Combining AFM13, a Bispecific CD30/CD16 Antibody, with Cytokine-Activated Blood and Cord Blood–Derived NK Cells Facilitates CAR-like Responses Against CD30⁺ Malignancies

Cancer progression is associated with an ineffective antitumor response, mediated by multiple mechanisms, including downregulation of HLA antigens (1), upregulation of checkpoints, and secretion of suppressive cytokines (2–4). Therefore, promising immunotherapy strategies have been developed to enhance the activity of T and natural killer (NK) cells against cancer targets, including the use of cytokines, checkpoint inhibitors, genetic engineering of cells to express a chimeric antigen receptor, or the use of bispecific antibodies (5–12).

NK cells are cytotoxic innate lymphoid cells that play an important role in immunosurveillance and display potent effector responses against infected, stressed, and malignant cells (13). Unlike T cells, NK cells do not express antigen-specific receptors to recognize their target; instead, their activation and effector functions are mediated either by interactions between germline encoded activating or inhibitory receptors and their respective ligands on target cells (14, 15), including CD16A (FcγRIIIa)-mediated antibody-dependent cellular cytotoxicity (ADCC) and cytokine production (16, 17). NK-cell functionality and activation thresholds are also modulated via cytokine receptors (18).

Deficiency in NK number and function is associated with cancer development (19). NK cells from patients with cancer have impaired effector function as shown in multiple disease settings (2, 10, 20). Mechanisms include downregulation of activating receptors, loss of CD16 expression, upregulation of inhibitory receptors, and induction of exhaustion by the tumor microenvironment (21–23). For these reasons, many investigators have explored the use of allogeneic NK cells for immunotherapy and have shown these cells to be safe and effective at killing cancers, especially myeloid malignancies (6, 24, 25).

Allogeneic NK cells can be derived from multiple sources, each with their unique biology (5, 26). Some of the most commonly used sources of allogeneic NK cells currently being investigated in the clinic include peripheral blood (PB) from healthy donors (HD) and cord blood (CB; refs. 26, 27). PB-NK cells may be used with minimal cytokine activation (24), following expansion ex vivo in the presence of cytokines with or without feeder cells (28, 29), or can be briefly activated with IL12, IL15, and IL18 to induce differentiation into memory-like (ML) NK cells (30), with enhanced responses when stimulated through cytokine or activating receptors, including CD16 (31, 32). This is a promising strategy that was shown to be safe and induced remissions in patients with high-risk acute myeloid leukemia (AML; refs. 6, 12). CB-derived NK cells are usually expanded ex vivo to generate sufficient number of cells for infusion (27–29). A novel approach with CAR19/IL15-engineered CB-NK cells was recently reported as safe and able to induce complete remission in patients with relapsed or refractory B-cell malignancies (33).

Immune cell engagers have been developed to overcome barriers related to deficient cancer recognition and NK-cell activation. AFM13 is a tetravalent bispecific antibody construct with two binding sites for each CD30 and CD16A, designed for the treatment of CD30⁺ malignancies. It binds CD16A on the surface of NK cells, thus activating and recruiting them to CD30-expressing tumor cells to mediate tumor cell killing (34, 35). Here, we investigate the use of AFM13 to endow NK cells with anti-CD30 specificity coupled with a high-affinity CD16A trigger, analyzing single NK-cell responses via mass cytometry. Furthermore, we evaluate combining AFM13 with cytokine-induced ML differentiation with both adult PB-NK cells and with expanded CB-NK cells, to identify promising strategies to translate into the clinic for CD30⁺ leukemias and lymphomas.

Human PB mononuclear cells (PBMC) from anonymous HDs were obtained from leukoreduction filters after platelet apheresis and isolated by Ficoll density gradient centrifugation. CB units for research were obtained from the MD Anderson Cancer Center Cord Blood Bank (Houston, TX). PBMC from patients with Hodgkin lymphoma (HL) were collected on an Institutional Review Board–approved clinical protocol (Lab00-099). CB mononuclear cells were isolated from fresh CB units by Ficoll density gradient centrifugation. The protocol for the ex vivo expansion of CB-NK cells using K562 feeder cells was published previously (7, 33, 36). Cytokine-induced ML NK cells were generated from conventional peripheral blood NK cells (cNK) utilizing activation (16 hours) with rhIL12, rhIL15, and rhIL18 as described previously (6). After the activation, NK cells were washed and cultured for 6 days to allow ML differentiation. Preactivated expanded (P+E) CB-NK cells were generated following the same pre-activation protocol with IL12, IL15, and IL18 prior to expansion with universal antigen presenting cells (uAPC) and IL2 as above.

