD56 acquisition. Expression of ID2, which is important for NK-cell maturation [28,29], increased from day 14 onward, after addition of IL-15 to the culture (Fig. 2B). Eomesodermin (EOMES), another factor important for NK-cell maturation, was upregulated from week 3 onward. Finally, expression of GATA-3 and the Ewing's sarcoma-associated transcript 2 (EAT-2; SH2D1B), required to develop cytotoxic functions [30,31], was increased in cells cultured in the presence of SR1 (Fig. 2B).

FIG. 2.

SR1 reduces expression of aryl hydrocarbon receptor repressor (AhRR) and increases expression of several transcription factors important for NK-cell differentiation and NK-cell effector functions. Cells generated from CD34⁺ HSPCs in the presence or absence of SR1 were collected at different time points. (A) Kinetics of CD56 expression in the cultures used for weekly mRNA isolation was determined using FCM. Mean ± standard error of the mean (SEM) of 4–6 different donors. (B) Expression levels of AhRR and transcription factors relative to GAPDH at different time points during culture were determined using quantitative reverse transcriptase polymerase chain reaction (qRT-PCR). Mean ± SEM of 4–6 different donors. (C) Expression levels of AhRR and TOX relative to GAPDH after 7 days of culture were determined using qRT-PCR. Mean ± SEM of six different donors. (D) Expression levels of AhRR and transcription factors in CD56⁺ cells, which were FACS sorted from cultures after 5 weeks of culture, were determined using qRT-PCR. Mean ± SEM of four different donors. *P < 0.05, paired one-tailed Student's t-test. FACS, fluorescence-activated cell sorting.

To investigate whether the increased expression levels of these transcription factors reflected a difference between NK cells generated in the presence or absence of SR1, or resulted from the different compositions of the cultures, CD56⁺ NK cells, generated in the presence or absence of SR1, were sorted by the FACS from HSPC-NK cultures after 5 weeks. Subsequently, gene expression levels of the previously mentioned transcription factors were determined. We did not observe increased expression of the NK-cell differentiation factors, TOX, ID2, and EOMES, suggesting that SR1 increases the number of NK-cell progenitors resulting in more NK cells, but it does not intrinsically change HSPC-NK cells (Fig. 2D). In addition, expression levels of GATA-3 and EAT-2 were similar in CD56⁺ NK cells cultured in the presence of SR1. Expression of GZMB and PERF was slightly higher but not significantly increased. Furthermore, IFN-γ was not increased in NK cells cultured in the presence of SR1, but these cells were not exposed to target cells before mRNA analysis.

Collectively, these data demonstrate that addition of SR1 to our ex vivo culture system blocks function of AhR, resulting in the upregulation of several transcription factors that are required for NK-cell differentiation and maturation.

SR1-generated HSPC-NK cells have an activated and a mature phenotype

To further elucidate the effect of SR1 on NK-cell activation status and function, we investigated the influence of SR1 on the phenotype of our ex vivo-generated CD56⁺ NK cells. After 35 days of culture in the presence of SR1, NK-cell phenotype was analyzed using FCM (Fig. 3A–D and Supplementary Fig. S1C). SR1-generated NK cells expressed high levels of NKG2A, which indicates transition between stage 3 and stage 4 NK-cell progenitors [32,33]. CD16, important for antibody-dependent cell-mediated cytotoxicity, was expressed on 22.7% ± 2.6% of the CD56⁺ cells. Furthermore, we found high expression levels of the activating markers, DNAM-1, NKG2D, NCRs, and TRAIL. Expression of KIRs was observed in a low percentage of CD56⁺ cells. However, expression levels were similar as observed for UCB-NK cells generated in our culture system [5,6,32]. Importantly, our NK cells also expressed high levels of CD62L and chemokine (C-X-C motif) receptor 3 (CXCR3), which are involved in homing to the lymphoid organs and trafficking of NK cells toward inflammation in vivo [7,34,35]. Interestingly, the presence of CD62L, CXCR3, DNAM-1, and TRAIL was significantly higher in CD56⁺ cells generated in the presence of SR1 (Fig. 3C, D). Increased expression of DNAM-1 and TRAIL suggests that NK cells generated in the presence of SR1 are more active compared with those generated in the absence of SR1. We did not find significant differences in expression of the other molecules in the cultures with SR1 compared with those without SR1 (data not shown). Altogether, these results indicate that SR1-generated HSPC-NK cells consist of a mixture of immature and mature NK cells expressing various activating receptors similar to our previously described UCB-derived NK cells [6].

