The family of PIM serine/threonine kinases includes three highly conserved oncogenes, PIM1, PIM2, and PIM3, which regulate multiple prosurvival pathways and cooperate with other oncogenes such as MYC. Recent genomic CRISPR-Cas9 screens further highlighted oncogenic functions of PIMs in diffuse large B-cell lymphoma (DLBCL) cells, justifying the development of small-molecule PIM inhibitors and therapeutic targeting of PIM kinases in lymphomas. However, detailed consequences of PIM inhibition in DLBCL remain undefined. Using chemical and genetic PIM blockade, we comprehensively characterized PIM kinase–associated prosurvival functions in DLBCL and the mechanisms of PIM inhibition–induced toxicity. Treatment of DLBCL cells with SEL24/MEN1703, a pan-PIM inhibitor in clinical development, decreased BAD phosphorylation and cap-dependent protein translation, reduced MCL1 expression, and induced apoptosis. PIM kinases were tightly coexpressed with MYC in diagnostic DLBCL biopsies, and PIM inhibition in cell lines and patient-derived primary lymphoma cells decreased MYC levels as well as expression of multiple MYC-dependent genes, including PLK1. Chemical and genetic PIM inhibition upregulated surface CD20 levels in an MYC-dependent fashion. Consistently, MEN1703 and other clinically available pan-PIM inhibitors synergized with the anti-CD20 monoclonal antibody rituximab in vitro, increasing complement-dependent cytotoxicity and antibody-mediated phagocytosis. Combined treatment with PIM inhibitor and rituximab suppressed tumor growth in lymphoma xenografts more efficiently than either drug alone. Taken together, these results show that targeting PIM in DLBCL exhibits pleiotropic effects that combine direct cytotoxicity with potentiated susceptibility to anti-CD20 antibodies, justifying further clinical development of such combinatorial strategies.

Significance:

These findings demonstrate that inhibition of PIM induces DLBCL cell death via MYC-dependent and -independent mechanisms and enhances the therapeutic response to anti-CD20 antibodies by increasing CD20 expression.

Diffuse large B-cell lymphoma (DLBCL) is the most common type of aggressive B-cell lymphoma in adults. DLBCL exhibits highly heterogeneous clinical behavior and a complex molecular background. Depending on their transcriptomic profiles, DLBCLs can be classified into distinct categories: germinal center –like (GCB) and activated B-cell–like (ABC) subtypes, that differ also in clinical behavior (1, 2). However, recent studies highlight far more complex molecular background with multiple low-frequency mutations, somatic copy-number alterations, and structural variants (3–6). Despite such striking molecular heterogeneity, the combination of anti-CD20 monoclonal antibody rituximab with chemotherapy (R-CHOP) remains the standard of care in the first-line DLBCL treatment. However, about one of three of patients is refractory to first-line therapy or relapses after initial response, underscoring the need for better treatment modalities.

Recent studies indicate that targeting oncogenic drivers and activated signaling pathways is a promising strategy to improve treatment outcome in DLBCL patients (3, 4, 7). MYC is one of the key oncogenes in DLBCL development. MYC encodes a leucine zipper transcription factor that modulates a broad spectrum of cellular genes responsible for enhancing cell proliferation, cellular metabolism, growth, angiogenesis, metastasis, genomic instability, stem cell self-renewal, reduced differentiation, and immunogenicity (8). MYC-rearranged tumor cells remain addicted to and reliant on its continuous expression (9), indicating that targeting MYC might be therapeutically beneficial. However, direct MYC-targeting strategies have been largely ineffective, highlighting the need for other, indirect approaches.

The family of PIM serine/threonine kinases includes 3 highly conserved oncogenes (PIM1–3), overexpressed in multiple solid tumors and hematologic malignancies. PIMs modulate the activity of proteins involved in regulation of translation, transcription, cell cycle, metabolism, survival, immune escape (10–15), and amplify the tumor-promoting potential of other oncogenes, such as MYC. Double transgenic Eμ-Pim1/Eμ-Myc mice exhibit highly accelerated lymphomagenesis, with B-cell tumors arising in utero or around birth (16, 17), indicating that Pim kinases are essential Myc partners (18). The PIM–MYC oncogenic partnership is not limited to lymphoid tumors but is also present in solid cancers. For example, PIM inhibition in MYC-addicted triple-negative breast cancer cells arrests the growth of patient-derived tumor xenografts and Myc (mouse)-driven transgenic breast cancer mouse models by blocking MYC expression and inhibiting MYC transcriptional activity (19).

However, prior attempts to target PIMs in the clinic were hampered by the off-target effect associated with unacceptable toxicity profiles of first-generation PIM inhibitors (SGI-1776; ref. 20). Although newer pan-PIM inhibitors AZD1208 and LGH447(PIM447) were well tolerated, they exhibited insufficient on-target activity and limited efficacy when used as single agents in patients with advanced solid cancers or refractory AML. However, the pan-PIM inhibitor PIM447 demonstrated moderate therapeutic efficacy in patients with relapsed/refractory multiple myeloma (21, 22), indicating that identifying the tumor types that are more sensitive to PIM inhibition is critical for a rational planning of PIM inhibitor trials. Moreover, PIM kinase inhibitors sensitize tumor cells to chemo/radiotherapy (23, 24) or other small-molecule kinase inhibitors (25), suggesting that rather than being used as single agents, PIM inhibitors might be considered a backbone of combination therapies.

In this study, we evaluated the activity of a novel pan-PIM inhibitor SEL24/MEN1703 in DLBCL preclinical models. MEN1703 is currently being tested in a clinical trial in patients with AML (DIAMOND-01/CLI24-001 trial, NCT03008187) and exhibits a favorable safety profile. Using chemical and genetic PIM blockade, we comprehensively characterized the PIM kinase-associated prosurvival functions in DLBCL and the mechanisms of PIM inhibitor activity. We characterize here MYC-dependent and -independent consequences of PIM inhibition in DLBCL. Importantly, we demonstrate that PIM inhibition in DLBCL cells increases the expression of CD20 antigen in a MYC-dependent mechanism, and synergizes with anti-CD20 antibody rituximab in a mouse lymphoma xenograft model. Thus, PIM inhibition exhibits pleiotropic effects that combine direct cytotoxicity with increased surface CD20 expression and increased susceptibility to anti-CD20 antibodies—crucial drugs improving DLBCL outcome.

Cell lines, culture conditions, and chemicals

Human DLBCL cell lines DHL4, DHL6, U2932, and Burkitt lymphoma (BL) cell line Raji were maintained in RPMI-1640 medium supplemented with 10% FBS, and DLBCL cell lines LY1, LY7, HBL-1, and B-ALL cell line SEM were grown in IMDM supplemented with 10% FBS. The pan-PIM inhibitor MEN1703 was provided by Menarini Ricerche. Pan-PIM inhibitors SGI-1776 and PIM447, MYC inhibitor 10058-F4, and proteasome inhibitor MG132 were purchased from Selleckchem. All chemicals were dissolved in DMSO (Sigma-Aldrich). In vehicle-control experiments, maximal final DMSO concentration was 0.1% (control for 10 μmol/L dose).

Antibodies and oligonucleotides

Antibodies used in immunoblot, capillary nanoimmunoassay, chromatin immunoprecipitation, and IHC analyses are listed in Supplementary Table S3. Primers used in gene expression and chromatin immunoprecipitation experiments are listed in Supplementary Table S4. siRNAs used in knockout experiments are listed in Supplementary Table S5.

Proliferation, cell death/apoptosis, cell-cycle, and caspase activation assays

Cellular proliferation was measured by the CellTiter96 AQueous nonradioactive cell proliferation assay (MTS, Promega). EC50 values were calculated using GraphPad Prism Version 6 (GraphPad software). All experiments were conducted in triplicates. For assessment of apoptosis, cells were washed in PBS, suspended in 1× Annexin V binding buffer (Becton Dickinson; BD), stained with Annexin V–FITC and propidium iodide (PI) (BD), 7AAD-only (GFP-expressing DHL4 MYC-mutant T58A cells), or PI-only [cells investigated in complement-dependent cytotoxicity (CDC) and antibody-dependent cellular cytotoxicity (ADCC) assays] and analyzed by flow cytometry using Cytoflex flow cytometer (Beckman Coulter). Caspase activity was assessed using Caspase-Glo kits (Promega, cat. no. G8090, G8200, and G8210), according to the manufacturer's instructions. Briefly, DLBCL cells were incubated with 1.5 μmol/L MEN1703 or DMSO for 24 hours. Thereafter, 2.5 × 104 of cells were mixed with Caspase-Glo substrate, placed into 96-well BRAND microplate (Sigma-Aldrich), and the luminescence was measured after 90 minutes using Tristar LB 941 microplate reader (Berthold). All experiments were conducted in triplicates. For cell-cycle analysis, cells were fixed in 70% ethanol, stained with PI, and analyzed by flow cytometry.