The tumor cell lines Karpas 299 and HuT-78 cells were chosen for this study based on their high expression of CD30. Karpas 299 was purchased from Sigma-Aldrich. Daudi, K562, and Raji cells were obtained from ATCC. The HuT-78 cell line was provided by John DiPersio (Washington University, St. Louis, MO). The cell lines were cultured in RPMI medium supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% l-glutamine. Karpas 299 and Daudi cells were used as targets for the cytotoxicity assays. All cells lines were maintained according to ATCC instructions and were Mycoplasma negative.

Details of the flow cytometry to measure cytokine production and degranulation, ⁵¹Chromium (⁵¹Cr)-release assay, and Incucyte real-time assay are described in details in the Supplementary Materials and Methods.

For immunofluorescence detection of AFM13 on the surface of NK cells, CB-NK cells were loaded with AFM13 for 1 hour, washed, and cultured for another 24 to 72 hours at 37°C in the presence of IL2, harvested, and stained with rat anti-AFM13 (clone 7, Affimed), followed by staining with goat anti-rat IgG (Alexa Fluor 647, Jackson ImmunoResearch). Images were obtained using imaging flow cytometry (Amnis Image Stream Mark II) and analyzed by the Ideas software (Luminex). The retention of AFM13 on the surface of NK cells was determined by the colocalization of CD16 and AFM13 signals. Imaging flow cytometry (Amnis Image Stream Mark II) was also used to assess immunologic synapse formation between AFM13-loaded CB-NK cells and Karpas 299 by staining the cells with a biotinyalated anti-pericentrin primary antibody (Novusbio) followed by streptavidin-conjugated APC-Cy7 (BioLegend) and AF 488-conjugated phalloidin (Thermo Fisher Scientific) to measure F-actin. Images were analyzed with the IDEAS software (Miltenyi Biotech).

Total RNA was collected from 5 million expanded CB-NK cells (with or without pre-activation) and extracted and purified using RNeasy Plus Mini Kit (Qiagen). SmartSeq2 RNA sequencing (RNA-seq) libraries were prepared and data analyzed as described in the Supplementary Materials and Methods.

Supplementary Tables S1 and S2 list antibodies used for the characterization of PB-NK cells and CB-NK cells, respectively, and custom conjugations were described previously (6, 37). Mass cytometry data were collected on a CyTOF2 mass cytometer (Fluidigm) and analyzed using Cytobank as described previously (6, 38–40). Detailed information regarding staining, acquisition, and analysis is provided in Supplementary Materials and Methods.

NSG mice (10–12 weeks old; Jackson Laboratories) were irradiated with 300 cGy and inoculated intravenously with FFLuc-labeled Karpas cells (1 × 10⁵) on day 0. Treatment groups received 10 × 10⁶ CB-NK cells or CB-NK cells loaded with AFM13 or 100 μg/mL of AFM13 alone through tail vein injection on day 0. Mice received recombinant human IL2 injection intraperitoneally three times per week (10,000 units/mouse). Mice were subjected to weekly bioluminescence imaging (BLI; Xenogen-IVIS 200 Imaging system; Caliper).

The Student t test was used to compare quantitative variables. Two-way ANOVA was used for comparison between samples and longitudinal timepoints followed by Bonferroni or Sidak multiple comparisons. Probabilities of survival were calculated using the Kaplan–Meier method. Statistical significance was assessed with the Prism 8.0 software (GraphPad Software, Inc.). P values <0.05 were considered significant.

Data may be shared after request to the corresponding authors by e-mail: Katayoun Rezvani: krezvani@mdanderson.org and Todd A. Fehniger: tfehnige@wustl.edu.

