Targeting the CXCR4 pathway using a novel anti-CXCR4 IgG1 antibody (PF-06747143) in chronic lymphocytic leukemia

The CLL-B cells were collected from blood samples at the Moores-UCSD Cancer Center in compliance with the Declaration of Helsinki and after approval of the UC San Diego Institutional Review Board (IRB) [24].Peripheral blood mononuclear cells (PBMC) from CLL patients were isolated using Ficoll-Hypaque gradient density centrifugation (Cat# 17-1440-03, GE Healthcare Life Science). For caspase activation assays, the CLL-B cells were purified by positive selection using Dynabeads CD19 pan B (Cat# 11143D, Invitrogen) and DETACHaBEAD CD19 (Cat# 12506D, Invitrogen) according to the manufacturer’s protocol. For the other assays, fresh or frozen PBMCs were used and cells were stained with CD19/CD5 antibodies for detection of double positive CLL-B cells.

Primary leukemic cells from CLL patients were cultured in RPMI supplemented with 10% heat-inactivated FBS (fetal bovine serum, Catalog # FB-02, Omega Scientific, Tarzana, CA) and 1% antibiotic at a density of 3 × 10⁵ cells per milliliter at 37 °C and 5% CO₂. The cells were either cultured in 96-well round bottom plates (Catalog # 3596, Corning, NY) alone or co-cultured with NK-tert stromal cells (RIKEN, Yokohama, Japan) at a ratio of 20:1 (CLL: stroma-NK-tert) in RPMI with 1% Penn-Strep and 10% FBS [25].

The CXCR4 phenotyping of CLL-B, stroma-NK-tert, normal B, and T cells was done by flow cytometry using a 1:50 dilution with rat anti-human CD184 (CXCR4) PE Mab (Catalog # 551966, clone:2B11, BD Biosciences). The isotype control antibody was PE Rat IgG_2b, κ (Catalog # 12-4031-83, clone: eB149/10H5, eBioscience).

The parental CXCR4 antagonist antibody, m15, was derived from immunization of Balb/c mice with CHO cells transfected with human CXCR4. The heavy and light chain variable domains of m15 were then cloned into human IgG1 or hinge stabilized IgG4 and light κ backbone, to generate chimeric m15-IgG1 and m15-IgG4. m15 was subsequently humanized by CDR grafting/affinity maturation and cloned into human IgG1/κ constant domains to create PF-06747143.

Experiments were performed on a Biacore^TM T200 surface plasmon resonance biosensor (GE Life Sciences). The binding to human CXCR4 was determined using human CXCR4-enriched lipoparticles (Integral Molecular) compared to null particles. Lipoparticles were diluted into 10 mM HEPES, 150 mM NaCl, 1 mg/mL BSA, pH 7.4 buffer to concentrations between 0.015 to 0.04 units/mL and captured for 5 min onto flow cells. A threefold dilution series of Fab was evaluated and dissociation was monitored for 10 min. The data were fit to a 1:1 Langmuir with mass transport model using Biacore T200 Evaluation Software Version 2.0.

The ability of PF-06747143 or m15-IgG1 to inhibit CXCL12-induced calcium flux was evaluated in human T cell leukemia Jurkat cells using the Fluo-NW Calcium assay kit (Life Technologies). Cells were plated in 384-well plates at 70,000 cells per well in quadruplicates and incubated with m15-IgG1 parent antibody and PF-06747143, upon stimulation with CXCL12 at 8 nM (EC80) (Invitrogen), for 110 min. Calcium flux was then measured for 95 s using a FLIPR Tetra (Molecular Devices).