Gene expression profiling and analysis of MS4A1 gene expression

DHL4, HBL-1, and U2932 cells were treated in biological duplicates with DMSO or 1.5 μmol/L MEN1703 for 24 hours. RNA was extracted using a GeneMATRIX Universal RNA purification kit (EURx). RNA quality was assessed using Agilent Bioanalyzer, and subsequently profiled on an Illumina Human HT_12 V3 Expression BeadChip by the AROS Applied biotechnology A/S (Denmark). The gene-by-sample expression matrix used for subsequent analyses was obtained from the raw “.IDAT” files using GenePattern (https://www.genepattern.org). Top 8,714 transcripts were selected by variation filtering using GenePattern's PreprocessDataset module, and by setting min fold change at ≥ 1.5 and min delta at 100. Expression data sets are available under GEO accession number GSE158120. Gene set enrichment analysis (GSEA) was performed using GSEA software (Broad Institute, Cambridge, MA; www.gsea-msigdb.org/gsea) using default settings, 1,000 permutations per phenotype and gene sets available in Molecular Signature Database (MSigDB).

MS4A1 transcript abundance in tumors with/without MYC-Ig translocation was evaluated in Hummel (26) and Green data sets (ref. 27; Hummel: GEO accession number GSE4475; Green: https://doi.org/10.1101/674259). In the Hummel data set, MS4A1 gene expression was evaluated in DLBCL and BL cases with/without MYC rearrangement. In this analysis, DLBCL and BLs with MYC translocation confirmed by FISH (n = 55) were compared with DLBCL lacking the MYC translocation (n = 127). In the MR Green data set, MS4A1 gene expression was evaluated in DLBCL and BLs from Nebraska cases with/without MYC rearrangement (n = 112; ref. 27). In this analysis, BL and DLBCLs with the translocation confirmed in NGS hybrid-capture sequencing (n = 17) were compared with DLBCLs lacking the MYC-Ig translocation (n = 95). MS4A1 Affymetrix probe sets 210356_x_at and 217418_x_at were used in this analysis.

Animal studies

For the in vivo assessment of MEN1703 single-agent activity, 1 × 107 of LY7 or U2932 cells in a 30% Matrigel Matrix (Corning, cat. no. 354230) were inoculated subcutaneously into NU/J and SCID/beige CB17 mice, respectively. The experiments were approved by the 1st Local Committee for Experiments with the Use of Laboratory Animals, Wroclaw, Poland (Permission no. 1/2013). Two weeks after inoculation, animals with established disease (tumor volume ≥100 mm3, n = 12 for U2932, n = 10 for LY7) were divided into 2 cohorts with equal average tumor sizes and given 50 mg/kg MEN1703 or vehicle (H2O) for 13–15 days, via oral gavage using a flexible cannula (Instech Laboratories, Inc.). Tumor growth was measured every second day using digital calipers. After the last dose, mice were euthanized, tumor tissues harvested and proteins isolated. To assess the effects of PIM inhibition on rituximab activity in vivo, the 6- to 7-week-old SCID Fox Chase female mice (Charles River Laboratories) were intravenously inoculated with 2×106 RAJI cells, modified to express Firefly Red Luciferase as described previously (28). Mice were treated daily with H2O or MEN1703 (50 mg/kg) via oral gavage for 18 days (starting on day 1 after inoculation of RAJI cells) or/and intraperitoneally (i.p.) with isotype control antibody or rituximab (10 mg/kg; five times, starting on day 5 after inoculation of tumor cells). Bioluminescence was measured once a week (starting from day 5 after first antibody administration). Mice were i.p. injected with D-luciferin (150 mg/kg), anesthetized with isoflurane and visualized using IVIS Imaging System (Xenogen). Images were analyzed with the Living Image 4.2 software (Caliper Life Science). Statistical significance was determined with the two-way ANOVA test and Statistica v12 software package. All animal studies were performed according to the protocol approved by the local bioethical committee (approval #WAW2/056/2020).

CDC assay

Following treatment with DMSO or pan-PIM inhibitors, 1 × 105 DHL4 and Raji cells were suspended in 200 μL RPMI medium supplemented with 10% human AB Rh+ serum (as a source of complement) and plated in duplicates into 96-well plate. Cells were incubated with increasing concentrations of rituximab for 1 hour. Cell viability was assessed by PI staining and flow cytometry. Cell survival was calculated as a percentage of control cells incubated without antibody ± SD. Statistical differences were determined using two-way ANOVA.

Antibody-dependent cellular phagocytosis assay

Antibody-dependent cellular phagocytosis (ADCP) experiments were performed as described previously (29), with modifications. Briefly, mononuclear cells from healthy donors' buffy coats were isolated by Histopaque centrifugation and monocytes purified using CD14 Microbeads (Miltenyi, cat. no. 130-050-201). To differentiate into macrophages, monocytes were cultured in the TexMACS medium (Miltenyi, 130-097-196) supplemented with 50 U/mL penicillin, 50 U/mL streptomycin, and 100ng/mL CSF-1 (R&D Systems, cat. no. 216-MC-005) followed by 2-day culture period in CSF-1–deficient medium. DLBCL cells were CFSE labeled and incubated for 3 days in the presence of pan-PIM inhibitors or DMSO as indicated. Thereafter, the cells were suspended in ice-cold and un-supplemented RPMI-1640 medium at 1 × 106/mL, and coated for 30 minutes with 1 μg/mL rituximab or IgG isotype control (BioLegend, 403501) at 4°C. Thereafter, the differentiated macrophages were washed with PBS, and the rituximab-coated DLBCL cells were added to the wells at a 1:5 effector:target ratio. After 2-hour incubation at 37°C/5% CO2, free-floating, not phagocytosed cells were washed away with PBS and phagocytosis analyzed using Eclipse Ti2 fluorescent microscope (Nikon). The phagocytic index was determined as the number of ingested CFSE-labeled DLBCL cells per 100 macrophages in three independent replicates in a representative experiment.

Statistical analyses

Statistical analyses were done using GraphPad Prism software (GraphPad Software Inc.) or R software package (version 4.0.4). Two-sided Student t test, Mann–Whitney, two-way ANOVA, and one-way ANOVA tests were used. Appropriate tests are provided in figure legends. P < 0.05 was considered statistically significant. The P values were marked with the asterisks on the charts (*, P < 0.05; **, P < 0.01; ***, P < 0.001.).

Additional detailed description of methods is available in the online Supplementary Data.

SEL24/MEN1703 attenuates DLBCL cell growth in vitro and in vivo

We first evaluated PIM1-3 expression in 57 DLBCL diagnostic biopsies by immunohistochemistry (IHC) and in a panel of six DLBCL cell lines by immunoblotting. Seventy percent (40/57) of diagnostic specimens were positive for at least one PIM isoform (hereafter PIM-positive). PIM-positive cases were classified more frequently as ABC than GCB subtype (80% vs. 44%, P = 0.01, Fisher exact test; Fig. 1A and B). All cell lines expressed at least two PIM isoforms (Supplementary Fig. S1). We next assessed in vitro and in vivo consequences of PIM kinase inhibition for DLBCL growth and survival. siRNA-based knockdown of all three PIM isoforms in DHL4 and U2932 cells markedly increased the number of apoptotic cells (Supplementary Fig. S2A). In subsequent analyses, we used a pan-PIM/FLT3 inhibitor, MEN1703, currently undergoing clinical trials (30). Because MEN1703 also targets FLT3 (30), we first confirmed complete absence of FLT3 in DLBCL cell lines (Supplementary Fig. S2B). MEN1703 induced cell-cycle arrest in five of six DLBCL cell lines (Supplementary Fig. S2C) and decreased cell proliferation in a dose-dependent manner in all tested GCB- and ABC-type cell lines (EC50 range, 0.29–1.16 μmol/L; Fig. 1C; Supplementary Fig. S2D). MEN1703 triggered activation of caspases 3/7, 8, and 9, induced PARP cleavage and apoptosis (Fig. 1D; Supplementary Fig. S2E–S2G). In murine xenograft models representing GCB (Ly7) and ABC (U2932) DLBCL subtypes, MEN1703 inhibited tumor growth by 58% and 93%, respectively (two-way ANOVA, P < 0.0001; Fig. 1E).

Figure 1.