FIG. 3.

NK cells generated in the presence of SR1 have an active and a mature phenotype. NK cells were generated from CD34⁺ progenitor cells in the presence or absence of SR1. After 5 weeks, a phenotypical analysis was performed by FCM. (A) Phenotype of a representative NK-cell product. Density plot of live cells gated on forward scatter/side scatter. (B) Expression levels of several NK-cell-specific surface antigens on live CD56⁺ NK cells from 3 to 8 different donors (mean ± SEM) are shown. (C) Expression levels of NK-cell-specific antigens on CD56⁺ NK cells generated in the presence (gray histograms) or absence (black histogram) of SR1 from one representative donor compared with isotype controls (white histogram). Numbers represent % positive cells. (D) Expression levels of NK-cell-specific antigens on CD56⁺ NK cells generated in the presence or absence of SR1 from 3 to 7 different donors (mean ± SEM) are shown. *P < 0.05, **P < 0.01, paired one-tailed Student's t-test.

SR1-generated HSPC-NK cells have efficient IFN-γ production capacity and cytolytic activity against AML and MM cells

Next, we investigated the functional activity of SR1-generated NK cells and compared them with NK cells generated in the absence of SR1. After 5 weeks of culture, NK-cell products were harvested and cocultured overnight with AML or MM tumor cell lines. Target cell-induced IFN-γ production was determined. We found that our SR1-generated HSPC-NK cells have a good IFN-γ producing capacity, which is improved in cells generated in the presence of SR1 (Fig. 4A). Subsequently, we compared the killing capacity of NK cells generated in the presence or absence of SR1. Therefore, 1 × 10⁴ CD56⁺ NK cells were cocultured overnight with different AML cell lines. We observed a significantly improved killing of K562 and HL-60 cells and also a trend toward improved killing of THP-1 cells by SR1-generated NK cells (Fig. 4B). To rule out an effect of non-NK cells in the cultures, we repeated the killing experiments using sorted NK cells. Therefore, CD56⁺ NK cells were sorted from the NK-cell products after 5 weeks of culture. Next, NK cells were cocultured overnight with AML or MM tumor cell lines as described earlier. Importantly, we also observed higher killing of K562, HL-60, and RPMI8226 cells by sorted SR1-generated NK cells (Fig. 4C). In addition, Granzyme B production was enhanced in SR1-generated HSPC-NK cells compared with NK cells generated in the absence of SR1 (Fig. 4D). These results indicate that addition of SR1 to our culture system not only increases the number of CD56⁺ cells, but also enhances the functional activity of the CD56⁺ NK cells.

FIG. 4.

NK cells generated in the presence of SR1 are functionally active. NK cells were generated from CD34⁺ HSPCs in the presence or absence of SR1. After 5 weeks, the functional activity of the cells was investigated. (A–F) NK cells were cocultured overnight with the leukemia cell lines, K562, HL-60, or THP-1, or the multiple myeloma (MM) cell lines, RPMI8226, U266, or UM9, at an effector:target (E:T) ratio of 0.3:1, 1:1, or 3:1. (A) Interferon-γ levels were determined by enzyme-linked immunosorbent assay (ELISA) in the coculture supernatants of 1 × 10⁵ CD56⁺ NK cells and 1 × 10⁵ target cells. Mean ± SEM of 3–4 donors generated in the presence or absence of SR1 are shown. (B) Specific killing of leukemia cell lines at an E:T ratio of 0.3:1 was determined in an FCM-based cytotoxicity assay. Mean ± SEM of three donors generated in the presence or absence of SR1 are shown. (C, D) After 5 weeks of culture, CD56⁺ NK cells were FACS sorted from the cultures and cocultured with leukemia cell lines or the MM cell line RPMI8226 at an E:T ratio of 1:1. Cultures were supplemented with interleukin (IL)-15 (5 ng/mL). (C) Specific killing of the cell lines was determined in an FCM-based cytotoxicity assay. Data are depicted as mean ± SD. (D) Granzyme B production by the CD56⁺ NK cells was measured by ELISA. Data are depicted as mean ± SD. (E, F) Degranulation of SR1-generated CD56⁺ cells after coculture at an E:T ratio of 1:1, determined by FCM as the percentage of CD107a expressing cells. One representative donor (E) and mean ± SEM of 4–7 donors (F) are shown. (G) Specific killing of leukemia or MM cell lines by SR1-generated NK cells at different E:T ratios was determined in an FCM-based cytotoxicity assay. Mean ± SEM of 3–7 donors are shown. (H) Specific killing of primary acute myeloid leukemia (AML) cells by SR1-generated NK cells from five different patients (No. 1 AML-M2, No. 2 AML-M2, No. 3 AML-M4, No. 4 AML-M5, and No. 5 AML-M0) was determined for five different HSPC donors. Specific killing was determined after 1, 2, and 3 days of coculture in an FCM-based cytotoxicity assay at an E:T ratio of 3:1. Data are displayed as mean ± SD of triplicate samples. *P < 0.05, **P < 0.01, ***P < 0.001, paired one-tailed (A, B, F) or an unpaired two-tailed (C, D) Student's t-test.