Cell death was evaluated by flow cytometry analysis using CD19/CD5/Annexin V antibodies [26]. Specific induced cell death (SICD) calculation was used in order to discriminate the antibody/compound-specific induced cell death from background or spontaneous cell death observed in the vehicle-treated groups. The calculation of % SICD was performed using the following formula: % SICD = (Compound-induced cell death − Vehicle spontaneous cell death)/(100 − Vehicle spontaneous cell death) × 100.

m15-IgG1 was tested in combination with different standard of care (SOC) agents currently used for treatment of CLL. F-ara-A, bendamustine, rituximab, and ibrutinib were evaluated in combination with 200 nM of m15-IgG1. CLL-B cells were treated for 48 h at 37 °C either cultured alone or co-cultured with stroma-NK-tert cells. The combination data and level of synergism was analyzed using CompuSyn software (ComboSyn, Inc., NJ, USA). The data derived from this analysis were expressed as combination index (CI), which offers definition for additive (CI = 1), synergism (CI < 1), and antagonism (CI > 1) in drug combination [27].

For analysis of ADCC in B-CLL patient primary cells, the ADCC Reporter Bioassay kit from Promega (Catalog #G7010) was used, per instructions from the manufacturer. The ADCC Reporter Bioassay uses engineered Jurkat cells stably expressing the FcγRIIIa receptor, V158 (high affinity) variant, and a NFAT (nuclear factor of activated T cells) pathway response element driving expression of firefly luciferase as effector cells. The transfected Jurkat cell line was grown in RPMI containing G-418 sulfate solution (Catalog # V8091) and hygromycin (Catalog # 10687010, 50 mg/mL solution). The ADCC buffer (99.5% RPMI 1640 with L-glutamine and 0.5% super low IgG FBS) was prepared using RPMI supplemented with super low IgG defined fetal bovine serum (catalog # SH30898, Hyclone). The luciferase assay system was used as a readout (Catalog # G7940, Promega). Different concentrations of antibodies IgG1 control, PF-06747143, rituximab and obinutuzumab were added to the effector/target cell 1:1 ratio mixtures. A total of 75,000 for effector and target cells were incubated for 6 h at 37 °C in a humidified CO₂ incubator. Following incubation, the plate was equilibrated to ambient temperature for 15 min. Bio-Glo™ luciferase assay reagent was added and incubated at room temperature for 30 min. The luminescence was detected using an Infinite 200 Microplate Reader (Tekan), and the results are expressed in relative light units (RLU).

PF-06747143 was added to CLL-B cells (1 × 10⁶/mL) in RPMI media with 5% active human serum [28, 29] or inactivated human serum, which was incubated at 56 °C for 30 min. The heat-inactivated/normal human serum-treated cells were incubated for 4 h at 37 °C with increasing concentrations of PF-06747143. Cytotoxicity was determined by flow cytometry using CD19/CD5/Annexin V staining. % SICD was calculated according to the following formula: 100 × (% viable cells with inactivated serum − % viable cells with native serum)/(% viable cells with inactivated serum).

Cytoskeletal reorganization (F-actin polymerization) was evaluated in CLL samples activated by CXCL12 and treated with PF-06747143 or control agents [4].

PF-06747143 was assessed for its ability to inhibit CXCL12-induced chemotaxis in primary CLL-B cells derived from CLL patients using a transwell migration assay [30].

To evaluate the mechanism of cell death induced by PF-06747143, CLL-B cells were purified from patient-derived PBMCs and tested for caspase activation including caspases 3, 8, and 9 using the ApoTarget Caspase Colorimetric Protease Assay Sampler kit (Cat # KHZ1001, Invitrogen, Frederick, MD) according to the manufacturer’s instructions. Z-VAD-FMK, a caspase inhibitor (Cat # G7231, Promega Corporation), was used as control [31].

CLL-B cells were seeded at 2.5 × 10⁵/mL in RPMI media and treated with antibodies for 4 h at 37 °C and 5% CO₂ in 24-well plates. The generation of ROS was detected using dihydroethidium (HE) staining (Catalog # D1168, Sigma-Aldrich, St. Louis, MO) as described previously [32]. The samples were then analyzed by flow cytometry followed by data analysis using FlowJo software.