PIM inhibition is toxic to DLBCL cells in vitro and in vivo. A, IHC analysis of PIM1–3 expression in DLBCL biopsies (n = 57). Shown are representative GCB and ABC cases, PIM1–3 negative (#14), PIM1–3 positive (#21 and #28), and PIM1 positive and PIM2–3 negative (#49). Original magnification, ×600. B, Summary of the IHC analysis of PIM expression performed in a panel of 57 patients with DLBCL. C, Effects of MEN1703 on DLBCL cell line proliferation. Cells were incubated with DMSO or 0.8 μmol/L MEN1703 for 72 hours and cellular proliferation was determined by the MTS assay. Bars represent mean of proliferation relative to DMSO treatment. Error bars, ±SD of three independent replicates in a representative experiment. P values were determined using the two-sided t test: *, P < 0.05; ***, P < 0.001. Dose–response curves and calculated EC50 values for MEN1703 are presented in Supplementary Fig. S2D. D, Cellular apoptosis in MEN1703-treated DLBCL cells. After 72 hours of incubation with DMSO or MEN1703 (1.5 μmol/L), apoptosis was measured by Annexin V/PI staining, followed by flow cytometric analysis. Each point represents percentage of apoptotic cells in DMSO- or MEN1703-treated cell lines.Annexin V–positive or Annexin V/PI- positive cells were considered apoptotic. Annexin V/PI dot plots are shown in Supplementary Fig. S2G. E, Tumor growth kinetics in MEN1703 or vehicle–treated mice. LY7 and U2932 cells were suspended in 30% Matrigel and injected into NU/J and SCID/beige CB17 mice, respectively. When tumor volume reached ≥100 mm3, LY7 xenograft-bearing mice were treated once daily with MEN1703 (50 mg/kg) or H2O (control) via oral gavage, and U2932 xenograft-bearing mice were treated twice a day with the same dose of MEN1703 or H2O. Tumor growth kinetics in the control and MEN1703 mice were compared by the two-way ANOVA test. Each experimental group consisted of 5 to 6 mice. Error bars, ±SEM. F, PIM inhibition downregulates PIM-specific 4EBP1 and S6 phosphorylations and MCL1 protein levels. Cells were incubated for 6 hours with DMSO or 1.5 μmol/L MEN1703, harvested, and lysed. Expression of p4EBP1 (S65) and pS6 (S235/236) and MCL1 was analyzed by immunoblotting. GAPDH served as a loading control. G, MEN1703 decreases BAD-S112 phosphorylation in DLBCL cells. Following 1.5-hour incubation with DMSO or 1.5 μmol/L MEN1703, pBAD-S112 protein levels were assessed by immunoblotting. GAPDH served as a loading control. Data in C-D and F–G are representative of three independent experiments.

Figure 1.

PIM inhibition is toxic to DLBCL cells in vitro and in vivo. A, IHC analysis of PIM1–3 expression in DLBCL biopsies (n = 57). Shown are representative GCB and ABC cases, PIM1–3 negative (#14), PIM1–3 positive (#21 and #28), and PIM1 positive and PIM2–3 negative (#49). Original magnification, ×600. B, Summary of the IHC analysis of PIM expression performed in a panel of 57 patients with DLBCL. C, Effects of MEN1703 on DLBCL cell line proliferation. Cells were incubated with DMSO or 0.8 μmol/L MEN1703 for 72 hours and cellular proliferation was determined by the MTS assay. Bars represent mean of proliferation relative to DMSO treatment. Error bars, ±SD of three independent replicates in a representative experiment. P values were determined using the two-sided t test: *, P < 0.05; ***, P < 0.001. Dose–response curves and calculated EC50 values for MEN1703 are presented in Supplementary Fig. S2D. D, Cellular apoptosis in MEN1703-treated DLBCL cells. After 72 hours of incubation with DMSO or MEN1703 (1.5 μmol/L), apoptosis was measured by Annexin V/PI staining, followed by flow cytometric analysis. Each point represents percentage of apoptotic cells in DMSO- or MEN1703-treated cell lines.Annexin V–positive or Annexin V/PI- positive cells were considered apoptotic. Annexin V/PI dot plots are shown in Supplementary Fig. S2G. E, Tumor growth kinetics in MEN1703 or vehicle–treated mice. LY7 and U2932 cells were suspended in 30% Matrigel and injected into NU/J and SCID/beige CB17 mice, respectively. When tumor volume reached ≥100 mm3, LY7 xenograft-bearing mice were treated once daily with MEN1703 (50 mg/kg) or H2O (control) via oral gavage, and U2932 xenograft-bearing mice were treated twice a day with the same dose of MEN1703 or H2O. Tumor growth kinetics in the control and MEN1703 mice were compared by the two-way ANOVA test. Each experimental group consisted of 5 to 6 mice. Error bars, ±SEM. F, PIM inhibition downregulates PIM-specific 4EBP1 and S6 phosphorylations and MCL1 protein levels. Cells were incubated for 6 hours with DMSO or 1.5 μmol/L MEN1703, harvested, and lysed. Expression of p4EBP1 (S65) and pS6 (S235/236) and MCL1 was analyzed by immunoblotting. GAPDH served as a loading control. G, MEN1703 decreases BAD-S112 phosphorylation in DLBCL cells. Following 1.5-hour incubation with DMSO or 1.5 μmol/L MEN1703, pBAD-S112 protein levels were assessed by immunoblotting. GAPDH served as a loading control. Data in C-D and F–G are representative of three independent experiments.

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To demonstrate the inhibitor's on-target activity, we next evaluated the activity of known PIM kinase substrates following MEN1703 treatment. Genetic (siRNA) and chemical (MEN1703) PIM inhibition decreased 4EBP1 phosphorylation at the PIM-specific S65 residue, S6 phosphorylation at S235/236, and blocked BAD S112 phosphorylation (Supplementary Fig. S3A; Fig. 1F and G; ref. 31). Consistent with these in vitro data, p4EBP1 and pS6 levels were markedly downregulated in explanted xenografts from MEN1703-treated animals (Supplementary Fig. S3B), indicating that PIM inhibition in DLBCL cells decreases cap-dependent protein translation. In line with this observation, expression of MCL1, regulated mainly at the translational level (32), significantly decreased after PIM inhibition (Fig. 1F). Given the previous studies revealing synergistic effects of simultaneous MCL1 and BCL2 inhibition in DLBCL cells (33, 34), we evaluated in vitro interaction between MEN1703 and venetoclax and found synergistic activity of the combinatorial treatment (Supplementary Fig. S3C).

PIM inhibition decreases MYC-dependent gene expression and inhibits DLBCL cell proliferation and survival

To obtain an unbiased insight into the mechanisms underlying cytotoxicity of PIM inhibition, we assessed global gene expression profiles of three representative DLBCL cell lines (DHL4, HBL-1, and U2932) following 24 hours treatment with MEN1703. GSEA analyses of PIM inhibitor–treated DLBCL cells revealed downregulated expression of multiple MYC-dependent genes including those involved in cell-cycle control and proliferation (Fig. 2A; Supplementary Fig. S4; Supplementary Table S1).

Figure 2.

Toxic effects of PIM inhibition in DLBCL cells are partially mediated by MYC. A, GSEA enrichment plots of MYC target genes in DLBCL cell lines (DHL4, HBL-1, and U2932) treated with DMSO or 1.5 μmol/L MEN1703 for 24 hours and subjected to gene expression analysis. Presented are gene sets available under MSigDB systematic names: M1249, M16789, and M5926. The positions of the MYC target genes are significantly skewed toward the left end of the sorted list, reflecting their decreased expression in MEN1703-treated cell lines. Corresponding heatmaps including core enrichment genes are presented in Supplementary Fig. S4. B, IHC analysis of PIM1–3 and MYC expression in DLBCL biopsies (n = 57). Shown are representative GCB and ABC cases, PIM1–3 and MYC negative (#14), PIM1 positive and MYC negative (#49), and PIM1–3 and MYC positive (#21 and #28). Original magnification, ×600. C, Summary of the IHC analysis of PIM and MYC expression performed in a panel of 57 patients with DLBCL. MYC-high tumors were defined by expression of MYC in ≥30% of lymphoma cells (n = 35); MYC-low tumors were defined by MYC expression in less than 30% of the malignant cells (n = 22).PIM expression was more abundant in MYC-high than MYC-low tumors (P < 0.0001, Fisher exact test). D, siRNA-mediated PIM1, PIM2, and PIM3 silencing in the DHL4 cell line. Cells were electroporated with nontargeting siRNA (Ctr) or siRNAs targeting individual PIM isoforms and cultured for 24–48 hours. Afterward, protein levels of PIM kinases and MYC were assessed by immunoblotting. GAPDH served as a loading control. PIM1 and MYC immunoblots were performed on the same PVDF membranes and have the same loading control (GAPDH). E, siRNA-mediated PIM1, PIM2, and PIM3 protein knockdown in DHL4 and U2932 cells. Cells were electroporated with a nontargeting siRNA or with a siRNA cocktail targeting all three PIM kinases. Thereafter, cells were cultured for 48 hours, and expression of PIM1, PIM2, PIM3, and MYC was assessed by immunoblotting. F, Inhibition of PIM kinases decreases MYC levels in DLBCL cells. Cells were incubated for 6 hours with DMSO alone or with increasing doses (1–5 μmol/L) of MEN1703 (MEN), harvested, and lysed. Expression of MYC was analyzed by immunoblotting. GAPDH served as a loading control. G, MEN1703 decreases MYC expression in primary DLBCL cells. Patient-derived primary DLBCL cells were incubated with DMSO or 2.5 μmol/L MEN1703 for 24 hours. Thereafter, intracellular MYC levels were determined by flow cytometry. Bar graphs depict median fluorescence intensities of MYC relative to DMSO treatment. Error bars, ±SD of two independent replicates in a representative experiment. P values were determined using the two-sided t test: *, P < 0.05; **, P < 0.01; ***, P < 0.001. H, Effects of MEN1703 on proliferation of MYC-wild-type and MYC T58A-mutant DHL4 cells. Parental DHL4 (MYC_WT) and MYC-mutant (T58A) cells incubated with DMSO or 0.5 μmol/L MEN1703 for 72 hours and cellular proliferation was determined by MTS assay. Bars represent mean proliferation relative to DMSO treatment. Error bars, ±SD of three independent replicates in a representative experiment. The P values were determined using the two-sided t test: ***, P < 0.001. I, Cell death in MEN1703-treated MYC WT and T58A-mutant DHL4 cells. After 48 hours of incubation with DMSO or 1.5 μmol/L MEN1703, cell death was measured by 7AAD staining, followed by flow cytometric analysis. Data in D–F and H–I are representative of three independent experiments.

