The extracellular domain is necessary for full transformation by FGFR2 fusions. To ascertain the role of FGFR2-fusion ECDs, we developed BaF3 and NIH3T3 fibroblast cell lines expressing FGFR2 fusions: FGFR2-BICC1 (the most common fusion found in ICC), FGFR2-AHCYL1, and FGFR2-PHGDH proteins. Expression of FGFR2 fusions resulted in IL-3–independent growth of BaF3 cells and transformation of NIH3T3 cells (Figure 1A and Supplemental Figure 1A; supplemental material available online with this article; https://doi.org/10.1172/JCI182417DS1); growth of these cells was attenuated by the FGFR inhibitor (FGFRi) infigratinib (Supplemental Figure 1A). Transformation and proliferation of the FGFR2-fusion expressing lines were further enhanced by the FGFR2 ligand FGF10 (Figure 1, A and B). To measure receptor dimerization, we utilized NanoBiT assays that detect protein interactions by proximity-mediated luciferase complementation (18) (Figure 1C). We validated expression of full-length FGFR2-WT and FGFR2-AHCYL1 coupled to the NanoBiT fragments LgBiT and SmBiT (Supplemental Figure 1, B and C) and assayed luminescent activity upon coexpression. Complementation-based luciferase activity of FGFR2 fusions was significantly higher than that of FGFR2-WT (Figure 1D), indicating ligand-independent dimerization. Nonetheless, addition of FGF10 significantly enhanced receptor dimerization of FGFR2-WT and FGFR2-AHCYL1 (Figure 1D). These data indicate that the FGFR2-fusion ECD is functional and enhances fusion receptor activation through ligand-mediated dimerization.

The extracellular domain is necessary for full transformation by FGFR2 fusions. (A) Transformation assays showing cumulative population doublings in BaF3 cells expressing FGFR2-PHGDH (12 days) and FGFR2-AHCYL1 (15 days) with or without FGF10 (100 ng/mL) or IL-3 (10 ng/mL), as indicated (n = 3). (B) Growth of BaF3 cells expressing FGFR2-PHGDH and FGFR2-AHCYL1 analyzed by CellTiter-Glo at 5 days after IL-3 removal (n = 5). (C) Illustration of the dimerization assay using FGFR2-fusion NanoBiT constructs. Large BiT and Small BiT subunits are fused to the C-terminus of FGFR2 fusions. SP, signal peptide; TM, transmembrane; KD, kinase domain; FP, fusion partner;PM, plasma membrane. (D) HEK-293T cells expressing FGFR2-WT and FGFR2-AHCYL1 fused to LgBiT alone or fused to LgBiT and SmBiT were used to quantify the receptor dimerization in the presence or absence of FGF10. Shown is the fold increase over FGFR2-LgBiT activity alone (n = 5). (E) Illustration of FGFR2-BICC1 constructs with D1 (Ig1), D2 (Ig2), D3 (Ig3), or D2+D3 (Ig2+Ig3) deletions in the ECD. (F) Representative images of focus formation assays of NIH-3T3 cells expressing FGFR2 WT or the indicated ECD deletion variants. Scale bar: 250 μm. (G) Quantification of number of colonies from F (n = 6). (H) Growth of NIH3T3 cells overexpressing FL, D1, D2, D3, and D2+3–deleted FGFR2-BICC1 constructs as measured by Incucyte at 5 days after plating (n = 5). (I) Dimerization of FGFR2-BICC1 D1, D2, D3, or D2+D3 ECD–deleted constructs in HEK-293T cells compared with full-length FGFR2-BICC1. Fold change in luminescence over FGFR2-WT–LgBiT is shown (n = 5). (J) Immunoblotting of FGFR2 downstream pathway effectors in HEK-293 cells expressing FGFR2-BICC1 ECD deletion constructs. All data are mean ± SEM. Data are representative of 1 out of 3 independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 by 1-way ANOVA multiple comparisons.

Next, we asked whether subdomains of the ECD were required for FGFR2-fusion dimerization, cell growth, and transformation. To this end, we generated FGFR2 fusions with deletions of the D1, D2, and D3 subdomains (Figure 1E). Since the D2 and D3 domains are necessary and sufficient for ligand binding, we also generated D2 + D3 deletion constructs. Each ECD deletion was expressed in NIH3T3 cells lacking endogenous FGFR2, and we performed colony formation and proliferation assays. Comparable expression of each construct was observed via immunoblotting (Supplemental Figure 1, D and E). D1, D2, D3, and D2 + D3 deletions each reduced growth (35%–77% growth inhibition) and transformation capacity (36%–50% reduction) compared with full length (FL) FGFR2-fusion expressing cells (Figure 1, F–H). Specifically, deletion of D2 of the FGFR2 ECD had a pronounced impact on cell growth and transformation, suggesting that D2 may play a prominent role in the oncogenicity of FGFR2-BICC1. Thus, the ECD is required for full transformation by FGFR2 fusions.