The bispecific nature of AFM13 supports activation of NK cells through its CD16-binding domain along with an enhanced tumor recognition mediated by CD30 binding on the tumor cells. However, the quality of single NK-cell responses within a heterogeneous NK-cell population to respond to CD16 engagement with AFM13 remains unclear. To confirm the effect of AFM13 in enhancing tumor recognition and NK-cell functionality (34, 35, 41), cNK from HD PBMC were incubated with HuT-78 (CD30⁺) with or without AFM13 labeling. Flow cytometry confirmed a specific and robust increase in IFNγ, TNF, and surface CD107-positive NK cells with the HuT-78 plus AFM13 condition (Supplementary Fig. S1A–S1D). Control stimulation with Raji cells (CD30⁻) or Raji plus AFM13 did not induce significant responses by cNK cells (Supplementary Fig. S1A–S1D). We hypothesized that the underlying NK-cell repertoire and subsets affect AFM13-triggered responses. To define the multidimensional profiles of cNK cells responding to AFM13, mass cytometry was employed to assess the expression of phenotypic and function-relevant NK-cell markers modulated upon AFM13 triggering (Supplementary Table S1). AFM12 (binding to CD16A and CD19) was used as negative control, as HuT-78 cells do not express CD19 (Fig. 1A). Density viSNE maps (Fig. 1B) demonstrate that HuT-78+AFM13 stimulation induced a unique multidimensional profile, which was markedly altered compared with control conditions (unstimulated, HuT-78, or HuT-78+AFM12). The composite viSNE plot highlights these differences, where HuT-78+AFM13 stimulated cells are predominantly located in a distinct island (Fig. 1B) and serves as a reference for the investigation of function (Fig. 1C). Functional mass cytometry analysis showed that HuT-78+AFM13 significantly induced IFNγ, TNF, MIP1α, and surface CD107a (as a surrogate of degranulation) in cNK cells (Fig. 1C and D). As expected, AFM12 did not upregulate cytokine secretion or degranulation (Fig. 1D), indicating that enhanced NK-cell functionality upon AFM13 triggering is related to simultaneous binding of both CD16A on NK cells and CD30 on target cells, rather than CD16A binding alone on the NK cells. The short-term functional changes induced by AFM13-triggering correlated with activation-related decreases of CD16A and CD62L and upregulation of CD25 and more modestly, NKp44 (Fig. 1E). We also investigated the impact of AFM13 on the quality of the NK-cell response measured via polyfunction (combinations of IFNα, TNF, MIP1α, and CD107a). HuT-78+AFM13 was found to upregulate polyfunctional responses in cNK cells while HuT-78+AFM12 activated NK cells with single functional features (Fig. 1F). Thus, mass cytometry functional assays identify highly activated NK-cell islands upon AFM13 triggering, and an increase in polyfunctional NK cells, compared with control conditions.

To examine the phenotype of NK cells responding to AFM13, SPADE (Spanning-tree Progression Analysis of Density-normalized Events) analysis was employed with clustering of 17 markers associated with NK-cell maturation and differentiation (Fig. 2A; Supplementary Table S1). Visualization trees from SPADE analysis were used to identify nodes with higher and lower IFNγ median expression in response to AFM13 within individual NK-cell donors (Fig. 2B–D). Within the IFNγ^high nodes, we identified mature (KIR⁺CD57⁺) and immature NK cells (NKG2A/CD94⁺) responding to AFM13 triggering as demonstrated in three representative donors. In one donor, expression of KIR3DL1 marked NK cells with the highest production of IFNγ (node 1; Fig. 2B), but minimal KIR2DL2/3. Because HuT-78 lack the HLA-Bw4 ligand for KIR3DL1, this result suggests that licensed KIR3DL1⁺ NK cells that lack their inhibitory ligand (HuT-78 lack HLA-Bw4 motifs) exhibited the strongest response when triggered with AFM13 in this individual. In addition, node 8 in this donor and nodes 4 and 10 in a second donor (Fig. 2C) also exhibited a robust IFNγ expression and expressed markers of terminally matured NK cells (CD57⁺KIR2DL2/3⁺NKG2A⁻). In a different individual (Fig. 2D), nodes with the greatest IFNγ production (2 and 3), expressed CD57, CD94, NKG2A, and variable KIR3DL2/3. This demonstrates that both immature and mature NK cells responded robustly to AFM13 triggering. CD16 expression was found to be similar in both CD56^dim mature (NKG2A⁻CD57⁺) and CD56^dim immature (NKG2A⁺CD94⁺KIR⁻CD57⁻) NK cells and both subsets were equally triggered by AFM13 to produce similar levels of IFNγ (Supplementary Fig. S2). When analyzing cNK cells from all individuals tested in mass cytometry functional assays, IFNγ^high nodes had additional functional markers (e.g., MIP1α, CD107a degranulation; Fig. 2E), and were enriched for mature (CD57⁺KIR⁺) NK cells, compared with IFNγ^low NK cells (Fig. 2F). Thus, blood cNK cells exhibited heterogeneous responses within and across multiple individuals; advanced NK-cell maturation state and potentially loss of iKIR ligands were associated with AFM13-triggered responses. In addition, the correlative evidence of KIR and HLA-Bw4 interaction supports a role for specific inhibitory receptor and ligand interactions impacting AFM13-triggered responses in vitro. This will be further evaluated in future studies that manipulate inhibitory receptors or their ligands within functional assays.