Figure 2.

Toxic effects of PIM inhibition in DLBCL cells are partially mediated by MYC. A, GSEA enrichment plots of MYC target genes in DLBCL cell lines (DHL4, HBL-1, and U2932) treated with DMSO or 1.5 μmol/L MEN1703 for 24 hours and subjected to gene expression analysis. Presented are gene sets available under MSigDB systematic names: M1249, M16789, and M5926. The positions of the MYC target genes are significantly skewed toward the left end of the sorted list, reflecting their decreased expression in MEN1703-treated cell lines. Corresponding heatmaps including core enrichment genes are presented in Supplementary Fig. S4. B, IHC analysis of PIM1–3 and MYC expression in DLBCL biopsies (n = 57). Shown are representative GCB and ABC cases, PIM1–3 and MYC negative (#14), PIM1 positive and MYC negative (#49), and PIM1–3 and MYC positive (#21 and #28). Original magnification, ×600. C, Summary of the IHC analysis of PIM and MYC expression performed in a panel of 57 patients with DLBCL. MYC-high tumors were defined by expression of MYC in ≥30% of lymphoma cells (n = 35); MYC-low tumors were defined by MYC expression in less than 30% of the malignant cells (n = 22).PIM expression was more abundant in MYC-high than MYC-low tumors (P < 0.0001, Fisher exact test). D, siRNA-mediated PIM1, PIM2, and PIM3 silencing in the DHL4 cell line. Cells were electroporated with nontargeting siRNA (Ctr) or siRNAs targeting individual PIM isoforms and cultured for 24–48 hours. Afterward, protein levels of PIM kinases and MYC were assessed by immunoblotting. GAPDH served as a loading control. PIM1 and MYC immunoblots were performed on the same PVDF membranes and have the same loading control (GAPDH). E, siRNA-mediated PIM1, PIM2, and PIM3 protein knockdown in DHL4 and U2932 cells. Cells were electroporated with a nontargeting siRNA or with a siRNA cocktail targeting all three PIM kinases. Thereafter, cells were cultured for 48 hours, and expression of PIM1, PIM2, PIM3, and MYC was assessed by immunoblotting. F, Inhibition of PIM kinases decreases MYC levels in DLBCL cells. Cells were incubated for 6 hours with DMSO alone or with increasing doses (1–5 μmol/L) of MEN1703 (MEN), harvested, and lysed. Expression of MYC was analyzed by immunoblotting. GAPDH served as a loading control. G, MEN1703 decreases MYC expression in primary DLBCL cells. Patient-derived primary DLBCL cells were incubated with DMSO or 2.5 μmol/L MEN1703 for 24 hours. Thereafter, intracellular MYC levels were determined by flow cytometry. Bar graphs depict median fluorescence intensities of MYC relative to DMSO treatment. Error bars, ±SD of two independent replicates in a representative experiment. P values were determined using the two-sided t test: *, P < 0.05; **, P < 0.01; ***, P < 0.001. H, Effects of MEN1703 on proliferation of MYC-wild-type and MYC T58A-mutant DHL4 cells. Parental DHL4 (MYC_WT) and MYC-mutant (T58A) cells incubated with DMSO or 0.5 μmol/L MEN1703 for 72 hours and cellular proliferation was determined by MTS assay. Bars represent mean proliferation relative to DMSO treatment. Error bars, ±SD of three independent replicates in a representative experiment. The P values were determined using the two-sided t test: ***, P < 0.001. I, Cell death in MEN1703-treated MYC WT and T58A-mutant DHL4 cells. After 48 hours of incubation with DMSO or 1.5 μmol/L MEN1703, cell death was measured by 7AAD staining, followed by flow cytometric analysis. Data in D–F and H–I are representative of three independent experiments.

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Because these observations suggest a functional link between PIM kinases and MYC-dependent oncogenic program in DLBCL cells, we asked whether PIMs and MYC are coexpressed in DLBCL patients. In the investigated series of DLBCL diagnostic sections, 67% (38/57) of cases were double MYC/PIM-positive. Notably, 100% of cases with high MYC expression (MYC present in ≥30% of lymphoma cells, n = 35) were PIM-positive, whereas 86% of cases with undetectable or low MYC expression were PIM-negative (MYC detected in <30% of the cells, n = 22; P < 0.0001, Fisher exact test; Fig. 2B and C; Supplementary Table S2). Consistent with these data, primary high-grade B-cell lymphoma (HGBL) diagnostic samples (n = 5) exhibited high MYC expression in ≥90% of the malignant cells and were uniformly double- or triple- PIM-positive (Supplementary Fig. S5).

To investigate the mechanisms underlying MYC/PIMs coexpression in DLBCL cells, we knocked down MYC using siRNA or with an MYC-MAX dimerization inhibitor, 10058-F4, facilitating MYC degradation (35), and evaluated the expression of PIM isoforms. MYC depletion markedly downregulated PIM expression in DLBCL cell lines (Supplementary Fig. S6A and S6B). Taken together, these results demonstrate that MYC increases PIM expression in DLBCL and HGBL.

PIM1 and PIM2 were previously shown to increase MYC stability and protein expression level (36, 37). To investigate the role of PIMs in regulation of MYC abundance in DLBCL cells, we transiently silenced PIM1, -2, and/or -3 expression using siRNA and assessed MYC protein level by immunoblotting. In DHL4 cells, PIM1 and PIM2 knockdown decreased MYC levels by 23% and 14%, respectively (Fig. 2D). In marked contrast, simultaneous PIM1, -2, and -3 depletion downregulated MYC levels to 35% of baseline in DHL4 and to 53% in U2932 cells (Fig. 2E), indicating that effective decrease of MYC levels in DLBCL cells cannot be achieved by inhibition of the individual PIM isoforms. Consistent with these observations, MEN1703 decreased MYC protein level in all tested cell lines in a dose- and time-dependent manner (Fig. 2F; Supplementary Fig. S7A and S7B). At the earliest time points (2–6 hours), PIM inhibition did not significantly increase apoptosis, indicating that MYC downregulation is not a consequence of cell death (Supplementary Fig. S7C). Other available pan-PIM inhibitors, SGI-1776 and PIM447, exhibited similar activity in depleting MYC (Supplementary Fig. S7D). A proteasome inhibitor, MG132, partially protected MYC from PIM inhibitor induced degradation, indicating that PIMs regulate MYC abundance at least in part by increasing protein stability and/or blocking its proteasomal degradation (Supplementary Fig. S7E). Importantly, MEN1703 decreased MYC expression in explanted DLBCL xenografts and in patient-derived primary DLBCL cells (Supplementary Fig. S3B; Fig. 2G; Supplementary Fig. S7F).

To determine whether PIM inhibition–mediated cytotoxicity requires MYC degradation, we modified DHL4 cells to express T58A MYC mutant. MYC T58 phosphorylation facilitates subsequent ubiquitin-mediated degradation; thus, T58A mutant is more resistant to degradation than WT MYC (37, 38). DHL4 cells expressing the mutant were partially protected from PIM inhibitor–induced proliferation block (19% vs. 50% in MYC WT vs. T58A mutant-expressing cells, respectively; Fig. 2H). The T58A mutant moderately attenuated induction of apoptosis (36% vs. 50% in MYC WT vs. T58A mutant-expressing cells; Fig. 2I). These results demonstrate that cytotoxic activity of PIM inhibition involves both MYC-dependent and MYC-independent mechanisms.