Signaling by FGFR2-WT is initiated by binding of FGF ligands to the D2 and D3 domains leading to receptor dimerization and activation. To test the domain requirement for activity of FGFR2 fusions, we utilized NanoBiT complementation and immunoblotting assays. The D2-, D3-, and D2 + D3–deleted FGFR2 fusions showed significantly impaired dimerization in the presence or absence of FGF10 ligand (Figure 1I). In keeping with the autoinhibitory function of the D1 domain (19), loss of the D1 domain enhanced receptor dimerization. Finally, we assessed the downstream pathway activation of the ECD deletion constructs by immunoblotting. Compared with the FL construct, expression of the D2, D3, and D2 + D3 deletion derivatives showed markedly impaired FGFR2 signaling (reduced p-FGFR (Y653/654), p-FRS2(Y436), and p-ERK(T202/Y204)), whereas the D1 deletion increased FGFR2 signaling output correlating with the observed increase in dimerization (Figure 1, I and J, and Supplemental Figure 1F). Together, these data demonstrate that the FGFR2-fusion ECD is necessary for full transformation of FGFR2 fusions. We further identify an autoinhibitory function of the D1 domain, deletion of which activates ERK leading to diminished viability, consistent with previous observations of activation-dependent lethality we and others observed in BRAF and NRAS mutant setting (20, 21).

Development of candidate biparatopic antibodies directed against FGFR2. To determine whether biparatopic antibodies can disrupt the function of FGFR2 fusions, we identified and produced 6 optimized FGFR2 antibodies (22–25), including the parental antibody of bemarituzumab, an ADCC-enhanced FGFR2 antibody in phase III trials (26). Available data suggested these antibodies likely bind to distinct epitopes in the ECD of FGFR2b, the primary isoform of FGFR2 fusions expressed in ICC (3). We compared and validated the reported binding epitopes and binding affinities, ascertaining FGFR2 binding by flow cytometry and bio-layer interferometry (BLI) octet analysis. We determined the apparent binding affinities of parental antibodies A–F, finding equilibrium dissociation constants (Kd) ranging from 0.15 nM–32.79 nM (Figure 2A). To validate their binding epitopes, NIH3T3 cells expressing FGFR2-fusion constructs with deletions in D1, D2, D3, or D2 + D3 (Figure 1E) were analyzed by flow cytometry The data showed that antibody A bound to all constructs, antibody B bound to all except the D1-deleted construct, antibodies C and D bound to all but the D2-deleted construct, and antibodies E and F bound to all except the D3-deleted construct (Figure 2B and Supplemental Figure 2A). These data defined the following binding epitopes: antibody B (D1), antibodies C and D (D2), antibodies E and F (D3), and antibody A (outside the D1–D3 domains, likely involving the N-terminus), consistent with prior reports (23). BLI-octet epitope binning analysis by pairwise cross competition corroborated our findings, showing antibodies A and B with unique binding epitopes while antibody C, D and antibody E, F pairs having overlapping epitopes (Figure 2, C and D, and Supplemental Figure 2B).

Development of candidate biparatopic antibodies directed against FGFR2. (A) Anti-FGFR2 antibodies (Ab-A, Ab-B, Ab-C, Ab-D, Ab-E, and Ab-F) binding to SNU16 cells (FGFR2 amplification) by flow cytometry and their associated apparent Kd values. Anti-hIgG1-FITC secondary antibody was used to detect FGFR2 parental antibodies A–F (n = 3). (B) Flow cytometry analysis using anti-hIgG1-FITC secondary antibody to detect FGFR2 parental antibodies A–F. Binding epitopes of parental antibodies A–F along the FGFR2 ECD were identified using full-length, D1, D2, D3, and D2+3–deleted FGFR2-BICC1 overexpressing NIH3T3 cell lines shown in Figure 1. (C) Epitope binning through cross competition assay. BLI-Octet Epitope clustering diagrams showing cluster dendrogram with au (approximately unbiased) P values and bp (bootstrap probability) value (%). Distance represents correlations and cluster method is average. (D) α-fold predicted structure of FGFR2 ECD showing D1, D2, D3, and D1-D2 flexible linker as well as 6 FGFR2 parental antibody binding epitopes A–F. (E and F) Viability of FGFR2-PHGDH–overexpressing BaF3 cells upon treatment with increasing concentrations of antibody A–F in the presence or absence of FGF10 ligand (n = 9). All data are mean ± SEM. Data are representative of 1 out of 2 independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 by 1-way ANOVA multiple comparisons.