As blood cNK cells exhibited varied AFM13-triggered responses within different NK-cell maturation states and iKIR-defined subsets, several types of NK cells were evaluated for response to AFM13. These included HD and HL patient PB cNK cells, as well as cord-blood expanded (CB-NK; ref. 27) and PB cytokine-induced ML (30) NK cells, which are currently being tested in cellular therapy clinical trials for hematologic malignancies. AFM13 significantly augmented PB cNK cytotoxicity against Karpas 299 lymphoma in HD and to a lesser extent PB cNK from patients with relapsed/refractory HL who received multiple treatments including brentuximab. In contrast, AFM13-triggered CB-NK cells exhibited only a modest trend for increased cytotoxicity (Fig. 3A). Brief activation of PB-NK cells with IL12, IL15, and IL18 results in ML differentiation and improves cytokine production and killing of hematologic cancer targets (6, 30). We hypothesized that IL12/15/18 activation and consequent ML differentiation (Supplementary Fig. S3A and S3B) may positively impact AFM13-triggered responses. In a second set of experiments, PB cNK cells were compared with PB ML NK cells, with and without AFM13, for killing of CD30⁺ HuT-78 lymphoma targets. While cNK cells again demonstrated improved killing with AFM13 triggering, ML NK cells exhibited significantly enhanced killing compared with cNK cells (Fig. 3B; Supplementary Fig. S4A), consistent with previous studies showing their enhanced ADCC responses (31). We next evaluated the impact of IL12/15/18 activation followed by expansion of CB-NK cells on AFM13-triggered Karpas 299 cell killing (Fig. 3C). At all concentrations of AFM13 tested, IL12/15/18 P+E CB-NK cells exhibited enhanced killing over control expanded CB-NK cells. Addition of AFM13 did not increase the cytotoxicity of PB cNK cells or P+E CB-NK cells against CD30⁻ targets (Supplementary Fig. S4B and S4C). These data indicate that activation with IL12/15/18 followed by ML differentiation of PB-NK cells or expansion of CB-NK cells enhanced AFM13-triggered killing. In addition, PB ML NK cells (Fig. 3D) and IL12/15/18 P+E CB-NK cells (Fig. 3E) also produced IFNγ, TNF, and degranulated (surface CD107a) effectively to CD30⁺ targets.

Because IL12/15/18 pre-activation has not previously been reported with CB-NK cell expansion, we next evaluated potential mechanisms responsible for their enhanced functionality. Flow cytometry analysis of P+E CB-NK cells compared with control expanded CB-NK cells showed similar CD16 expression (Supplementary Fig. S5), indicating that CD16 expression changes were not the underlying mechanism for the enhanced AFM13 response. RNA-seq of purified CB-NK cells was performed comparing the transcriptome profiles of P+E CB-NK versus expanded CB-NK cells after 14 days of culture (Fig. 4A). P+E CB-NK cells had higher expression of select chemokines and cytokine genes, such as CCL3, CCL4, CCR5, and TNF (Fig. 4B and C). Gene set enrichment analysis (GSEA) confirmed enrichment of genes involved in the IFNγ response, TNF signaling, IL2/STAT5 signaling, IL6/JAK/STAT3 signaling, and mTOR pathway as well as genes related to inflammatory immune responses (Fig. 4D and E). Taken together, these data suggest that pre-activation of CB-NK cells with IL12, IL15, and IL18 prior to expansion results in increased signaling pathways related to NK-cell effector function, indicating a potential mechanism of enhanced AFM13 response.