JVM-13 tumor cell line [33, 34], purchased from ATCC, was stably transfected with the luciferase gene. The cells were cultured in RPMI media with 10% FBS. To establish a JVM-13 disseminated model, 1 × 10⁶ cells per mouse were implanted via tail vein injection in female SCID beige mice (Charles River). Tumor burden was monitored via bioluminescence imaging (BLI) (IVISÒ 200) throughout the study. When the tumor burden (mean BLI) reached 7.2 × 10⁶ photons/s, on day 19, mice were randomly assigned into four groups and treated with (1) IgG1 negative control Ab or (2) PF-06747143, dosed subcutaneously at 10 mg/kg, once a week, for a total of 6 doses; (3) bendamustine, dosed intraperitoneally at 30 mg/kg, on days 19 and 20, followed by another 2-day treatment cycle 28 days later; and (4) combination of PF-06747143 and bendamustine. Mice were euthanized according to the IACUC guidelines once they developed disease-related symptoms such as hind leg paralysis.

The statistical analysis was carried out using GraphPad Prism software (v. 5.0c; San Diego, CA). The statistical differences for the mean values were analyzed using one way ANOVA and are indicated with *, p < 0.05; **, p < 0.01; ***, p < 0.001; and ****, p < 0.0001. Tumor model survival analysis was performed using Kaplan-Meier followed by a long-rank (Mantel-Cox) test.

Expression of CXCR4 was evaluated by flow cytometry in primary B-CLL cells from patients, as well as normal B, T, and stroma-NK-tert cells. A representative example of CXCR4 staining is shown in Fig. 1a. CXCR4 expression was higher in CLL-B cells compared to normal B and T cells. We then assessed CXCR4 expression levels in high- and low-risk CLL patient groups. CLL patients were stratified into high risk (ZAP-70 ≥ 20%, IgVH ≥ 98% homology) and low risk (ZAP-70 < 20%, IgVH < 98% homology) based on ZAP-70 expression and IgVH homology status [35]. The average mean fluorescence (ΔMFI) for CXCR4 expression in CLL samples ranged from 263.73 to 2401.7, regardless of high- or low-risk status. The ΔMFI in high-risk patients was 1497.23 ± 195.89 and in low-risk patients, it was 1533.73 ± 178.54. Upon comparison between low- and high-risk CLL patients, no significant difference in CXCR4 expression levels was observed (p = 0.8601). Overall, CXCR4 expression was significantly higher in CLL-B cells as compared to normal B or T cells (10- and 22-fold, respectively; p < 0.0001) (Fig. 1b). Furthermore, CXCR4 expression in leukemia and lymphoma cell lines including K562, MEC1, Namalwa, Raji, JeKo-1, and Jurkat showed a broad range of expression levels, with ΔMFIs of −2.35, 0.01, 73.41, 526.72, 725.45, and 1358.5, respectively (Additional file 1: Figure S1).

To determine the binding specificity and affinity of PF-06747143 and its parental antibody m15 to human (h) CXCR4, we employed surface plasmon resonance. The monovalent Fab fragments of each antibody were incubated with hCXCR4-enriched lipoparticles. The apparent equilibrium dissociation constant (K _D ) was 0.36 nM for PF-06747143 and 0.67 nM for the parental antibody, m15, indicating that both antibodies have potent and comparable affinity to hCXCR4 (Table 1). None of the Fabs bound to lipoparticles lacking hCXCR4 (data not shown). In a separate experiment, PF-06747143 was fluorescently labeled (PF-06747143-PE) and binding to CHO cells expressing hCXCR4 (CHO-hCXCR4) was compared to binding to cells transfected with empty vector (CHO-parental). PF-06747143-PE bound specifically to CHO cells expressing hCXCR4 (CHO-hCXCR4) but not to the CHO-parental cells (Fig. 2a), demonstrating selective binding to CXCR4-expressing cells.

