Healthy-donor (HD) source leukocytes were purchased from Versiti Blood Services (Columbus, OH, USA). Whole-blood samples from untreated, low-count (<60,000 cells/μL) CLL patients were provided by the Leukemia Tissue Bank of The Ohio State University Comprehensive Cancer Center, obtained according to the Declaration of Helsinki and under protocols approved by the Institutional Review Board at The Ohio State University. Peripheral blood mononuclear cells (PBMCs) were isolated from each sample using lymphocyte separation medium (Corning, Corning, NY, USA), followed by density gradient centrifugation, and then washed with RPMI (Gibco, Thermo-Fisher Scientific, Whaltham, MA, USA). For PBMC experiments, cells were counted using the Cellometer Auto 2000 (Nexcelom Biosciences, Lawrence, MA, USA) and resuspended in RPMI media supplemented with 10% FBS (VWR, Radnor, PA, USA), 2 mM L-glutamine (Gibco) and 100 U/mL Penicillin-Streptomycin (Gibco) at 10x10⁶ cells/mL.

To obtain HD or CLL-patient monocytes (CD14⁺ cells), as well as CLL cells (CD19⁺), PBMCs were incubated with magnetic CD14⁺ or CD19⁺ selection beads (Miltenyi Biotec, Waltham, MA, USA) for 15 minutes on ice. Following this, cells were centrifuged and resuspended in MACS buffer (Miltenyi Biotec) and added to an LS selection column. After three washes, cells were recovered from the column, washed with incomplete RPMI, counted, and resuspended at 3x10⁶ cells/mL in supplemented RPMI media.

For stimulation, cells were plated and treated with L18-MDP (MDP) (InvivoGen, San Diego, CA, USA). A concentration of 1 µg/mL was selected after testing a concentration-response curve in isolated monocytes, and is similar to a previous report (49). Cells were incubated at 37°C, 5% CO₂, for 24 hours unless indicated otherwise. For inhibition experiments, monocytes were treated with DMSO (as inhibitor vehicle control) or: Bay 11-7085 at 5 µM (Millipore Sigma, St. Louis, MO, USA), trametinib at 10 nM (Selleck Chemicals, Houston, TX, USA), or SB202190 at 1 µM (Selleck Chemicals) for 30 minutes before MDP stimulation. Dimethyl sulfoxide (DMSO; Sigma Aldrich, St. Louis, MO, USA) was used to prepare stock solutions and the final concentration of DMSO was under 1 μL/mL.

Cultured cells were collected and then mRNA extracted using the Norgen Total RNA extraction kit (Norgen Biotek, ON, Canada), according to the manufacturer’s instructions. Collected mRNA was quantified using the nanodrop ND100 (Thermo-Fisher Scientific), and then reverse transcribed using the high-capacity reverse transcriptase kit (Applied Biosystems, Thermo-Fisher Scientific) into cDNA. Then, quantitative real-time PCR (qPCR) was done using SYBR green (Applied Biosystems) and the QuantStudio™ 3 machine (Life Technlologies). RCN (relative copy number) values were calculated for each target, normalizing to a GAPDH housekeeping control according to the equation RCN = 2 ^(-ΔCt) x 100, where ΔCt = Ct_problem – Ct_GAPDH) (50). Primers used are shown in Table 1 .

Cells were collected and lysed using 70 µL TN1 lysis buffer for every 1 x 10⁶ cells, as previously described (51). Protein was quantified using the DC Protein Assay according to manufacturer’s instructions (Bio-Rad, Hercules, CA, USA). Samples were boiled with 5x SDS sample buffer (150 mM Tris; 11.5% SDS; 0.05% bromophenol blue; 50% glycerol; 1% 2-mercaptoethanol). Samples were then size-separated using SDS-PAGE. Proteins were transferred to a nitrocellulose membrane using the Trans-Blot Turbo Transfer System (Bio-Rad). Membranes were blocked with the LI-COR blocking solution (LI-COR Biotechnology, NE, USA) and incubated with primary antibodies overnight at 4°C in LI-COR antibody diluent. The following primary antibodies were utilized: phospho-p42/44 (polyclonal, product #9101; or clone D13.14.4E; Cell Signaling Technology), phospho-MAPKAPK2 (polyclonal, product #3041; Cell Signaling Technology), phospho-p38 (clone D3F9; Cell Signaling Technology), phospho-p65 (clone 93H1; Cell Signaling Technology) mouse anti-human GAPDH (clone D4C6R; Cell Signaling Technology), rabbit anti-human FcϵRIγ (product 06-727; Upstate, Sigma-Aldrich), and mouse anti-human calreticulin (clone FMC 75, Enzo Life Sciences, Farmingdale, NY, USA). After probing with the corresponding fluorescently-conjugated secondary antibody (IRDye 800CW goat anti-rabbit, or IRDye 680RD goat anti-mouse antibodies; LI-COR) for 1 hour at room temperature, membranes were developed using the Odyssey CLx (LI-COR). For densitometry analysis, Fiji was used (52).