Mass cytometry was used to investigate if AFM13 can induce changes in the phenotype of P+E CB-NK cells using a second panel (Supplementary Table S2). P+E CB-NK cells were cocultured with AFM13 and tumor targets for 4 hours and then analyzed using CyTOF (Supplementary Fig. S6). viSNE-based analysis revealed segregation in the cell clusters of P+E CB-NK cells treated with AFM13, with or without tumor targets (Supplementary Fig. S6A and S6B). Cells loaded with AFM13 had evidence of activation (increased expression of TRAIL, NKp44, and CD69; Supplementary Fig. S6C). These data are consistent with the notion that through CD16 engagement, AFM13 can activate NK cells as it was previously shown in PB-NK cells (41). No significant differences in NK cell–associated markers other than the ones described above were observed in the studied conditions (Supplementary Table S2).

When combining AFM13 with donor NK-cell therapy, one approach to maximize initial NK-cell triggering is preloading effector cells with AFM13 prior to infusion. To address this, we investigated the feasibility and activity of precomplexed AFM13-NK cells by testing the stability of AFM13 binding on P+E CB-NK cells over time. To test this, NK cells were incubated with different concentrations of AFM13 (1–100 μg/mL) for 1 hour and then washed twice with PBS prior to performing functional studies. Regardless of the AFM13 concentration used, AFM13-loaded and washed NK cells were as efficient as loaded and unwashed NK cells at killing Karpas 299 targets (Fig. 5A). Furthermore, IncuCyte monitoring of NK-cell cytotoxicity over 24 hours revealed similar killing kinetics of AFM13-loaded and washed and AFM13-loaded and unwashed NK cells against Karpas 299 targets (Fig. 5B). Thus, AFM13 binding to the surface of P+E CB-NK cells is stable and retains activity to enhance tumor recognition and killing.

Retention of AFM13 on the surface of NK cells over time was evaluated using a mAb directed against AFM13. Multiparameter flow cytometry showed that NK cells loaded with AFM13 (100 μg/mL) for 1 hour and then washed retained AFM13 on their surface for least 72 hours in culture (Fig. 5C). As expected, AFM13 was detected mainly on CD16⁺ NK cells (Fig. 5C). These results were confirmed using an imaging flow cytometer where we found that AFM13 was colocalized with CD16 on the surface of NK cells (Fig. 5D). Interestingly, AFM13 was partially internalized from the cell membrane into the cytoplasm (Fig. 5D and E). However, the retention of AFM13 on the surface of the NK cell remained the same over time, depicting thereby that 1-hour incubation of AFM13 with NK cells (Fig. 5A) could have the same effect as incubating AFM13 with NK cells either for 24 hours or 48 hours. Nonetheless, to test whether AFM13 internalization affects the ability of NK cells to kill tumor targets, NK cells were loaded with AFM13, washed, and incubated for 1 hour, 48 hours, or 72 hours in media plus rhIL2, with unwashed cells as control. All AFM13-loaded groups (washed and unwashed) exhibited significantly higher cytotoxicity against Karpas 299 lymphoma targets at all timepoints compared with unloaded P+E CB-NK cells (Fig. 5F), suggesting that partial internalization of AFM13 does not affect the enhanced cytotoxic activity of NK cells. To provide additional mechanistic data to support the importance of combined CD16/AFM13 engagement in mediating the antitumor response, we used imaging flow cytometry to confirm significantly higher colocalization of CD16 at the immunologic synapse between CB-NK cells and Karpas cells in the presence of AFM13 compared with a control IgG antibody (Fig. 5G and H). In addition, we confirmed that in the absence of NK cells, the addition of the CD30 bispecific engager failed to induce apoptosis of the Karpas tumor cells as demonstrated in Supplementary Fig. S7.