PIMs modulate MYC-dependent expression of PLK1 in DLBCL cells

GSEA analyses of PIM inhibitor–treated DLBCL cells revealed downregulated expression of multiple MYC-dependent genes involved in cell-cycle control (Supplementary Fig. S4). Among these, transcript levels of PLK1 were uniformly decreased following PIM inhibition in all tested cell lines (Supplementary Table S1). Because PLK1 is a master regulator of mitosis (39, 40), we assessed PLK1 transcript and protein abundance following PIM inhibition in a panel of six DLBCL cell lines. In accordance with the microarray data, MEN1703 decreased PLK1 transcript and protein levels (Fig. 3A and B). We further confirmed the specificity of these effects in U2932 and DHL4 cells with siRNA-mediated knockdown of PIMs (Fig. 3C).

Figure 3.

PIM inhibition downregulates PLK1 in an MYC-dependent manner in DLBCL cells. A, PIM inhibition decreases PLK1 protein expression. DLBCL cells were incubated for 24 hours with DMSO or with 1.5 μmol/L MEN1703. PLK1 protein levels were assessed by immunoblotting. B, MEN1703 downregulates PLK1 transcription. Cells were incubated with DMSO alone or with 1.5 μmol/L MEN1703 for 6 hours. Thereafter, expression of PLK1 was assessed by qPCR. Bars represent mean of PLK1 expression relative to DMSO treatment. Error bars, ±SD of n = 3 replicates. The P values were determined using the two-sided t test: *, P < 0.05; **, P < 0.01; ***, P < 0.001. C, PIM1–3 knockdown decreases PLK1 protein levels. U2932 and DHL4 cells were electroporated with siRNA cocktail targeting all three PIM (3xPim) kinases or with the nontargeting control. Thereafter, cells were cultured for 48 hours, and PLK1 protein abundance was assessed by immunoblotting. PLK1 immunoblots in C were performed on the same cell lysates and the same PVDF membrane as MYC immunoblots in Fig. 2E and have the same loading control (GAPDH). D, MYC inhibition decreases PLK1 expression. U2932 and DHL4 cells were electroporated with MYC-targeting siRNA or with a nontargeting control. Thereafter, cells were cultured for 48 hours and PLK1 protein abundance was assessed by immunoblotting. E, PLK1 expression in MEN1703-treated MYC WT and MYC-T58A-mutant DHL4 cells. Parental DHL4 (MYC_WT) and MYC-mutant (T58A) cells were incubated with DMSO or 1.5 μmol/L MEN1703 for 48 hours and PLK1 protein abundance was determined by immunoblotting. GAPDH served as a loading control in A, C–E. F, PIM inhibition decreases MYC binding to PLK1 promoter. DHL4 and U2932 cells were treated overnight with DMSO or 1.5 μmol/L MEN1703. MYC binding to PLK1 promoter region −231/−147 located upstream TSS (+1) was determined by chromatin immunoprecipitation and quantitative PCR. Bars represent mean CT values normalized to control IgG, relative to DMSO treatment ±SD of n = 3 replicates. P values were determined using the two-sided t test: *, P < 0.05. G, EC50 dose–response curves for BI6727 in DLBCL cell lines. Cellular proliferation was determined by the MTS assay after 72 hours of incubation. The individual data points in dose–response curves represent the mean ±SD of n = 3 replicates. Calculated IC50 values are indicated below the plots. Data in A–G are representative of three independent experiments.

Figure 3.

PIM inhibition downregulates PLK1 in an MYC-dependent manner in DLBCL cells. A, PIM inhibition decreases PLK1 protein expression. DLBCL cells were incubated for 24 hours with DMSO or with 1.5 μmol/L MEN1703. PLK1 protein levels were assessed by immunoblotting. B, MEN1703 downregulates PLK1 transcription. Cells were incubated with DMSO alone or with 1.5 μmol/L MEN1703 for 6 hours. Thereafter, expression of PLK1 was assessed by qPCR. Bars represent mean of PLK1 expression relative to DMSO treatment. Error bars, ±SD of n = 3 replicates. The P values were determined using the two-sided t test: *, P < 0.05; **, P < 0.01; ***, P < 0.001. C, PIM1–3 knockdown decreases PLK1 protein levels. U2932 and DHL4 cells were electroporated with siRNA cocktail targeting all three PIM (3xPim) kinases or with the nontargeting control. Thereafter, cells were cultured for 48 hours, and PLK1 protein abundance was assessed by immunoblotting. PLK1 immunoblots in C were performed on the same cell lysates and the same PVDF membrane as MYC immunoblots in Fig. 2E and have the same loading control (GAPDH). D, MYC inhibition decreases PLK1 expression. U2932 and DHL4 cells were electroporated with MYC-targeting siRNA or with a nontargeting control. Thereafter, cells were cultured for 48 hours and PLK1 protein abundance was assessed by immunoblotting. E, PLK1 expression in MEN1703-treated MYC WT and MYC-T58A-mutant DHL4 cells. Parental DHL4 (MYC_WT) and MYC-mutant (T58A) cells were incubated with DMSO or 1.5 μmol/L MEN1703 for 48 hours and PLK1 protein abundance was determined by immunoblotting. GAPDH served as a loading control in A, C–E. F, PIM inhibition decreases MYC binding to PLK1 promoter. DHL4 and U2932 cells were treated overnight with DMSO or 1.5 μmol/L MEN1703. MYC binding to PLK1 promoter region −231/−147 located upstream TSS (+1) was determined by chromatin immunoprecipitation and quantitative PCR. Bars represent mean CT values normalized to control IgG, relative to DMSO treatment ±SD of n = 3 replicates. P values were determined using the two-sided t test: *, P < 0.05. G, EC50 dose–response curves for BI6727 in DLBCL cell lines. Cellular proliferation was determined by the MTS assay after 72 hours of incubation. The individual data points in dose–response curves represent the mean ±SD of n = 3 replicates. Calculated IC50 values are indicated below the plots. Data in A–G are representative of three independent experiments.

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Because PLK1 is transcriptionally regulated by MYC (41), we asked whether MYC downregulation induced by PIM inhibition is responsible for the decreased PLK1 expression in DLBCL cells. After confirming that MYC siRNA decreases PLK1 protein expression (Fig. 3D), we compared PLK1 protein levels in MYC WT and T58A mutant–transduced DHL4 cells treated with MEN1703. The inhibitor decreased PLK1 levels in MYC WT cells by 53%, and by 31% in MYC T58A cells, indicating that the pan-PIM inhibitor regulates PLK1 expression at least partially via MYC (Fig. 3E). Consistent with this, PIM inhibition markedly reduced MYC binding to the PLK1 promoter region containing the E-box motif [−231/−147 upstream of PLK1 transcription start site (TSS); Fig. 3F]. Finally, a potent and selective PLK1 inhibitor volasertib markedly suppressed DLBCL cell proliferation at low-nanomolar concentrations, indicating that PLK1 downregulation in MEN1703-treated cells is an important mechanism of inhibitor activity (Fig. 3G).

PIM–MYC axis represses CD20 expression in B-NHL cells

Previous studies in murine B-cell lymphomas and BL suggested that MYC suppresses CD20 expression (42, 43). We thus hypothesized that PIM inhibition might induce CD20 expression in DLBCL and other B-NHLs through downregulation of MYC. First, to characterize the role of MYC in CD20 regulation in primary human lymphomas, we investigated MS4A1(CD20) mRNA levels in a data set including 182 BL and DLBCL patients (26) and in an independent data set from 116 BL and DLBCL patients (contributed by Dr. M.R. Green; ref. 27). Because MYC translocation to immunoglobulin locus is associated with increased MYC levels and activity (8), we divided BL and DLBCL patients into two groups, harboring wild-type MYC or MYC–Ig translocation (Tx). In both data sets, MS4A1(CD20) mRNA levels were significantly higher in MYC WT patients than in those with MYC–Ig translocation (Fig. 4A). To further investigate the role and mechanism of MYC in MS4A1 regulation, we evaluated CD20 protein levels in DHL4 and RAJI (BL) cells following siRNA-mediated MYC knockdown and found that MYC inhibition increases CD20 levels (Fig. 4B). Likewise, in a lymphoblastic P493-6 B cells carrying tetracycline-regulated MYC (44), MYC downregulation increased MS4A1/CD20 mRNA level and surface abundance in a time-dependent manner (Fig. 4C; Supplementary Fig. S8A).

Figure 4.