After PBMC or monocyte stimulation, cells were collected and washed with PBS. Then, cells were transferred into v-bottom plates and incubated with whole human IgG (Jackson ImmunoResearch Labs, West Grove, PA, USA) at 10 µg/mL in PBS for 15 minutes on ice. After Fc block, the specific antibody cocktail was added ( Table 2 ). Samples were acquired using the LSR-Fortessa (BD Biosciences) at the Flow Cytometry Shared Resource at The Ohio State University Comprehensive Cancer Center and analyzed using FlowJo version 10.7.2 (Ashland, OR, USA). Supplementary Figure 1 shows an example of the gating used in monocyte analysis.

Cultured cells were centrifuged, and cleared supernatants collected and used for ELISA with the Human TNFα DuoSet ELISA kit (R&D Systems, Minneapolis, MN, USA) according to manufacturer’s instructions. For acquisition, the BioTek Synergy HTX plate reader was used (Agilent, Santa Clara, CA, USA).

To determine antibody-mediated binding of targets, opsonized sheep red blood cells (sRBCs; Colorado Serum Company, Denver, CO, USA) were prepared as previously described (53). sRBCs where washed with PBS until the supernatant appeared clear. Then, 2 μL of PKH26 Red Fluorescent Dye (Sigma-Aldrich), diluted in Diluent C, were added to the cell pellet and thoroughly mixed. The reaction was stopped with a 1:1 volume of FBS, and sRBCs were washed with PBS until clear. Fluorescent sRBCs were then opsonized with rabbit anti-sheep antibody (Sigma-Aldrich) on ice for 2 hours before washing.

Upon monocyte stimulation with MDP at 1 μg/mL for 24 hours, cells were used for rosetting analysis, as previously described (53). Briefly, monocytes were divided and incubated with 1 uL of packed, opsonized or non-opsonized, fluorescent sRBCs. Cells were mixed and spun down, then incubated on ice for 1 hour. After incubation, cells were fixed with 1% paraformaldehyde in PBS for 15 minutes before analysis. The number of bound targets per 100 monocytes, as well as the percent of active cells (monocytes with bound sRBCs) were counted through microscopy using the Olympus BX41 (Olympus Lide-Science/Evident, Tokyo, Japan). Samples were counted in a blinded fashion by two different subjects. The number of bound non-opsonized targets in control samples was subtracted from the number of bound opsonized targets.

sRBCs were stained with the pH sensitive Protonex Red 600 dye (AAT Bioquest, Pleasanton, CA) at 4 μM in Hanks Buffered Saline Solution (HBSS; Gibco). The staining was done for 30 minutes at 37°C and then sRBCs were washed 3 times with PBS. sRBCs were then opsonized with antibody as stated above.

For measuring phagocytic activity, monocytes were treated with DMSO or pathway inhibitors for 30 minutes before stimulation with MDP at 1 μg/mL for 24 hours. Then, cells were washed and stained for contrast with carboxyfluorescein succinimidyl ester (CFSE, Thermo-Fisher Scientific) at 0.5 μM in PBS for 10 minutes at room temperature; the reaction was stopped with FBS and cells spun down. Cells were resuspended in complete media to perform the assay. Monocytes were mixed with 1 μL of packed, fluorescently-labeled, opsonized sRBCs for 45 minutes at 37°C, then washed and fixed with paraformaldehyde at a final concentration of 1%. The amount of phagocytosed sRBCs was counted per 100 cells under the microscope, and the number of cells with ingested events was also recorded as active phagocytes. The counts were done independently by two or three different subjects in a blinded fashion and averages calculated. As control, phagocytosis of non-opsonized, fluorescent sRBCs was checked for negativity. Some donors and results are shared among different phagocytosis assays.