To initially assess the safety of P+E CB-NK preloaded with AFM13 in vivo, immunodeficient NSG mice were injected intravenously with 10 × 10⁶ AFM13-loaded (and washed as in Fig. 6) P+E CB-NK cells (AFM13-NK) and monitored for toxicity, including daily weight measurements for more than 10 months. We did not find any evidence of toxicity during the follow-up or on histologic examination at necropsy (Supplementary Fig. S8A and S8B). Next, the antitumor activity of 10 × 10⁶ AFM13-complexed NK cells in lymphoma bearing mice was tested (Fig. 6A). Mice received (i) Karpas 299 cells alone, (ii) Karpas 299 cells plus unloaded P+E CB-NK cells (10 × 10⁶), (iii) Karpas 299 cells plus AFM13-complexed NK cells, or (iv) Karpas 299 cells + 100 μg/mL of AFM13 (intravenously without NK cells). All mice received human IL2 (10,000 units/mouse) intraperitoneally on the day of NK-cell injection and every 2–3 days to support human NK-cell survival in NSG mice. Compared with control mice, infusion of the AFM13–NK complex improved tumor control measured via BLI (Fig. 6B) and survival (Fig. 6C). No differences in mouse weight was observed between the different mouse conditions, suggesting minimal cytokine release and no graft versus host disease (Fig. 6D). Collectively, these data support that the AFM13-complexed NK cells are capable of safely controlling tumor cells in vivo.

In this study, we have shown that AFM13, a tetravalent bispecific antibody directed against CD16A and CD30, potentiate NK-cell cytotoxicity and cytokine production of IL12/15/18-induced ML mature blood NK cells and preactivated and expanded CB-NK cells against CD30-expressing lymphoma targets both in vitro and in vivo. Multidimensional analysis of PB cNK cells triggered by AFM13 plus CD30⁺ lymphoma targets revealed responses that were polyfunctional, and exhibited a heterogeneity of response at the single cell and individual donor levels. However, a consistent pattern of response spanning analysis of all donors revealed the greatest responses occurring in mature PB cNK cells.

Comparisons of distinct types of NK cells revealed a source-dependent response profile to AFM13-based stimulation. While cNK blood NK cells exhibited enhanced cytotoxicity in response to AFM13 alone (41) or AFM13-dependent target recognition, this appeared decreased in cNK cells from patients with HL in vitro. Furthermore, the established IL12, IL15, and IL18-induced ML NK-cell differentiation of PB-NK cells resulted in significantly enhanced AFM13-dependent killing. This finding was also evident upon IL12/15/18 activation followed by expansion of CB-NK cells, with enhancement of AFM13-triggered cytotoxicity. Thus, this is the first study that identifies immunotherapeutic strategies that combine activation via the IL12, IL15, and IL18 receptors of two NK-cell types with AFM13 triggering, resulting in augmented NK-cell responses to CD30⁺ cancers preclinically.

Prior work has defined the induction of NK cells with ML properties following combined cytokine activation. Cytokine-induced ML responses were identified using a murine adoptive transfer system, where IL12, IL15, and IL18-activated splenic NK cells differentiated in syngeneic mice, and produced increased IFNγ in response to cytokine restimulation (42). Human PB-NK cells briefly activated with IL12, IL15, and IL18 and allowed to differentiate in vitro exhibited enhanced function when restimulated with cytokines, with activating receptors or with tumor targets (30), and may also be engineered with a CAR (11). Multiple preclinical studies have shown that human and murine NK cells exhibit enhanced anticancer properties of ML NK cells (6, 30, 31, 43–45). PB ML NK cellular therapy was shown to safely induce complete remissions in 47% of evaluable patients with relapsed/refractory AML with expansion and persistence of donor ML NK cells in the blood and bone marrow of patients for at least 21 days after infusion (6, 12). Our finding that CB-NK cells preactivated with IL12, IL15, and IL18 prior to expansion had enhanced AFM13-mediated killing could be explained by the lack of complete maturation of expanded CB-NK cells (46). Furthermore, while IL2 can drive the proliferation of CB-NK cells, IL15 was necessary for the cells to acquire full effector function (37). Here, RNA-seq data demonstrated that activation with IL12/15/18 prior to expansion of CB-NK cells induces a phenotype distinct from expanded CB-NK cells, and resemble aspects of cytokine-induced ML NK cells (6). RNA changes included upregulation of IFNγ and TNF signaling as well as the STAT5 and mTOR pathways. Future studies will focus on unraveling molecular mechanisms that result in ML NK-cell programs by various NK-cell types.