PIM kinases repress CD20 expression through MYC. A, The median MS4A1/CD20 expression for probes 1 (210356_x_at) and 2 (217418_x_at) in two independent sets of DLBCL and BL patients [Hummel, n = 182, GEO accession number GSE4475 (26) and Green, n = 116 (27)], grouped according to the MYC–Ig translocation status. Tx, translocation present; WT, translocation absent. Horizontal line, median; bars, ±25–75 percentile; whiskers, ±range. P values were calculated using Mann–Whitney test: **, P < 0.01; ***, P < 0.001. B, MYC knockdown increases surface CD20 expression. Cells were electroporated with an MYC-targeting siRNA or a nontargeting siRNA (control) and cultured for 96 hours. Thereafter, CD20 expression was assessed by flow cytometry. Bars represent median fluorescence intensity (MFI) of CD20 calculated as a percentage of control siRNA-transfected cells CD20 MFI. Error bars, ±SD of two independent replicates in a representative experiment. P values were determined using the two-sided t test: **, P < 0.01; ***, P < 0.001. C, MYC downregulation increases CD20 levels in lymphoblastic cell line P493–6. Cells were left untreated or were stimulated with 10 ng/mL doxycycline for 24–96 hours to repress MYC. CD20 surface abundance was assessed by flow cytometry. Bars, MFI of CD20. Error bars, ±SD of two independent replicates in a representative experiment. P values were determined using ANOVA and Tukey HSD post hoc test: **, P < 0.01; ***, P < 0.001. D, PIM inhibition decreases MYC binding to MS4A1 promoter. DHL4 cells were treated with DMSO or 1.5 μmol/L MEN1703 for 24 hours. MYC binding to MS4A1 promoter regions −313/−198 and −126/−39 located upstream TSS (+1) was determined by chromatin immunoprecipitation and quantitative PCR. Bars, mean CT values normalized to control IgG. Error bars, ±SD of three independent replicates in a representative experiment. P values were determined using the two-sided t test: *, P < 0.05; ***, P < 0.001. E, PIM1–3 knockdown increases CD20 surface abundance. DHL4 cells were electroporated with siRNA cocktail targeting all three PIM (3xPim) kinases or with a control siRNA. CD20 surface abundance was assessed by fold change after 96 hours. Bars represent CD20 MFI relative to control siRNA ±SD of n = 2 replicates. F, PIM inhibition induces CD20 surface expression. DHL4 and RAJI cells were incubated with DMSO or 1.5 μmol/L MEN1703 for 48 hours. Afterward, CD20 expression was assessed by flow cytometry. Bars represent CD20 MFI relative to DMSO treatment ±SD of n = 2 replicates. G, PIM inhibition induces CD20 surface expression in an MYC-dependent manner. DHL4 parental (MYC_WT) and MYC-mutant (T58A) cells were incubated with DMSO or 1.5 μmol/L MEN1703 for 48 hours. CD20 expression was assessed by fold change after 24 hours. Bars represent CD20 MFI relative to DMSO treatment ±SD of n = 2 replicates in a representative experiment. P values in A–C and E–G were determined using the two-sided t test: *, P < 0.05; **, P < 0.01; ***, P < 0.001. Data in B–G are representative of three independent experiments.

Figure 4.

PIM kinases repress CD20 expression through MYC. A, The median MS4A1/CD20 expression for probes 1 (210356_x_at) and 2 (217418_x_at) in two independent sets of DLBCL and BL patients [Hummel, n = 182, GEO accession number GSE4475 (26) and Green, n = 116 (27)], grouped according to the MYC–Ig translocation status. Tx, translocation present; WT, translocation absent. Horizontal line, median; bars, ±25–75 percentile; whiskers, ±range. P values were calculated using Mann–Whitney test: **, P < 0.01; ***, P < 0.001. B, MYC knockdown increases surface CD20 expression. Cells were electroporated with an MYC-targeting siRNA or a nontargeting siRNA (control) and cultured for 96 hours. Thereafter, CD20 expression was assessed by flow cytometry. Bars represent median fluorescence intensity (MFI) of CD20 calculated as a percentage of control siRNA-transfected cells CD20 MFI. Error bars, ±SD of two independent replicates in a representative experiment. P values were determined using the two-sided t test: **, P < 0.01; ***, P < 0.001. C, MYC downregulation increases CD20 levels in lymphoblastic cell line P493–6. Cells were left untreated or were stimulated with 10 ng/mL doxycycline for 24–96 hours to repress MYC. CD20 surface abundance was assessed by flow cytometry. Bars, MFI of CD20. Error bars, ±SD of two independent replicates in a representative experiment. P values were determined using ANOVA and Tukey HSD post hoc test: **, P < 0.01; ***, P < 0.001. D, PIM inhibition decreases MYC binding to MS4A1 promoter. DHL4 cells were treated with DMSO or 1.5 μmol/L MEN1703 for 24 hours. MYC binding to MS4A1 promoter regions −313/−198 and −126/−39 located upstream TSS (+1) was determined by chromatin immunoprecipitation and quantitative PCR. Bars, mean CT values normalized to control IgG. Error bars, ±SD of three independent replicates in a representative experiment. P values were determined using the two-sided t test: *, P < 0.05; ***, P < 0.001. E, PIM1–3 knockdown increases CD20 surface abundance. DHL4 cells were electroporated with siRNA cocktail targeting all three PIM (3xPim) kinases or with a control siRNA. CD20 surface abundance was assessed by fold change after 96 hours. Bars represent CD20 MFI relative to control siRNA ±SD of n = 2 replicates. F, PIM inhibition induces CD20 surface expression. DHL4 and RAJI cells were incubated with DMSO or 1.5 μmol/L MEN1703 for 48 hours. Afterward, CD20 expression was assessed by flow cytometry. Bars represent CD20 MFI relative to DMSO treatment ±SD of n = 2 replicates. G, PIM inhibition induces CD20 surface expression in an MYC-dependent manner. DHL4 parental (MYC_WT) and MYC-mutant (T58A) cells were incubated with DMSO or 1.5 μmol/L MEN1703 for 48 hours. CD20 expression was assessed by fold change after 24 hours. Bars represent CD20 MFI relative to DMSO treatment ±SD of n = 2 replicates in a representative experiment. P values in A–C and E–G were determined using the two-sided t test: *, P < 0.05; **, P < 0.01; ***, P < 0.001. Data in B–G are representative of three independent experiments.

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In addition to its well-defined role of transcriptional activator, MYC has been shown to repress transcription of certain genes by binding to its cognate canonical or noncanonical E-box motifs localized within promoter regions (45). MYC associated with two identified noncanonical E-boxes (CACCTG) located upstream MS4A1 TSS at positions −245/−240 and −45/−40 (Supplementary Fig. S8B), and the binding was markedly reduced by MEN1703 (Fig. 4D).

Because MYC is a potent modulator of transcription of numerous microRNAs (miRNA; ref. 46), we also assessed MYC-dependent and miRNA-driven mechanisms regulating CD20 expression. We first screened a curated list of known 59 MYC-upregulated miRNAs (47) for potential interactions with MS4A1 transcript and identified miR-222 matching sequences in the 3′UTR of the MS4A1 gene. In DHL4 cells incubated with MYC inhibitor 10058-F4 or with MEN1703, miR-222 levels markedly decreased (Supplementary Fig. S9A). To investigate whether miR-222 represses MS4A1 through direct 3′UTR interaction, wild-type 3′UTR fragment or its seed-sequence mutants were cloned into a luciferase reporter plasmid. In cells cotransfected with the vector containing wild-type 3′UTR MS4A1 sequence and miR-222 mimic, luciferase activity was markedly decreased, compared with a nontargeting control (Supplementary Fig. S9B). In contrast, miR-222 did not block luciferase expression in cells transfected with reporter vectors carrying 3′UTRs with mutated seed sequences. In DHL4 cells transfected with miR-222 mimic, but not the nontargeting control, mRNA and surface CD20 expression markedly decreased (Supplementary Fig. S9C and S9D).

We next examined the effects of genetic and pharmacologic PIM inhibition on CD20 expression in additional B-NHL models. In line with the increased transcript and CD20 protein levels observed after MYC inhibition (Fig. 4B and C), PIMs siRNA or PIM inhibitors (MEN1703, SGI-1776 and PIM447) markedly induced MS4A1/CD20 mRNA and protein in DLBCL and BL cells (Fig. 4E and F; Supplementary Fig. S8C and S8D). PIM-mediated CD20 induction was MYC-dependent, because MEN1703-treated DHL4 cells modified to express MYC T58A mutant did not show increased CD20 expression (Fig. 4G). In addition, PIM inhibition increased surface CD20 expression in primary lymphoma cells of different subtypes (CLL, Richter's transformation, DLBCL, and MCL; Supplementary Fig. S8E). Taken together, our results indicate that PIM–MYC axis suppresses CD20 expression in B-cell lymphomas.