To determine whether targeting FGFR2-fusion ECDs with anti-FGFR2 antibodies impaired their oncogenic activity, we treated BaF3 cells expressing FGFR2-PHGDH with each FGFR2 antibody. Antibodies against the ligand-binding domain (antibodies C, D, E, and F) inhibited FGF-stimulated growth (Figure 2E), supporting the notion that FGF ligands augment FGFR2-fusion activity and that the ECD is necessary for FGFR2 fusion–driven growth. In the ligand-independent setting, only antibody F inhibited FGFR2-PHGDH–driven BaF3 cell growth (Figure 2F). Antibodies B, D, and E had marginal impacts on cell growth in this setting, while antibodies A and C exhibited agonistic activity and promoted ligand-independent growth (Figure 2F). Consistent with its agonist activity, antibody C increased dimerization of FGFR2-AHCYL1 and FGFR2-BICC1 (Supplemental Figure 2C). As is the case with antibodies against the MET receptor that agonize and dimerize the receptors (14), the ligand-independent growth-promoting effects of antibodies A and C may result from unique binding epitopes eliciting antibody-induced dimerization. In addition, the differential activity of antibodies C and D suggests that they bind to distinct epitopes within the D2 domain.

We next asked whether FGFR2 biparatopic antibodies might have enhanced potency and might avoid ligand-independent agonism. We used controlled Fab-arm exchange to generate full IgG1 FGFR2 antibodies that simultaneously bind 2 different epitopes on the FGFR2 ECD (27). Here, complementary IgG Fc mutations force heterodimer formation between distinct IgG-formatted antibodies while maintaining heavy and light chain pairing. We produced each of the 6 parental antibodies with the reciprocal mutations to create 15 unique biparatopics from all pairwise combinations (Figure 3, A and B). In mass spectrometry analysis each biparatopic antibody showed greater than 95% purity with minimal residual parental antibody (as in Supplemental Figure 3, A and B). In all, we validated the binding affinities as well as binding epitopes of the 6 parental antibodies and generated 15 biparatopic antibodies for further characterization.

Identification of potent tumor growth–inhibiting biparatopic antibodies via unbiased screening. (A) Illustrations showing strategy for biparatopic antibody generation. (B) A diagram showing all 15 possible biparatopic antibody pairs that were generated from 6 parental antibodies A–F. (C and D) Viability of FGFR2-AHCYL1 overexpressing BaF3 cells upon treatment with IgG1, biparatopic antibodies, and their parental antibodies in the absence (C) and presence of FGF10 (D) (n = 2). Data are representative of 1 out of 2 independent experiments. (E) Binding affinities (Kd, nM) of parental antibodies (gray) compared with biparatopic antibodies (blue) from MSD-SET assay. Biparatopic antibodies bpAb-B/D and bpAb-B/C showed apparent binding affinities (apparent Kd) of 0.07 nM (orange bar) and 0.18 nM (pink bar), respectively (n = 2). Data are representative of 1 independent experiment. (F) Representative binding curves illustrating the binding avidity between FGFR2-PHGDH expressing NIH3T3 cells and antibody B, D, C or biparatopic antibody bpAb-B/C and bpAb-B/D via acoustic force spectroscopy (n = 4–6). Data are representative of 1 independent experiment.

Unbiased screening identifies potent, tumor growth–inhibiting biparatopic antibodies. We next assessed antiproliferative activity in FGFR2-fusion driven BaF3 cells with or without addition of ligand. Of the 15 biparatopic antibodies tested, 7 (46%) and 11 (73%) outperformed parental antibodies at inhibiting growth of FGFR2-AHCYL1–driven BaF3 cells in the absence or presence of FGF10 ligand, respectively (Figure 3, C and D). A second BaF3 model driven by an FGFR2-PHGDH fusion yielded similar results (Supplemental Figure 3, C and D). Notably, bpAb-B/C and bpAb-B/D were the most potent of the 21 parental and biparatopic antibodies in the viability assays. Importantly, the efficacy of pairwise mixtures of the parental antibodies differed from and did not predict the potency of their respective biparatopic antibodies (Supplemental Figure 3, E and F), suggesting that distinct modes of action are enabled by the biparatopic format.