Calcium flux is triggered upon activation of CXCR4 by its ligand, CXCL12. We next evaluated the ability of PF-06747143 and its parental antibody, m15, expressed as a chimeric human IgG1 antibody (m15-IgG1), to inhibit calcium flux induced by CXCL12. The Jurkat T cell leukemia line, which expresses high levels of CXCR4 (Additional file 1: Figure S1), was incubated with CXCL12 (EC₈₀ at 8 nM) to stimulate calcium flux. A titration of PF-06747143 and m15-IgG1 was performed. Both PF-06747143 and m15-IgG1 blocked CXCL12-induced calcium flux in a dose-dependent manner, with similar IC_50s of 1.41 and 1.13 nM, for PF-06747143 and m15-IgG1, respectively. These results show that both CXCR4 antibodies have potent and comparable CXCL12 antagonistic activity (Fig. 2b).

Next, we evaluated if bivalency was required for PF-06747143 to inhibit calcium flux. To this end, a bivalent form of PF-06747143, which has no constant Fc region [F(ab’)₂], and a monovalent form of the antibody [Fab], were generated and compared to PF-06747143, which is a bivalent full-length antibody (PF-06747143 FL). (Additional file 2: Figure S2). Similar CXCL12-induced calcium flux inhibition was observed for all three forms of PF-06747143 tested, indicating that the functional CXCL12 antagonistic activity is not dependent on bivalent binding or Fc constant region of the antibody.

m15-IgG1 was evaluated for its ability to trigger cell death upon binding to primary CLL-B cells expressing CXCR4 or to the MEC1 (CLL) cell line, which has no detectable CXCR4 expression (ΔMFI = 0.01) (Fig. 3a). Cells were incubated with increasing concentrations of m15-IgG1 or control IgG1 antibody and analyzed for cell death using flow cytometry. CLL-B cells underwent cell death upon treatment with m15-IgG1 (2–2000 nM) in a dose-dependent manner, while MEC1 cells did not show evidence of cell death, even in presence of high concentrations of the antibody (Fig. 3b), indicating that the CXCR4 antibody cell death is CXCR4 expression dependent.

To determine whether m15-IgG1 could induce cell death in presence of stromal cells, CLL-B cells were cultured with stroma-NK-tert cells in presence of increasing concentrations of m15-IgG1 and cell death was evaluated after 48 h. F-ara-A, an agent that inhibits DNA synthesis and is a cornerstone for the treatment for CLL patients [36] as well as AMD3100, a CXCR4 small molecule inhibitor [37], were evaluated for comparison. m15-IgG1 induced cell death of leukemia cells cultured either alone or in presence of stromal cell support (stroma NK-tert cells), demonstrating the ability of the antibody to induce cell death in presence of stromal cells (Fig. 3c). Similar results were observed for various B-CLL patients (Additional file 3: Figure S3).

Moreover, m15-IgG1 was more potent at inducing cell death than F-ara-A (p < 0.0001) in CLL-B cells (Fig. 3c). AMD3100, which binds and inhibits signaling through CXCR4, did not induce cell death in CLL-B cells (Fig. 3c), indicating that binding and inhibition of the CXCR4 pathway is not sufficient to trigger cell death.

Next, we sought to determine whether the Fc constant region or backbone of the CXCR4 antibody played a role in the ability to induce B-CLL cell death. We compared m15-IgG1 to m15-IgG4, which is an antibody with the same antigen-binding regions from m15-IgG1, cloned in a human IgG4 constant region. Results from these studies (Additional file 4: Figure S4) demonstrate that both antibodies are capable to induce cell death in CLL-B cells to similar degrees, with no significant difference between m15-IgG1 and m15-IgG4, cultured alone or co-cultured with stroma cell support (p > 0.05). This indicates that the antibody constant region does not play a role in this cell death mechanism.

In addition to the high-risk and low-risk prognosis markers ZAP70 and IgVH, the 17p deletion in the TP53 gene is also considered a strong independent adverse prognostic factor for survival and is associated with the short median treatment-free survival, in CLL patients with CLL [38].