To further confirm the functional activity of SR1-generated NK cells, we investigated degranulation of these cells after overnight coculture with AML or MM tumor cell lines. We found marked degranulation of our SR1-generated NK cells upon coculture with the different cell lines (Fig. 4E, F). Most efficient degranulation was observed against MHC^neg K562 cells as well as the MHC expressing AML cell line HL-60. We also observed marked degranulation upon coculture with the MM cell lines, U266 and RPMI8226. The MM cell line UM9 was the least-potent cell line to induce degranulation.

Next, we performed cytotoxicity experiments using SR1-generated HSPC-NK cells harvested after 5 weeks of culture. For this, NK cells were cocultured overnight with different CFSE-labeled AML (K562, HL-60, and THP-1) and MM (UM9, U266, and RPMI8226) cell lines. The next day, specific killing was determined by FCM. We observed very efficient killing of most AML and MM cell lines, even at an E:T ratio of 1:1 (Fig. 4G). Subsequently, we investigated whether patient-derived primary AML blasts were susceptible to killing by SR1-induced NK cells. Therefore, we performed cytotoxicity assays with AML blasts from five different patients. Importantly, AML blasts could be potently killed within 3 days of coculture (Fig. 4H). We observed some variance in susceptibility between the different patients, but variance between HSPC-NK-cell donors was small, indicating that the quality of the SR1-induced NK-cell products is consistent.

Taken together, these data demonstrate that SR1-induced HSPC-NK cells, generated from G-CSF-mobilized CD34⁺ cells, mediate efficient IFN-γ production, degranulation, and cytolytic activity against hematological tumor cells. Furthermore, as expected by increased expression of activating markers, SR1 augments IFN-γ production and cytotoxic activity of ex vivo-generated HSPC-NK cells.

HSPC-NK-cell proliferation, viability, and function are inhibited by MPA but not by CsA

Increased NK-cell numbers after nonmyeloablative allo-SCT have been associated with decreased relapse rates, without an increased risk for GVHD [8,9]. Hence, adoptive transfer of ex vivo-generated NK cells shortly after allo-SCT is an attractive approach to improve patient outcome. Our therapeutic strategy is to apply HSPC-NK-cell adoptive transfer for high-risk patients treated with nonmyeloablative allo-SCT (Fig. 5A). For this purpose, we want to exploit 15–30 × 10⁶ CD34⁺ cells from the G-CSF-mobilized donor stem cell graft, which represents ∼5%–10% of the total graft, for ex vivo NK-cell generation. These HSPC-NK cells can be infused as a single infusion or multiple infusions after allo-SCT to improve the GVT effect. However, these allo-SCT patients are treated with immunosuppressive drugs to prevent GVHD, so the effect of these drugs on HSPC-NK-cell proliferation and viability should be known. For this reason, we investigated the effect of CsA and MPA, which are often used after nonmyeloablative allo-SCT, on SR1-induced NK cells. For this, week 5 HSPC-NK cells were cultured for 7 additional days in the presence or absence of therapeutic concentrations of MPA or CsA (Fig. 5B–F). After 7 days, we analyzed NK-cell proliferation and viability using FCM (Fig. 5B–D). We found that proliferation of HSPC-NK cells was already inhibited by MPA at a therapeutic concentration of 0.1 μg/mL. In contrast, CsA, even at a concentration of 1 μg/mL, did not inhibit HSPC-NK-cell proliferation (Fig. 5B, C). In addition, NK-cell viability was not affected by CsA, while MPA induced dose-dependent cell death (Fig. 5D). Next, we investigated the effect of MPA and CsA on HSPC-NK-cell function. For this, CD56⁺ NK cells cultured in the presence of MPA or CsA were cocultured overnight with K562 cells at an E:T ratio of 1:1. As a control, week 6 CD56⁺ NK cells were used. We observed impaired killing by NK cells cultured in the presence of MPA; however, CsA did not inhibit NK-cell-mediated killing (Fig. 5E). In addition, we observed impaired IFN-γ production after incubation with MPA, while incubation with CsA even enhanced IFN-γ production (Fig. 5F). These data indicate that adoptive transfer of SR1-generated HSPC-NK cells can be applied in patients treated with CsA, but infusion should be postponed until cessation of MPA therapy.