Multiple studies have shown that NK cells from patients with cancer, including patients with HL, have impaired cytotoxic function (35, 47, 48). Indeed, we confirmed upregulation of multiple inhibitory receptors, including PD-1, TIM 3, TIGIT, and Pan-KIR, and lower expression of Eomes on HL NK cells compared with PB-NK cells. In contrast, CD16 levels were equivalent in NK cells from patients with HL and HD (Supplementary Fig. S9), suggesting that the functional deficit observed in patients with HL is predominantly related to expression of multiple checkpoints, rather than to lower AFM13-binding to NK cells secondary to CD16 downregulation. This may in part explain the results of a phase I dose-escalation study of AFM13, where only 3 of 26 evaluable patients with heavily pretreated HL achieved partial remission (11.5%; ref. 49). These data support the notion that AFM13 will be more effective when combined with allogeneic NK cells from HDs. One approach supported by our study is to precomplex the NK cells with AFM13 ex vivo and to infuse the complexed product to the patient rather than to infuse the AFM13 and NK cells as two separate products. This strategy may prevent the in vivo dilution and uptake of AFM13 by the recipient endogenous NK cells, which are likely to have low cytotoxic activity. We have preactivated, expanded, and precomplexed CB-NK cells with AFM13 and confirmed their cytotoxicity against CD30⁺ tumors both in vitro and in vivo. AFM13-complexed NK cells retain the engager on their surface for at least 3 days, which may provide sufficient time for the AFM13-complexed NK cells to engage and kill targets, as observed in our preclinical mouse model. CB-derived and PB ML NK cells are a ready source of cells to combine with AFM13, that have the potential for broad availability. Furthermore, we have shown that CB-derived NK cells can be safely used in clinics even in the absence of HLA matching (33, 50).

Our approach of loading NK cells with an anti-CD16 bispecific antibody can be extended to other targets, transforming the precomplexed NK cells into de facto CAR-NK cells, thus providing a rapid approach to translate NK cells with CAR-like characteristics to the clinic. While antigen escape, including antigen loss or downregulation, may also occur after AFM13-complexed NK-cell immunotherapy, the capacity of NK cells to recognize and target tumor cells through their endogenous receptors may make disease escape less likely.

In conclusion, we have developed a novel approach to immunotherapy using NK cells complexed with AFM13. A clinical trial with off-the-shelf AFM13-complexed preactivated and expanded CB-NK cells in patients with relapsed/refractory CD30⁺ malignancies is underway and will elucidate if the promising in vitro and in vivo observations are reproducible in patients (Clinicaltrial.gov NCT04074746).

L.N. Kerbauy reports other support from Affimed during the conduct of the study, as well as personal fees from Takeda outside the submitted work. P.P. Banerjee reports other support from Takeda Pharmaceutical during the conduct of the study. M.M. Berrien-Elliott reports personal fees from Wugen outside the submitted work; in addition, M.M. Berrien-Elliott has a patent for 017001-PRO1 pending, licensed, and with royalties paid from Wugen. M. Becker-Hapak reports a patent for US20100159594A1 issued, licensed, and with royalties paid from Washington University. R. Basar reports personal fees from Takeda during the conduct of the study, as well as personal fees from Affimed outside the submitted work. M. Daher reports personal fees from Takeda outside the submitted work. E. Liu reports other support from Takeda and Affimed during the conduct of the study. S.O. Ang reports other support from Takeda and Affimed during the conduct of the study. R.E. Champlin reports grants from Affimed during the conduct of the study. Y.L. Nieto reports grants from AstraZeneca, BioSecura, and Affimed outside the submitted work. J. Koch is an employee of Affimed. W. Fischer is an employee of Affimed. O.K. Okamoto reports grants from FAPESP 2013/08028-1 and CNPq 307611/2018-3 during the conduct of the study; O.K. Okamoto is a visiting scholar at HIAE. E.J. Shpall reports personal fees and other support from Bayer HealthCare Pharmaceuticals, Novartis, and Magenta, as well as other support from Adaptimmune, Mesoblast, and Axio outside the submitted work; in addition, E.J. Shpall has a patent for Takeda licensed and a patent for Affimed licensed. T.A. Fehniger reports grants from Affimed during the conduct of the study. T.A. Fehniger also reports personal fees from Nektar; grants, personal fees, and other support from Wugen; other support from Indpata and Orca Bio; personal fees and other support from Kiadis; grants from Affimed; and grants and other support from Compass Therapeutics outside the submitted work. In addition, T.A. Fehniger has a patent for 15/983,275 pending and licensed to Wugen, a patent for PCT/US2019/060005 pending and licensed to Wugen, and a patent for 62/963,971 pending and licensed to Wugen. K. Rezvani reports other support from Affimed during the conduct of the study, as well as other support from Takeda Pharmaceutical Company outside the submitted work; K. Rezvani participates on the scientific advisory board for GemoAb, AvengeBio, Virogin, GSK, and Bayer. No disclosures were reported by the other authors.