To evaluate the ability of the CXCR4 antibody to induce cell death in leukemia cells from CLL patients with high risk (CLL-HR), low risk (CLL-LR), as well as with TP53 17pDel status, we treated samples from patients with these various genetic backgrounds with m15-IgG1 and evaluated cell death after 48 h of culture. Similar levels of cell death were induced by m15-IgG1 in CLL-LR, CLL-HR, TP53wt, or TP53mut/Del(17p) patient samples (Fig. 3d). This indicates that low-risk, high-risk, or TP53 statuses are not factors for sensitivity to m15-IgG1. Moreover, m15-IgG1-induced cell death was significantly higher than that with F-ara-A (p < 0.0001), which is known to be a TP53-dependent chemotherapy agent [39, 40]. F-ara-A did not induce cell death in TP53mut/Del(17p) even at supra-physiological concentrations (>3 μM).

Importantly, m15-IgG1 induced significantly lower levels of cell death in normal B and T lymphocytes compared with CLL samples (p < 0.0001) (Fig. 3d). These data suggest that m15-IgG1-induced cell death is dependent on the level of CXCR4 expressed in the cells.

To characterize the kinetics of cell death induced by the CXCR4 antibody, a washout experiment was carried out, where CLL-B cells were incubated with m15-IgG1 (200 nM) for increasing lengths of time, after which the antibody was washed out. Readouts were performed 48 h post-treatment initiation. Cell death was observed as early as 3 h and continued to increase until 48 h after m15-IgG1 was removed (Additional file 5: Figure S5). This effect was also independent of presence of stromal cells.

To determine if the CXCR4 antibody could offer additional benefit to available therapies, the m15-IgG1 antibody was evaluated in combination with SOC agents currently used in the treatment of CLL patients in the clinic. The percent cell death combinatorial effect was evaluated in both high-risk (HR) and low-risk (LR) CLL patient cells, in presence or absence of stromal cell support (Fig. 4). Overall, the presence of stromal cells did not appear to be a significant factor in the ability of m15-IgG1 to synergize with CLL SOC agents. Rituximab, an anti-CD20 antibody [41, 42], was tested at 2, 10, and 30 μg/mL. Synergistic cell death responses, in the range of 10–30%, were observed for both low- and high-risk patients, for all rituximab doses tested (Fig. 4a). Ibrutinib (IBM) or imbruvica, a BTK inhibitor [43], was tested at 0.1, 10, and 30 μM, and synergism was observed in 50–60% of high-risk patients and 50-80% of low-risk patients samples (Fig. 4b). Bendamustine, a chemotherapy agent derived from nitrogen mustard, commonly used in the treatment of CLL and lymphomas [44], was tested at 0.1, 30, and 90 μM. Synergistic effects were noted in 30–70% of high-risk samples and in 50–90% of low-risk patients (Fig. 4c). F-ara-A was tested at 0.3, 3, and 10 μM doses. Synergistic effects were observed in 30–60% of high-risk and in 50–70% of low-risk patient samples (Fig. 4d).

To establish if the humanized IgG1 CXCR4 antibody PF-06747143 and its parent antibody m15-IgG1 had comparable cell death activity, we performed a cell death study in leukemic B cells derived from B-CLL patient, comparing both antibodies. As shown in Fig. 5a, activity of both antibodies was very similar, with cell death induced at doses as low as 10 nM by both antibodies. Both antibodies induced similar degree of cell death, regardless of the presence of stromal cells.

Intact antibodies possess two binding regions, which allow for binding to two epitopes at the same time. To determine if binding to two epitopes (bivalency) was required for PF-06747143 to induce cell death, the following forms of the PF-06747143 were compared: bivalent full-length (PF-06747143 FL), bivalent with no Fc region (F(ab’)₂), and monovalent (Fab) (Fig. 5a). Comparable cell death (SICD) activity for the F(ab’)₂ form of PF-06747143 antibody and the intact antibody (FL) was observed, while the Fab form of the antibody did not induce cell death. These results indicate that bivalency, or ability to bind to two epitopes at once, is required for PF-06747143 ability to induce cell death.