PIM inhibition augments the activity of anti-CD20 antibodies in vitro and in vivo

Given the increased CD20 surface expression following PIM inhibition in B-NHL cells, we hypothesized that PIM inhibitors would augment the activity of anti-CD20 antibodies. To address this hypothesis, we assessed the consequences of PIM inhibition on CDC, ADCC, and ADCP of anti-CD20 antibodies in DLBCL cells. First, DHL4 and RAJI cells were pretreated with DMSO or MEN1703, incubated for 1 hour with increasing concentrations of rituximab in the presence of 10% human serum as a source of complement, and their viability was assessed using PI staining. Rituximab triggered higher complement-dependent toxicity in MEN1703-treated DHL4 and RAJI cells, compared with control (DMSO-treated) cells (two-way ANOVA, P < 0.0001; Fig. 5A). Similar results were obtained using SGI-1776 and PIM447 (Supplementary Fig. S10A). Calculated CIs demonstrated synergistic nature of rituximab combinations with every pan-PIM inhibitor used in the CDC assay.

Figure 5.

PIM inhibition potentiates CDC and ADCP of rituximab. Increased CDC triggered by rituximab in MEN1703-treated cells. DHL4 and RAJI were pretreated with DMSO or 1.5 μmol/L MEN1703 for 48 hours and then treated with rituximab (1 hour) in the presence of human AB Rh+ serum. Cell death was determined with PI staining and flow cytometry. Error bars, ±SD of two independent replicates in a representative experiment. Combination indexes (CI) were calculated using CompuSyn software. Statistical significance was determined using two-way ANOVA. B, PIM inhibition in DLBCL cells potentiates rituximab-dependent phagocytosis. After 72 hours of treatment with DMSO or MEN1703 (1 μmol/L), CFSE-labeled DHL4 and DHL6 cells were incubated with human peripheral blood–derived macrophages in the presence of IgG isotype control or rituximab (1 μg/mL). Presence of fluorescently labeled DLBCL cells within the macrophages was assessed by immunofluorescence microscopy (arrows). Bar graphs present the mean phagocytic index, determined as the number of ingested cells per 100 macrophages. Error bars, ±SD of three independent replicates in a representative experiment. P values were determined using the two-sided t test: **, P < 0.01. Data in A–B are representative of three independent experiments.

Figure 5.

PIM inhibition potentiates CDC and ADCP of rituximab. Increased CDC triggered by rituximab in MEN1703-treated cells. DHL4 and RAJI were pretreated with DMSO or 1.5 μmol/L MEN1703 for 48 hours and then treated with rituximab (1 hour) in the presence of human AB Rh+ serum. Cell death was determined with PI staining and flow cytometry. Error bars, ±SD of two independent replicates in a representative experiment. Combination indexes (CI) were calculated using CompuSyn software. Statistical significance was determined using two-way ANOVA. B, PIM inhibition in DLBCL cells potentiates rituximab-dependent phagocytosis. After 72 hours of treatment with DMSO or MEN1703 (1 μmol/L), CFSE-labeled DHL4 and DHL6 cells were incubated with human peripheral blood–derived macrophages in the presence of IgG isotype control or rituximab (1 μg/mL). Presence of fluorescently labeled DLBCL cells within the macrophages was assessed by immunofluorescence microscopy (arrows). Bar graphs present the mean phagocytic index, determined as the number of ingested cells per 100 macrophages. Error bars, ±SD of three independent replicates in a representative experiment. P values were determined using the two-sided t test: **, P < 0.01. Data in A–B are representative of three independent experiments.

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Next, we evaluated the effects of PIM inhibition on natural killer (NK) cell effector function and coincubated DMSO- or MEN1703-treated RAJI target cells and effector NK cells for 4 hours in the presence of rituximab or type II anti-CD20 mAb, obinutuzumab, and assessed target cell viability. PIM inhibition had no effect on the efficacy of rituximab- or obinutuzumab-mediated antibody-dependent NK cytotoxicity (Supplementary Fig. S10B). However, MEN1703 potentiated cell death triggered by obinutuzumab alone (Supplementary Fig. S10B).

Macrophage-mediated, ADCP is another mechanism of anti-CD20 antibody activity in B-NHLs (29, 48). Because MYC has been shown to induce CD47, a SIRP1α ligand and a phagocytosis-inhibitory signal (49), we speculated that PIM inhibition-mediated MYC downregulation would decrease CD47 surface expression and thus enhance ADCP. In line with previous reports (49), MYC knockdown in DHL4 cells downregulated CD47 levels (Supplementary Fig. S11A). Likewise, PIM inhibition decreased CD47 surface levels in an MYC-dependent manner in DHL4 and DHL6 cells (Supplementary Fig. S11B and S11C). To investigate the effects of PIM inhibition on ADCP, we coincubated CFSE-labeled, DMSO- or MEN1703-treated DHL4 and DHL6 cells with monocyte-derived macrophages in the presence of IgG isotype control or rituximab (1 μg/mL), and assessed the number of DLBCL cells ingested by macrophages. MEN1703 and other pan-PIM inhibitors potentiated rituximab-dependent phagocytosis of lymphoma cells (Fig. 5B; Supplementary Fig. S11D).

Given these in vitro results indicating that PIM inhibition boosts anti-CD20 antibodies' effector mechanisms, we evaluated MEN1703 interaction with rituximab in vivo in SCID Fox Chase mice, intravenously inoculated with RAJI cells expressing firefly luciferase gene (28). Xenograft-bearing mice were treated with vehicle (H2O) and 10 mg/kg isotype control antibody (control animals), 50 mg/kg MEN1703, 10 mg/kg rituximab, or combination of MEN1703 and rituximab for 18 days. Compared with mice treated with the pan-PIM inhibitor or rituximab alone, MEN1703/rituximab combination significantly reduced tumor growth measured as supravital tumor cells bioluminescence (ANOVA, P < 0.0001; Fig. 6A and B), further demonstrating that PIM inhibition increases the efficacy of the anti-CD20 antibody-based therapies.

Figure 6.

PIM inhibition improves activity of rituximab in vivo. A, Increased rituximab activity in MEN1703-treated mice. SCID Fox Chase mice (n = 10 per group) were inoculated with RAJI cells expressing Red Firefly Luciferase and treated with H2O and isotype antibody (10 mg/kg, controls), MEN1703 (50 mg/kg), rituximab (10 mg/kg), or combination of MEN1703 and rituximab for 18 days (arrows). Bioluminescence was measured using IVIS Spectrum system and Living Image software. Each experimental group consisted of 8–10 mice. Statistical significance was determined using ANOVA with Tukey HSD post hoc tests: **, P < 0.01; ***, P < 0.001. B, Raw bioluminescence of mice injected with Raji cells expressing Red Firefly Luciferase (scale for bioluminescent signals at the bottom). Each image is an automatic superposition of white light exposure to visualize mice position in the instrument and bioluminescence detection taking place in the dark. In the images #5 and #6 from the left in the first, second, third, and fourth rows as well in image #4 in the first row, mice were separated with cardboard black shields to prevent bioluminescence spillover.

Figure 6.

PIM inhibition improves activity of rituximab in vivo. A, Increased rituximab activity in MEN1703-treated mice. SCID Fox Chase mice (n = 10 per group) were inoculated with RAJI cells expressing Red Firefly Luciferase and treated with H2O and isotype antibody (10 mg/kg, controls), MEN1703 (50 mg/kg), rituximab (10 mg/kg), or combination of MEN1703 and rituximab for 18 days (arrows). Bioluminescence was measured using IVIS Spectrum system and Living Image software. Each experimental group consisted of 8–10 mice. Statistical significance was determined using ANOVA with Tukey HSD post hoc tests: **, P < 0.01; ***, P < 0.001. B, Raw bioluminescence of mice injected with Raji cells expressing Red Firefly Luciferase (scale for bioluminescent signals at the bottom). Each image is an automatic superposition of white light exposure to visualize mice position in the instrument and bioluminescence detection taking place in the dark. In the images #5 and #6 from the left in the first, second, third, and fourth rows as well in image #4 in the first row, mice were separated with cardboard black shields to prevent bioluminescence spillover.

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PIM kinases in cancer cells exhibit prosurvival activity associated with their ability to modulate protein translation, transcription, cell metabolism, cell cycle, and apoptosis (10–15, 30, 50–52). In addition to their own oncogenic functions, PIMs cooperate with other lymphoma oncogenes, such as BCL6 or MYC (16, 17, 44). In DLBCLs, increased PIM levels are associated with inferior prognosis, particularly in the ABC subtype (51). For this reason, PIM kinases have garnered considerable attention as potential therapeutic targets in hematologic malignancies, including B-NHL.

Consistent with earlier reports (51), we demonstrate that PIMs are more abundant in the ABC subtype than GCB, although a significant fraction of GCB cases (44%) is also PIM-positive. This largely COO-independent PIM distribution might be explained by their tight association with MYC. Although GCB tumors more frequently exhibit MYC rearrangements, MYC expression and MYC transcriptional signature are the features shared by ABC (MCD/cluster 1) and GCB (EZB-MYC+/cluster 3) tumors (3, 6). MYC inhibition reduced PIM expression in DLBCL-derived cell lines, indicating that MYC functions as a positive regulator driving PIM expression. Because PIMs reciprocally increase MYC levels in DLBCLs, these proteins are linked by a feed-forward circuit that can be exploited therapeutically.