L.N. Kerbauy: Data curation, formal analysis, investigation, writing–original draft, writing–review and editing. N.D. Marin: Data curation, formal analysis, investigation, writing–original draft, writing–review and editing. M. Kaplan: Data curation, formal analysis, investigation. P.P. Banerjee: Data curation, formal analysis, investigation. M.M. Berrien-Elliott: Data curation, formal analysis, investigation. M. Becker-Hapak: Data curation, formal analysis, investigation. R. Basar: Investigation. M. Foster: Investigation. L. Garcia Melo: Investigation. C.C. Neal: Investigation. E. McClain: Investigation. M. Daher: Investigation. A.K. Nunez Cortes: Investigation. S. Desai: Investigation. F.W. Inng Lim: Methodology, writing–review and editing. M.C. Mendt: Methodology, writing–review and editing. T. Schappe: Investigation. L. Li: Formal analysis, writing–review and editing. H. Shaim: Formal analysis, writing–review and editing. M. Shanley: Formal analysis, writing–review and editing. E.L. Ensley: Formal analysis, investigation. N. Uprety: Data curation, investigation. P. Wong: Investigation. E. Liu: Formal analysis, writing–review and editing. S.O. Ang: Formal analysis, writing–review and editing. R. Cai: Formal analysis, writing–review and editing. V. Nandivada: Investigation. V. Mohanty: Data curation, formal analysis, investigation. Q. Miao: Data curation, formal analysis, investigation. Y. Shen: Data curation, formal analysis, investigation. N. Baran: Investigation. N.W. Fowlkes: Investigation. K. Chen: Methodology, writing–review and editing. L. Muniz-Feliciano: Writing–original draft, writing–review and editing. R.E. Champlin: Conceptualization, writing–review and editing. Y.L. Nieto: Conceptualization, writing–review and editing. J. Koch: Data curation, formal analysis, investigation. M. Treder: Data curation, formal analysis, investigation. W. Fischer: Data curation, formal analysis, investigation. O.K. Okamoto: Conceptualization, writing–review and editing. E.J. Shpall: Conceptualization, writing–review and editing. T.A. Fehniger: Conceptualization, data curation, formal analysis, supervision, funding acquisition, writing–original draft, project administration, writing–review and editing. K. Rezvani: Conceptualization, data curation, formal analysis, supervision, funding acquisition, writing–original draft, project administration, writing–review and editing.

We thank the RNA sequencing core facility, which is supported by the Cancer Center Support (CORE), and the Research Animal Support Facility at MD Anderson Cancer Center. Figures created with biorender.com.

This work is supported by grants from the NIH (1 R01 CA211044-01, 5 P01CA148600-03, and P50CA100632-16). The MD Anderson Cancer Center Flow Cytometry and Cellular Imaging Facility and the RNA sequencing core facility is supported in part by a grant from the NIH, NCI (CA016672). The Siteman Flow Cytometry Core and the Immune Monitoring Laboratory of the Siteman Cancer Center in Washington University, School of Medicine, St. Louis, MO are supported by the NCI CCC support (P30CA091842). Additional support for the study includes NIH: K12CA167540 (M.M. Berrien-Elliott), SPORE in Leukemia P50CA171963 (T.A. Fehniger, M.M. Berrien-Elliott), and R01CA205239 (T.A. Fehniger). This study was partially supported by Affimed.

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