FIG. 5.

NK-cell viability, proliferation, and function are inhibited by mycophenolic acid (MPA) but not by cyclosporin A (CsA). (A) Suggested strategy for adoptive transfer of ex vivo-generated SR1-NK cells after nonmyeloablative allogeneic stem cell transplantation (allo-SCT). A patient is conditioned with nonmyeloablative conditioning; subsequently, the patient receives a T-cell-replete stem cell graft. Ten percent of CD34⁺ cells of the donor graft are used for NK-cell generation. To prevent graft-versus-host disease, patients are treated with MPA and CsA. After cessation of MPA (around day 28 after transplantation), NK cells can be infused as a single infusion or multiple infusions. (B–F) The effect of immunosuppressive drugs used after nonmyeloablative allo-SCT was investigated. NK cells were generated from CD34⁺ HSPCs in the presence of SR1. After 5 weeks, NK cells were cultured for 7 more days in the presence or absence of different dosages of MPA or CsA. (B) Proliferation of carboxyfluorescein diacetate succinimidyl ester (CFSE)-labeled NK cells cultured in the presence or absence of drugs measured by CFSE dilution in a representative donor. (C) Proliferation of NK cells in the presence or absence of drugs measured by CFSE dilution. Mean fluorescence intensity (MFI) relative to MFI of control cells ± SEM of four different donors is shown. (D) The number of living cells determined by forward scatter/side scatter and exclusion of 7-AAD-positive cells. Percentage of living NK cells cultured for 7 days in the presence or absence of drugs. Mean viability relative to control cells ± SEM of four different donors is shown. (E, F) 1 × 10⁵ CD56⁺ NK cells cultured in the presence or absence of MPA or CsA were cocultured overnight with K562 cells in the presence of IL-15 (5 ng/mL) at an E:T ratio of 1:1. (E) Specific killing of K562 cells was determined in an FCM-based cytotoxicity assay. Data are depicted as mean ± SD and are representative of two different donors. (F) IFN-γ levels were measured in the supernatant after overnight coculture using ELISA. Data are depicted as mean ± SD and are representative of two different donors. **P < 0.01, ***P < 0.001, one-way analysis of variance.

Discussion

NK cells are the first lymphocyte population recovering after allo-SCT and have several important functions shortly after allo-SCT [36]. First, they are involved in defense against viral infections, such as cytomegalovirus infections [37,38], which can cause high morbidity shortly after allo-SCT. Furthermore, high NK-cell numbers shortly after nonmyeloablative and T-cell-depleted allo-SCT have been associated with reduced relapse rates without an increased risk of GVHD [8,9], indicating that allogeneic donor NK cells have specific antitumor activity. Nevertheless, it was shown that early engrafting NK cells have decreased cytokine producing capacity [39]. So, to further exploit the beneficial effects of NK cells after allo-SCT, adoptive transfer of functional and rapidly maturating NK cells would be an attractive immunotherapeutic strategy. Since donor-derived NK cells will not be rejected posttransplant, multiple NK- cell infusions without the need for immunosuppressive chemotherapy to prevent rejection will be feasible in this setting. However, high numbers of functional NK cells of donor origin are needed to apply this strategy. Since isolation of sufficient numbers of NK cells from donors, without contaminating alloreactive T cells, is challenging, we investigated if high numbers of functional NK cells could be generated from G-CSF-mobilized CD34⁺ HSPCs using a cytokine-based ex vivo culture protocol. This protocol was developed by our group earlier, and high numbers of pure and functional NK cells with in vivo maturation capacity can be generated from UCB-derived CD34⁺ HSPCs using this GMP-compliant protocol [5 –7].