Despite the well-established role of MYC in driving cancer cell growth, direct MYC-targeting strategies have been largely ineffective, and no MYC-targeting agent has hitherto advanced to the clinical development. Thus, other indirect approaches might fill this therapeutic need. Given PIMs' role in accelerating MYC-driven lymphomagenesis, PIM inhibitors are rational candidates for this purpose. In line with reported partial redundancy of PIM isoforms in regulating MYC (16, 17, 37, 53), we show that only simultaneous depletion of all three PIMs markedly downregulated MYC and was associated with substantial cytotoxicity in DLBCL cells. Similar to genetic PIM inhibition, the pan-PIM inhibitor decreased MYC protein levels and attenuated MYC-dependent transcription. Consistent with the cell-cycle–promoting role of MYC, MEN1703 decreased expression of multiple genes engaged in cell-cycle control, including PLK1—the master mitotic regulator. Importantly, PLK1 is an adverse prognostic factor in DLBCL (54) and HGBLs (55), and an important node in the MYC proteomic network (55). As PLK1 stabilizes MYC via AKT–GSK3β pathway, PLK1 and MYC establish another feed-forward circuit. Thus, it is tempting to speculate that targeting PIMs and PLK1 would disrupt two orthogonal pathways that stabilize MYC and support DLBCL and survival. Importantly, PLK1-selective inhibitor volasertib exhibited activity at low-nanomolar range in DLBCL cell lines, prompting further combination studies.

In addition to their cell-intrinsic role, recent studies indicate that certain oncogenes, including MYC, also foster tumor immune privilege and immunosuppression (49, 56–58). Our previous studies demonstrated that PIM inhibition in Hodgkin lymphoma and CLL affects microenvironment composition and blocks tumor immune evasion (10, 50). In DLBCL and other lymphomas, modulation of tumor immunogenicity by PIM inhibitors is at least partially MYC-dependent. We demonstrate here that MYC represses the MS4A1 gene, encoding CD20 surface antigen, through two complementary mechanisms: direct repression of MS4A1 promoter and induction of MS4A1/CD20-targeting miR-222. Consistent with these mechanistic findings, MYC-rearranged tumors had lower levels of MS4A1 transcript. Complementary to the increased CD20 levels, PIM inhibitor decreased CD47 expression, a phagocytosis-inhibitory signal, in an MYC-dependent fashion. Consequently, PIM inhibition enhances the two major mechanisms of rituximab action, CDC and ADCP. Because ADCC is less dependent on the antigen density in lymphomas and is modulated by multiple additional mechanisms, such as KIR receptor–ligand interactions (59–61), it is not surprising that PIM inhibition did not further increase NK cell–mediated cell lysis. However, an afucosylated type II antibody, obinutuzumab, recognizing different epitope than rituximab and capable of directly inducing cell death, was more active when combined with PIM inhibitor, further highlighting the role of increased CD20 expression for the efficacy of anti-CD20 antibodies.

MYC-dependent mechanisms responsible for PIM inhibition–induced apoptosis in DLBCL cells are complemented by additional, MYC-independent mechanisms of cytotoxicity. PIM inhibition affects the activity of important regulators of protein translation and apoptosis, including 4EBP1, RPS6, and BAD. PIM blockade also reduces the expression of an antiapoptotic BCL2 family member, MCL1. As MCL1 drives resistance to selective BCL2 inhibitors (33, 62), these observations set the stage for combined targeting of PIMs and BCL2. In our preliminary studies, combination of MEN1703 and ABT-199 exhibited synergistic activity.

Taken together, we show that PIM blockage exhibits pleiotropic effects that combine direct cytotoxicity with an increased susceptibility to anti-CD20 antibody-based therapies (Fig. 7). Through a broad mechanism of action, PIM inhibition exhibits high potential for rational combinations with other targeted agents, such as venetoclax or volasertib. Given the tight association between MYC and PIMs expression in DLBCLs and the functional feed-forward loop linking these proteins, MYC expression might represent a potential biomarker selecting patients with DLBCL, and possibly other aggressive B-NHLs, who might benefit from a therapeutic approach including a PIM kinase inhibitor and rituximab. In particular, given the inhibition of MYC oncogenic program and synergy with venetocax, patients with HGBLs with MYC and BCL2 translocations (double-hit lymphomas) are a promising target group for the future clinical development of such combinatorial strategies.

Figure 7.

Proposed working model of cell-intrinsic and immunomodulatory effects of PIM blockade in DLBCL cells. PIM kinases support DLBCL cell growth, survival, and resistance to anti-CD20–based therapies through MYC-dependent and -independent mechanisms. PIM blockade impedes PIM-MYC feed-forward regulatory circuit, induces DLBCL cell death, and potentiates cytotoxic effector mechanisms of anti-CD20 antibodies.

Figure 7.

Proposed working model of cell-intrinsic and immunomodulatory effects of PIM blockade in DLBCL cells. PIM kinases support DLBCL cell growth, survival, and resistance to anti-CD20–based therapies through MYC-dependent and -independent mechanisms. PIM blockade impedes PIM-MYC feed-forward regulatory circuit, induces DLBCL cell death, and potentiates cytotoxic effector mechanisms of anti-CD20 antibodies.

Close modal

M. Szydłowski reports a patent for use of PIM kinase inhibitors to augment the efficacy of anti-Cd20 antibody-based therapies in hematologic malignancies and nonmalignant conditions pending. M. Pawlak reports personal fees from Oric Pharmaceuticals and 7N Sp. z o. o. Poland outside the submitted work. K. Brzózka reports personal fees from Ryvu Therapeutics during the conduct of the study; in addition, K. Brzózka has a patent for WO 2014/096388 licensed to Menarini Group. M.R. Green reports grants from Kite/Gilead, Sanofi, and Allogene, and other support from KDAc Therapeutics outside the submitted work. B. Chapuy reports grants from Gilead Sciences, personal fees from Jansen, Roche, AstraZeneca, BMS, Gilead, and Regeneron outside the submitted work. P. Juszczyński reports grants from Foundation For Polish Science and Polish National Science Center, personal fees from Ryvu Therapeutics during the conduct of the study; personal fees from Ryvu Therapeutics and Novartis outside the submitted work; in addition, P. Juszczyński has a patent for patent pending. No disclosures were reported by the other authors.

M. Szydłowski: Conceptualization, formal analysis, investigation, methodology, writing–original draft. F. Garbicz: Investigation, methodology. E. Jabłońska: Conceptualization and investigation. P. Górniak: Investigation and methodology. D. Komar: Investigation and methodology. B. Pyrzyńska: Investigation and methodology. K. Bojarczuk: Investigation and methodology. M. Prochorec-Sobieszek: Investigation. A. Szumera-Ciećkiewicz: Investigation. G. Rymkiewicz: Resources, contributed patient-derived samples. M. Cybulska: Investigation and methodology. M. Statkiewicz: Investigation and methodology. M. Gajewska: Investigation and methodology. M. Mikula: Investigation and methodology. A. Gołas: Investigation and methodology. J. Domagała: Investigation. M. Winiarska: Conceptualization. A. Graczyk-Jarzynka: Investigation. E. Białopiotrowicz: Investigation. A. Polak: Investigation. J. Barankiewicz: Resources, contributed patient-derived samples. B. Puła: Resources, contributed patient-derived samples. M. Pawlak: Data curation and visualization. D. Nowis: Conceptualization and investigation. J. Golab: Conceptualization. A.M. Tomirotti: Conceptualization. K. Brzózka: Conceptualization. M. Pacheco-Blanco: Resources and investigation. K. Kupcova: Resources and investigation. M.R. Green: Resources. O. Havranek: Resources and investigation. B. Chapuy: Conceptualization. P. Juszczyński: Conceptualization, formal analysis, supervision, and writing–original draft.

This work has been supported by the research grants from the Foundation for Polish Science (POIR.04.04.00-00-5C84/17-00), Polish National Science Centre (2016/22/M/NZ5/00668 and 2017/26/D/NZ5/00561), and iONCO grant from the Ministry of Science and Higher Education in Poland. The TMAs were constructed using the infrastructure funded by Operational Programme Innovative Economy 2007-2013, Priority II. R&D Infrastructure, Measure 2.3. Investments connected with development of IT infrastructure of Science (POIG.02.03.00-14-111/13). K. Kupcova, M. Pacheco-Blanco, and O. Havranek were supported by the following grant projects: GACR 20-01969Y (The Czech Science Foundation), PRIMUS/17/MED/9 and UNCE/MED/016 (both Charles University), and Progress Q26 (Ministry of Education Youth and Sports of the Czech Republic). The authors thank Dr. Zofia Pilch for outstanding help with animal experiments.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

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