Mice were housed in a vivarium at The Ohio State University, following all institutional guidelines. All experimental procedures were approved by The Ohio State University Institutional Animal Care and Use Committee.. An adoptive transfer Eμ-TCL1 mouse model of CLL was used (54, 55). Briefly, 1x10⁷ splenocytes from CLL-burdened Eμ-TCL1 mice were used to engraft C57BL/6 mice. After 2 weeks of disease development, treatments (5 mg/kg of MDP and/or 1 mg/kg αCD20 (Invivogen, San Diego, CA, USA, catalog #mcd20-mab10 -) were delivered intraperitoneally three times weekly, for 2 weeks. Engrafted C57BL/6 mice, left untreated, and nongrafted C57BL/6 mice served as controls. Mice were euthanized using a CO₂ chamber, followed by cervical dislocation. Spleens and peripheral blood were collected. For spleens, the tissue was weighed and disrupted using a 100 μm cell strainer (Corning); erythrocytes from both blood and disaggregated spleens were eliminated by centrifugation using lymphocyte separation medium. Cells were washed, resuspended in PBS and used for flow cytometry staining.

For flow cytometry staining, cells were placed in a v-bottom plate and stained as above using the antibodies listed for Panels 5 and 6 ( Table 2 ). For monocyte and macrophage phenotyping (Panel 6, iNOS and EGR2), we used the Foxp3 Intracellular staining kit (Thermo-Fisher Scientific) according to the manufacturer’s instructions, as described previously (56–58). For acquisition, Absolute Count Beads (BioLegend) were used.

Graphs were prepared using Excel (Microsoft Corporation, Redmond, WA). For two-group comparisons, a one-tailed t-test analysis for paired samples was performed using Excel. For multi-group comparisons, data were analyzed with the mixed effects model using SAS 9.4 (SAS Institute, Cary, NC, USA). Statistical analyses included only the engrafted groups (vehicle, MDP, αCD20 and combination) for mouse experiments, nongrafted controls were not compared. Data in graphs is shown as mean + standard deviation (S.D.).

Monocytes and macrophages are well-known responders to NOD2 agonists (40–42). To confirm this, we first treated PBMCs from HDs with MDP at 1 μg/mL for 24 hours. Flow cytometry was then used to measure different activation markers in monocytes and lymphocytes to determine which populations show an efficient response to NOD2 agonists. As shown in Figure 1A , monocytes showed a significant increase in CD86 and HLA-DR. Concurrently, there was a significant decrease in the M2-associated marker CD163. These results suggest that monocyte activation was skewed toward a proinflammatory phenotype. In T cells, we found that NOD2 agonist treatment increased CD86 in CD4⁺ T cells, which may suggest a slight activation of this population, as has been suggested previously (59) ( Supplementary Figure 2A ). B and NK cells were also examined, but only a reduction in NK-cell CD107a was observed ( Supplementary Figures 2C, D , respectively). Since changes on monocytes were significant, we decided to further test the effects of NOD2 in isolated monocytes. 24 hours following stimulation of NOD2, monocytes produced robust levels of TNFα, measured in the supernatants ( Figure 1B ). These results confirm earlier reports that monocytes are activated by NOD2 agonists (40, 49).

We next tested whether NOD2 stimulation would increase the phagocytic ability of monocytes, specifically in the context of FcγR-mediated responses. For this, we treated monocytes with MDP at 1 µg/mL for 24 hours and then incubated them with fluoresceinated, antibody-opsonized sRBCs. Here, cells were subjected to rosetting (initial binding stage of phagocytosis) and ingesting assays. Rosetting was done by incubating monocytes with the sRBCs at 4°C while incubation at 37°C was done in parallel to measure ingestion. As shown in Figure 1C , there was a significant increase in antibody-mediated binding (rosetting) in agonist-treated monocytes compared to controls, which is also reflected in the number of cells with bound sRBCs. Likewise, the number of ingested, opsonized sRBCs was significantly higher in treated monocytes ( Figure 1D ). There was also a significant increase in the number of actively phagocytosing monocytes. Taken together these results indicate that NOD2 stimulation leads to an increase in FcγR-mediated phagocytosis.