To determine the mechanism through which PF-06747143 induced cell death, caspase activation was evaluated. PF-06747143 treatment of CLL-B cells did not induce significant activation of caspase 3, 8, and 9 in CLL-B cells (Additional file 6: Figure S6). In a follow-up experiment, Z-VAD-FMK (Z-VAD), an irreversible pan-caspase inhibitor [45, 46], rescued caspase-dependent apoptosis in CLL-B cells after treatment with etoposide or F-ara-A, which are both known to induce caspase-dependent cell death. However, Z-VAD failed to rescue the CLL-B cells treated with PF-06747143, indicating that the mechanism of cell death induced by PF-06747143 is caspase independent (Fig. 5b).

Antibody-induced non-apoptotic cell death in human lymphoma and leukemia cells had been previously shown to be mediated through a ROS-dependent pathway, named programmed cell death (PCD) [32]. This novel cell death mechanism does not rely on caspase activation and is dependent on homotypic cell adhesion triggered upon antibody binding, followed by actin redistribution, lysosome membrane permeabilization, and ROS activation. Since we showed that PF-06747143 induction of cell death is caspase independent and bivalency dependent, we hypothesized that PF-06747143 cell death mechanism might involve ROS activation. To test this hypothesis, CLL-B cells were treated with PF-06747143 or controls and evaluated for ROS production after 4 h of treatment. CLL-B cells treated with H₂O₂, the positive control, showed high levels of ROS compared to untreated control (****, p < 0.0001), as expected. Similarly, CLL cells treated with PF-06747143 showed a rapid increase in cell death and ROS production, compared to the untreated or to the IgG1 control Ab groups (****, p < 0.0001). Of note, PF-06747143 treated cells had ROS levels similar to that of the positive antibody control obinutuzumab (p > 0.05), which has been reported to induce PCD associated with increased ROS levels [32]. Rituximab and F-ara-A, which are not known to induce PCD, did not show a significant increase in ROS (Fig. 5c).

To determine if binding to two epitopes (bivalency) was required for PF-06747143 ability to induce cell death via ROS production, the bivalent full-length (FL), the bivalent deprived of Fc region (F(ab’)₂), and the monovalent (Fab) forms of the antibody were evaluated. Comparable ROS activity was observed for the F(ab’)₂ form of PF-06747143 antibody and the intact antibody (FL), while the Fab form did not induce cell death or ROS production (Fig. 5c). These results indicate that bivalency, or ability to bind to two epitopes at once, is required for PF-06747143 to induce cell death via a mechanism that involves ROS induction.

Actin polymerization, or cytoskeletal reorganization, is a surrogate marker of cancer cell migration and metastatic potential, induced by the interaction of CXCR4 with CXCL12 [47]. Therefore, we sought to characterize PF-06747143 role in CXCL12-induced actin polymerization in CLL-B cells. Stimulation with CXCL12 induced an average increase in actin polymerization of 450%, relative to baseline (100%) (Fig. 6a). PF-06747143 significantly inhibited actin polymerization in a dose response-dependent manner. PF-06747143 at 100 and 1000 nM inhibited CXCL12-induced F-actin polymerization to levels below baseline at 85% and 75%, respectively (****, p < 0.0001). The small molecule inhibitor of CXCR4, AMD3100, was less potent than PF-06747143 in this assay, with very limited activity at 4 μM, which is its clinically relevant dose. At 40 μM, it showed moderate inhibition of 180%, which was similar to the inhibition observed at the lowest dose of PF-06747143 (10 nM) (Fig. 6a).

Since PF-06747143 was shown to be a potent inhibitor of cytoskeletal organization, we evaluated its ability to inhibit CLL-B cell migration driven by its ligand, CXCL12, using a transwell migration assay. CXCL12 induced chemotaxis of CLL-B cells, with an average increase >500% over baseline (Fig. 6b). PF-06747143 (10 nM–1 μM) significantly inhibited cell migration in a range from 40 to 80%, relative to CXCL12 induction only (****, p < 0.0001). The effect was dose dependent. CLL-B cells treated with AMD3100 (40 μM) had similar migration inhibition activity as PF-06747143 lowest dose (10 nM), but no significant effect was observed with AMD3100 at 4 μM (Fig. 6b).