We found that NK-cell expansion and differentiation from G-CSF-mobilized and BM-derived CD34⁺ cells in the absence of SR1 were very limited compared with those of UCB-derived CD34⁺ HSPCs observed in our previously developed culture protocol. Interestingly, we found that addition of the AhR antagonist SR1 to our cultures greatly enhanced NK-cell expansion and differentiation. As a result, we can expand G-CSF-mobilized CD34⁺ cells on average 268-fold using our newly developed SR1-based protocol. In addition, SR1-generated NK-cell products are 83% ± 9% pure. The remaining non-NK cells in the cultures represented mainly CD14⁺ and/or CD15⁺ mature monocytic and myelocytic cells. Probably due to SR1 addition, still some remaining low amount of CD34⁺ cells could be detected. However, contaminating CD3⁺ T cells were either not detectable or at very low frequency (0.1% ± 0.1%). Furthermore, we found that SR1-NK cells have a similar phenotype as our NK cells previously generated from UCB-derived CD34⁺ HSPCs, which has a highly active phenotype, characterized by expression of high levels of activating NK-cell receptors [5 –7]. Interestingly, expression levels of CD62L, CXCR3, DNAM-1, and TRAIL were even higher on NK cells generated in the presence of SR1. CD62L and CXCR3 are involved in homing to the lymphoid organs and trafficking toward inflammation in vivo, so high expression of these markers can contribute to trafficking of NK cells toward hematological tumor cells in the lymphoid organs [7,34,35]. In addition, it was described that CD62L indicates a unique subset of polyfunctional NK cells combining the ability to produce IFN-γ with cytotoxic properties [40]. DNAM-1 and TRAIL are activation markers, and increased expression levels of these markers suggest that SR1-generated NK cells are more active compared with NK cells generated in the absence of SR1. This was indeed confirmed in functional studies showing that SR1-induced NK cells have increased INF-γ production capacity and an increased capability to kill AML cell lines compared with NK cells generated in the absence of SR1. Furthermore, SR1-generated NK cells show efficient degranulation against AML and MM cell lines. In addition, even at low E:T ratios, these cells efficiently kill hematological tumor cell lines and, most importantly, patient-derived primary AML blasts. To further investigate the applicability of our SR1-NK-cell product after allo-SCT, we investigated the effect of the immunosuppressive drugs MPA and CsA, which are commonly used after nonmyeloablative allo-SCT, on our SR1-NK-cell product. Importantly, we found that therapeutic CsA concentrations do not affect HSPC-NK-cell viability, proliferation, or function, while exposure to therapeutic levels of MPA did have a negative effect. Exposure to CsA did even enhance IFN-γ production by the HSPC-NK cells. This is in accordance with the effect of CsA and MPA on naturally occurring NK cells described in the literature [41,42]. These results provide strong rationale to infuse SR1-generated HSPC-NK cells relatively short after allo-SCT, since MPA treatment is generally stopped at day 28 after allo-SCT. CsA, which is usually prescribed for at least 6 months, will most likely not affect NK-cell viability, proliferation, or function in vivo upon transfer.

Several methods to generate NK-cell products from donor origin for immunotherapy have been reported [43 –48]. In most studies, NK cells are isolated from apheresis products using magnetic cell sorting systems. However, poor recovery and viability are common problems in these procedures; therefore, NK cells require cytokine stimulation and expansion before infusion [2,46 –50]. NK-cell numbers up to 7.6 × 10⁸ have been reported using this method [49]. Nevertheless, large-scale apheresis procedures are necessary to obtain this amount of NK cells, and contaminating T cells can potentially cause GVHD. To circumvent these problems, NK-cell generation from CD34⁺ HSPCs is an attractive option. Yoon et al. recently reported safe infusion of NK cells generated using a feeder-free 6-week culture procedure after HLA-mismatched allo-SCT. Using this procedure, an average number of 9.28 × 10⁶ NK cells/kg was generated from 2.22 × 10⁶ donor CD34⁺ HSPCs/kg [45]. In our system, the average NK-cell expansion from CD34⁺ HSPCs was 235-fold (range, 115- to 904-fold). Therefore, if we use 10% of the G-CSF-mobilized CD34⁺ HSPCs isolated from donors for allo-SCT, we will have on average 0.5 × 10⁶ CD34⁺ cells/kg. In case of a 70 kg patient, we will get on average 8 × 10⁹ NK cells (>1 × 10⁸ NK cells/kg) for infusion. So we will be able to infuse large numbers of NK cells from donor origin using our SR1-based NK-cell generation protocol.