To test whether the increase in phagocytosis could be accounted for by a change in FcγR expression, we isolated monocytes from HDs and treated for 24 hours with vehicle or MDP at 1 µg/mL. Following this, RNA was isolated and transcripts for FcγRIa, FcγRIIa, FcγRIIIa, FcϵRIγ and FcγRIIb (the inhibitory FcγR) were evaluated by qPCR. Results ( Figure 2A ) showed that there was a significant increase in the expression of FcγRIIa and FcγRIIIa. Contrary to our expectation, however, RNA expression of FcγRI was significantly lower in MDP-treated monocytes. Transcripts of FcϵRIγ were significantly increased by MDP treatment ( Figure 2B , upper panel). Changes in FcγRIIb expression were not significant ( Supplementary Figure 3A ).

We also measured surface expression of FcγRs by flow cytometry. Consistent with an increase in transcripts, there was a significant increase in surface expression of FcγRIIa and FcγRIIIa in isolated monocytes after 24 hours with NOD2 agonist treatment ( Figure 2C ). Surprisingly, the surface expression of FcγRI was also significantly increased despite the observed reduction in transcript ( Figure 2C ). In addition to the increase in the geometric mean of expression of the activating FcγRs, we also observed a slight but significant increase in the percent of monocytes positive for FcγRI, as well as an increase of the monocyte population expressing FcγRIIIa ( Supplementary Figure 4 ). This indicates a phenotypic change into intermediate or non-classical monocytes after NOD2 stimulation (60).

Because surface expression of FcγRI depends on FcϵRIγ (7, 61), we also measured protein levels of FcϵRIγ using immunoblot analysis under the same stimulation conditions. Of note, we detected two bands when measuring FcϵR1γ, which has been seen by other authors previously and may correspond to either dimers or phosphorylation of the protein (7, 62); thus, densitometry analysis was done for both. As shown in Figure 2B (middle panel and lower-panel graph), MDP significantly increased protein levels of FcϵRIγ. Hence, despite reducing the transcript of FcγRI, MDP treatment led to greater surface expression of all three activating FcγRs on monocytes.

Previous studies have demonstrated that NF-κB, MEK and p38 are activated downstream of NOD2 (37–39). We therefore explored which of these pathways was required for MDP-mediated changes in FcγR expression and function. For this, HD monocytes were isolated and each of these three pathways was inhibited using pharmacologic inhibitors prior to treating the monocytes with MDP. To verify efficacy of the inhibitors, immunoblot analysis was performed with phospho-specific antibodies for NF-κB, ERK and MAPKAPK2 (downstream of p38), measured at 24 hours post-treatment ( Supplementary Figure 5 ). [Of note, no visible activation of p38 was detected at this time point after MDP stimulation, which may be due to p38 activity peaking earlier after stimulation (63–65)]. qPCR and flow cytometry were done to measure FcγR expression. As shown in Figure 3A and quantified in Supplementary Figure 6 , MDP-driven increases in FcγR surface expression were prevented by inhibition of NF-κB and p38 but not by inhibition of MEK. Changes in transcript after MDP were also largely prevented by inhibition of NF-κB and p38 ( Supplementary Figure 7 ). Consistent with the effects of these inhibitors on changes in FcγR expression, the MDP-mediated increase in phagocytosis was prevented with NF-κB and p38 inhibition ( Figure 3B ). Taken together, these data demonstrate that NF-κB and p38 are required mediators for enhancing FcγR expression and function downstream of NOD2.

CLL-patient monocytes are known to have altered phenotype compared to HDs (17). Thus, to test whether the effects of NOD2 stimulation seen in HD monocytes were reproducible in CLL-patient monocytes, we isolated monocytes from deidentified peripheral blood samples obtained from CLL patients and measured NOD2 expression using qPCR. As shown in Figures 4A, B , NOD2 was expressed in CLL-patient monocytes at similar transcript levels to HD monocytes. As with HD monocytes, CLL-patient monocytes were able to respond to MDP by producing TNFα after 24 hours of stimulation. Further, MDP treatment of CLL-patient monocytes also enhanced surface expression of FcγRI and FcγRIIIa, observed by flow cytometry ( Figure 4C ), and expression of FcϵRIγ, measured by immunoblot ( Figure 4D ). In addition, the transcriptional levels of FcγRIIb in CLL-patient monocytes remained unchanged after NOD2 stimulation ( Supplementary Figure 3B ). However, in contrast to HD monocytes, MDP treatment did not increase surface expression of FcγRIIa in CLL-patient monocytes after MDP treatment ( Figure 4C , middle graph). However, consistent with an overall increase in activating FcγRs, MDP treatment of CLL-patient monocytes led to a significantly higher level of phagocytosis ( Figure 4E ). In contrast to HD monocytes, the same increase in rosetting was not seen in patient monocytes ( Figure 4F ), although monocytes from 3 of the 4 donors showed higher levels with MDP treatment. This suggests that despite a lower overall response than that seen in HD monocytes, CLL-patient monocytes are capable of responding to MDP by increasing FcγR expression, cytokine production and phagocytic ability.