Therapeutic antibodies may rely on their constant region (Fc domain) ability to induce target cell killing via immune-mediated effector functions, such as ADCC and CDC to achieve efficacy. Human IgG1 and IgG3 antibodies have Fc domain sequences that can mediate potent effector functions, while human IgG4 and IgG2 antibodies display little or no effector function [48]. PF-06747143 is a humanized IgG1 antibody; therefore, ADCC and CDC studies were performed to characterize the antibody Fc-driven cytotoxic activity on CLL patient cells.

CDC cell death mechanism depends on the interaction between active serum complement proteins with the Fc region of the antibody, upon binding to the target cells. To evaluate if PF-06747143 could induce CDC in CLL-B cells, it was tested in the presence of active or inactive serum protein. As shown in Fig. 7a, PF-06747143 significantly (p < 0.0001) increased cytotoxicity in presence of active complement, relative to inactive complement, and this response was antibody dose dependent.

In the case of ADCC, effector cells bind to the Fc region of the antibody, when bound to target cells, and trigger cell lysis. To characterize the Fc-mediated ADCC activity of PF-06747143, CLL-B cells were incubated with the antibody in presence of effector cells. Two therapeutic antibodies, rituximab and obinutuzumab, that bind to CD20 and rely on ADCC as their main mechanism of action were used as positive controls. As expected, rituximab and obinutuzumab induced ADCC of CLL-B cells to high levels. PF-06747143 had comparable ADCC activity to that of the CD20 antibodies, inducing significant ADCC, when compared to the negative IgG1 control antibody (Fig. 7b). Taken together, these data demonstrate that PF-06747143, a humanized IgG1 antibody, has strong Fc-driven cytotoxic-dependent activity, leading to elimination of CLL-B cells.

To evaluate the role of the human IgG backbone in the ADCC activity of a CXCR4 antibody, we compared PF-06747143, m15-IgG1, and m15-IgG4, generated by cloning of the m15 variable domain in a human IgG4 backbone. The ADCC assay was performed using the CLL tumor cells (JVM-13), in presence of NK92158V effector cells. The m15-IgG1 antibody showed strong cytotoxicity, when compared to no activity with the m15-IgG4 antibody (Fig. 7c). This is in agreement with the expected diminished ADCC activity of an IgG4 antibody. In addition, PF-06747143 exerted similar cytotoxicity to that of m15-IgG1, confirming that the parent antibody (m15-IgG1) and its humanized antibody (PF-06747143) have comparable ADCC functional properties.

To determine if PF-06747143 is effective in eliminating CLL tumor cells in vivo, a disseminated model in which the tumor cells were implanted intravenously and migrate spontaneously to various sites in the body, including the BM and lymph nodes, was used (Fig. 8). Activity of PF-06747143 was evaluated as a monotherapy as well as in combination with bendamustine, a SOC agent approved for CLL treatment. Bioluminescence imaging performed on day 26 showed that PF-06747143 treatment as a monotherapy decreased tumor burden compared to hIgG1 control Ab and bendamustine groups. A combinatorial effect was observed when PF-06747143 was given with bendamustine (Fig. 8a). PF-06747143 strong tumor growth inhibition also resulted in significant survival benefit at the end of the study, day 75. The hIgG1 negative control Ab and bendamustine groups had median survival of 27 and 40 days, respectively, while PF-06747143 monotherapy group lived significantly longer, with median survival of 68.5 days (p < 0.0001 vs IgG1 control Ab and p < 0.0094 vs bendamustine) (Fig. 8b). The median survival for the PF-06747143 and bendamustine combination group was not reached by the last day of the study (day 75), demonstrating a combinatorial effect between these two agents. Taken together, these results show that PF-06747143 reduces BM and lymph node tumor burden and improves survival, as a monotherapy in a CLL disseminated tumor model. Moreover, its efficacy is improved in combination with bendamustine.