We next determined the apparent binding affinity of the biparatopic antibodies for FGFR2. Using the MSD-SET assay, we found that 80% (12 out of 15) of biparatopic antibodies, including bpAb-B/C and bpAb-B/D, had marked improvements (greater than 10-fold) in FGFR2 apparent binding affinities compared with their parental antibodies (Figure 3E). The remaining 3 biparatopic antibodies with lower affinities had binding epitopes either within the same ECD subdomain (D2 for bpAb-C/D; D3 for bpAb-E/F) or on subdomains that are the farthest apart (D1 and D3 for bpAb-A/E). These data suggest that the geometry of binding between antibodies and their epitopes plays an important role in achieving high apparent affinity binding. We next determined the binding avidity to FGFR2-expressing cells using acoustic force spectrometry. After binding of antibody-coated beads to FGFR2-PHGDH–expressing NIH3T3 cells on the chip, acoustic force ramp from 0 to 1,000 pN was applied and antibody detachment from cells was observed using real-time fluorescence imaging. bpAb-B/C and bpAb-B/D had markedly enhanced binding avidity compared with parental antibodies B, C, and D, confirming the affinity data (Figure 3F). Finally, we examined the kinetics of antibody association and dissociation using BLI-octet analysis. In addition to their enhanced binding avidity, antibodies bpAb-B/C and bpAb-B/D also exhibited slower off rates and higher apparent affinity (low Kd) compared with their parental antibodies B, C, and D (Supplemental Figure 3, G and H). Both bpAb-B/C and bpAb-B/D contain binding arms against epitope B, a flexible autoinhibitory ECD D1 (Figure 2D). Together, our data demonstrate that the majority of biparatopic antibodies against combinations of selected epitopes on the FGFR2 ECD have enhanced antitumor activity and cellular binding avidity compared with their parental antibodies. Based on these attributes we selected bpAb-B/C and bpAb-B/D for further characterization.

Biparatopic antibodies show superior inhibition of growth and transformation of FGFR2 fusion driven cholangiocarcinoma cell lines. We investigated the impact of biparatopic FGFR2 antibody candidates bpAb-B/C and bpAb-B/D on 2 patient-derived models of FGFR2 fusion + ICC, ICC13-7 (FGFR inhibitor–sensitive), and ICC21 (partially sensitive) (28). ICC13-7 and ICC21 express the endogenous FGFR2-OPTN and FGFR2-CBX5 fusions, respectively. Correlating with their activity in FGFR2-fusion expressing BaF3 cells, bpAb-B/C and bpAb-B/D have enhanced efficacy at inhibiting growth of ICC13-7 and ICC21 cells in the absence (Figure 4, A and C) and, even greater, in the presence (Figure 4, B and C), of FGF10 compared with the parental antibodies.

Biparatopic antibodies show superior inhibition of growth and transformation of a FGFR2 fusion–driven cholangiocarcinoma cell line. (A–C) Viability of cholangiocarcinoma cell line ICC13-7 or ICC21 upon treatment with biparatopic antibodies bpAb-B/C, bpAb-B/D, parental antibodies B, D, C, or IgG1 isotype in the absence (A and C) or presence (B and C) of FGF10 at 14 days after seeding (n = 3). (D and E) Proteome profiler human phospho-kinase array demonstrating levels of 43 phosphorylated human kinases in NIH3T3 cells overexpressing FGFR2-PHGDH treated with IgG1, bpAb-B/C, or bpAb-B/D for 5 hours (D). (E) Quantification of levels of p-FGFR1, p-FGFR2, p-FGFR3, and p-FGFR4 (white boxes) (n = 2). (F) Viability of CCLP-1 cells upon treatment with biparatopic antibodies bpAb-B/C, bpAb-B/D, parental antibodies B, D, C, or IgG1 isotype control (n = 3). (G and H) Immunoblot of ICC13-7 cells upon 5 hours after treatments with bpAb-B/C, or bpAb-B/D compared to the parental antibodies B, D, C in the absence (G) or presence (H) of FGF10 ligand. (I and J) Representative images of focus formation assays of FGFR2-PHGDH–expressing NIH3T3 cells upon treatments with parental antibodies B, D, C, biparatopic antibodies bpAb-B/C and bpAb-B/D, or IgG1 (I) as quantified by the number of colonies (J) (n = 3). Scale bar: 1000 μm. All data are mean ± SEM. Data are representative of 1 out of 2 independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 by 1-way ANOVA multiple comparisons.