In addition to monocytes, the effect of MDP on CLL cell activation and proliferation in patient samples was examined, as this could be a potential and undesirable side effect. PBMCs from CLL patients were treated with MDP for 24 hours, then expression of CD86 in CLL cells was measured using flow cytometry. Although there was some increase in CD86, it was not significant ( Supplementary Figure 8A ). We next isolated B/CLL cells and treated with vehicle or MDP, counting cell numbers and viability at different time points. As shown in Supplementary Figures 8B, C , the viability of CLL cells was not affected by MDP. These results suggest that NOD2 activation does not affect CLL cell viability. Taking cell number and percent viability together, results also suggest that MDP did not directly affect CLL-cell proliferation.

To test the effects of NOD2 stimulation on FcγR function in vivo, we used a mouse model that mimics human CLL (66, 67). C57/BL6 mice were engrafted with splenocytes from diseased Eμ-TCL1 mice, as described in the methodology. Disease was allowed to develop for 2 weeks. Animals were either treated with vehicle, treated with MDP, treated with a suboptimal dose of αCD20 antibody or treated with MDP plus αCD20 antibody in combination. Mice were sacrificed after 2 weeks of treatment. Healthy, nonengrafted mice were included as controls. CLL loads in peripheral blood and in the spleens were assessed. Results showed that white blood cell (WBC) counts in peripheral blood were significantly reduced by each single treatment, and dual treatment led to greater reduction than either single treatment ( Figure 5A ). There was also a significantly lower percentage of CD5⁺/CD19⁺ CLL cells after antibody treatment, and dual treatment led to a lower percentage than either single treatment ( Figure 5B ).

In addition to the leukemic load in the blood, we analyzed the spleen, as CLL cells tend to accumulate in this organ (68). Spleen weights were also significantly lower in mice treated with antibody, but MDP plus antibody led to spleen weights significantly lower than those for untreated or for either single treatment ( Figure 5C ). The number of CLL cells per milligram of spleen weight ( Figure 5D ) was also significantly lower with dual treatment compared to either single treatment or vehicle control. Furthermore, this was also observed in the percent of CD19⁺CD5⁺ cells from the viable CD45⁺ population (data not shown). An image of the spleens is shown in Figure 5E . These results suggest that the combination of NOD2 agonist plus therapeutic antibody may be more effective than antibody alone.

In addition to the leukemic burden, we tested whether MDP combined with antitumor antibody could modulate the phenotypes of monocytes and macrophages in vivo. For this, cells from the spleen were obtained and stained for flow cytometry to detect M1- and M2-related markers on monocytes and macrophages. Cells were gated as CD45⁺CD11b⁺Ly6G^- and further classified by expression of MHC-II and/or F4/80 as previously done (56). Percentages of cells expressing iNOS (M1) or EGR2 (M2) (58) were measured to assess M1 or M2 polarization. Results from nonengrafted, healthy control mice were also collected.

In MHC-II⁺ populations either negative or positive for F4/80 (mainly monocytes or macrophages, Figures 6A, B , respectively) there was a significant upregulation of iNOS with either single treatment, and dual-treatment led to significantly higher M1 marker iNOS than either single treatment or vehicle. Neither single treatment significantly downregulated the percent of myeloid cells expressing the M2 marker EGR2, but dual-treatment significantly reduced EGR2 compared to vehicle control and compared to either single treatment. In the F4/80⁺ group ( Figure 6C ), all treatments led to significantly higher iNOS versus vehicle, and combination treatment led to significantly higher iNOS than antibody-alone treatment.. Taken together, these data suggest that MDP/antibody combination treatment not only decreases CLL burden but also shifts myeloid phenotypes towards a more proinflammatory, and potentially M1-like state.