In the present study, we showed that the AhR antagonist SR1 greatly enhances NK-cell expansion and differentiation. AhR is a ligand-inducible transcription factor, which has extensively been studied in the context of its activation by environmental pollutants, such as dioxins and polycyclic aromatic hydrocarbons [51,52]. Until recently, little was known about the physiological role of AhR, but a growing body of evidence shows that AhR has multiple endogenous activators [15] and is involved in several physiological processes [22,53,54]. Examples of endogenous ligands are metabolites of dietary substances [15,55] and tryptophan metabolites like cinnabarinic acid [56], 6-formylindolo[3,2-b]carbazole (FICZ), which can be produced in the skin upon light exposure [57], and the indoleamine 2,3-dioxygenase 1 (IDO1) metabolite kynurenine [58]. AhR also regulates differentiation of various immune cells like Th17 cells [22], dendritic cells [17,18], regulatory T cells [19,20], and γδ T cells [21]. Recently, Hughes et al. reported that antagonism of AhR promotes differentiation of immature innate lymphoid cells into NK cells expressing EOMES and TBET [23]. In our SR1-based HSPC-derived NK-cell culture, we also observed upregulation of EOMES, which is involved in NK-cell maturation [29], and in addition, we found upregulated expression of TOX, ID2, GATA-3, and EAT-2 after AhR inhibition. TOX is important for early NK-cell development [27], ID2 is involved in NK-cell maturation [28], and GATA-3 and EAT-2 are important for NK-cell effector functions [30,31]. Interestingly, expression of TOX markedly decreased in the first week of culture in our culture system in the absence of SR1. In the presence of SR1, TOX expression is more preserved, resulting in significantly higher TOX expression after 1 week. This suggests that before induction of differentiation by addition of IL-15, early NK-cell progenitors are expanded in the presence of SR1, explaining increased NK-cell numbers generated in the presence of SR1, IL-15, and IL-2. To our knowledge, this is the first report of applying AhR blocking in an in vitro system to generate high numbers of functional NK cells.

In conclusion, we developed a cytokine-based ex vivo culture system, which enables us to generate high numbers of very potent NK cells from G-CSF-mobilized CD34⁺ HSPCs. Addition of SR1 to the culture system induced upregulation of multiple NK-cell-related transcription factors and resulted in generation of NK-cell products with very high cell numbers and purity. These NK cells have an active phenotype and are highly functional in vitro; therefore, they hold great promise for future adoptive HSPC-NK-cell therapy after allo-SCT.

Footnotes

Acknowledgments

The authors thank Rob Woestenenk for his help with flow cytometry and the colleagues of the Laboratory of Hematology, Stem Cell Laboratory, for the collection of patient materials. This work was supported by grants from ZonMw (Grant No. 40-41400-98-9018), the Dutch Cancer Society (KWF 2013–6150), and the Radboud University Medical Center.

Prior conference presentation of the submitted material: A part of these data were presented in an oral presentation at the European Society for Blood and Marrow Transplantation meeting, Italy, 2014, and at the Dutch Hematology Congress, the Netherlands, 2014. A part of these data were presented in a poster at the American Society of Hematology meeting, USA, 2014.

Author Disclosure Statement

No competing financial interests exist.

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Supplementary Material

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The Aryl Hydrocarbon Receptor Antagonist StemRegenin1 Improves In Vitro Generation of Highly Functional Natural Killer Cells from CD34 + Hematopoietic Stem and Progenitor Cells

Abstract

Introduction

Materials and Methods

Cell lines

Isolation of CD34+ stem and progenitor cells

Ex vivo expansion of CD34+HSPCs and differentiation into NK cells

RNA isolation and real-time quantitative reverse transcriptase polymerase chain reaction

Flow cytometry

Fluorescence-activated cell sorting of HSCP-NK cells

FCM-based cytotoxicity assays

Enzyme-linked immunosorbent assays

Proliferation assays

Statistical analysis

Results

SR1 enhances expansion of PB- and BM-derived CD34+ HSPCs and improves NK-cell differentiation

SR1 influences expression of transcription factors important for NK-cell differentiation and maturation

SR1-generated HSPC-NK cells have an activated and a mature phenotype

SR1-generated HSPC-NK cells have efficient IFN-γ production capacity and cytolytic activity against AML and MM cells

HSPC-NK-cell proliferation, viability, and function are inhibited by MPA but not by CsA

Discussion

Footnotes

Acknowledgments

Author Disclosure Statement

References

Supplementary Material

Isolation of CD34⁺ stem and progenitor cells

Ex vivo expansion of CD34⁺HSPCs and differentiation into NK cells

SR1 enhances expansion of PB- and BM-derived CD34⁺ HSPCs and improves NK-cell differentiation