To investigate whether cell growth inhibition caused by bpAb-B/C and bpAb-B/D were specific to inhibition of FGFR2 rather than other FGFRs, extracts from NIH3T3 cells expressing FGFR2-PHGDH were profiled using a phospho-RTK array. We found that bpAb-B/C and bpAb-B/D specifically inhibited phosphorylation of FGFR2 but not of FGFR1 or FGFR3 (Figure 4, D and E; minimal FGFR4 phosphorylation was detected in these cells). We also tested FGFR2 specificity using the CCLP-1 ICC cell line, which lacks an FGFR2 fusion and is driven by FGFR1 and FGF20 overexpression (3). Both bpAb-B/C and bpAb-B/D treatments had no significant impact on CCLP-1 cell viability, whereas the IC50 for FGFR1-3 inhibitor futibatinib is less than 1.5 nM (3) (Figure 4F). Thus, bpAb-B/C and bpAb-B/D inhibit FGFR2 with high specificity.

We next examined the effects of bpAb-B/C and bpAb-B/D on FGFR2-fusion–mediated signaling. Both bpAb-B/C and bpAb-B/D robustly decreased p-FGFR, p-FRS2, and p-ERK compared with their parental antibodies B, C, or D in a ligand-independent setting (Figure 4G, and Supplemental Figure 4, A, B, and E); additionally, bpAb-B/C and bpAb-B/D blocked FGF10-induced phosphorylation of FGFR, FRS2, and ERK (Figure 4H, and Supplemental Figure 4, A, B, and F). Similarly, bpAb-B/C and bpAb-B/D impaired downstream signaling in NIH3T3 cells expressing FGFR2-PHGDH, including p-FGFR, p-FRS2, p-AKT, and p-ERK (Supplemental Figure 4, C and D). Thus, bpAb-B/C and bpAb-B/D specifically inhibit downstream signaling by constitutively active FGFR2-fusion proteins.

We next assessed the ability of bpAb-B/C and bpAb-B/D to inhibit FGFR2-fusion–driven oncogenic activity via focus formation assays using FGFR2-PHGDH–transformed NIH3T3 fibroblasts (Figure 4I). Cells treated with bpAb-B/C and bpAb-B/D showed a dose-dependent decrease in transformation capacity (reduction in colony formation), whereas the parental antibodies and IgG1-treated control had no effect (Figure 4J). Collectively, these results highlight the specificity of the biparatopic antibodies toward FGFR2 and the marked improvement in the potency of FGFR2 inhibition when compared with bivalent monotopic antibodies.

Biparatopic antibodies show superior in vivo antitumor activity compared with the parental antibodies. We next tested the in vivo efficacy of bpAb-B/C and bpAb-B/D and their parental antibodies against subcutaneous tumors formed by FGFR2-PHGDH–transformed BaF3 cells in SCID mice. At a tumor size of approximately 250mm3, mice were randomized into 10 groups with 10 mice per treatment group. The antibodies were administered via intravenous tail vein injections twice per week for 4–6 weeks. Both bpAb-B/C and bpAb-B/D biparatopic antibodies potently suppressed tumor growth at 5, 15, and 25 mg/kg doses, whereas the parental antibodies (administered at 15 mg/kg) showed no antitumor activity (Figure 5, A and B). Pharmacokinetics analysis by ELISA demonstrated dose-proportional increases in the plasma concentration of the biparatopic antibodies, and, furthermore, considerably longer half life compared with small molecule inhibitors, consistent with their larger size (29, 30) (Supplemental Figure 5, A and B).

Biparatopic antibodies show superior in vivo antitumor activity compared with the parental antibodies. (A–D) Tumors of BALB/c scid mice (n = 10 per group) harboring BaF3 cells overexpressing FGFR2-PHGDH (A and B) or ICC13-7 (C and D) subcutaneous xenografts treated with parental and biparatopic antibodies. Results are represented in the waterfall plot illustrating changes in tumor volume at day 25 (A and B) or day 38 (C and D) after initial treatment (A and C) and as geometric mean of tumor volumes ± SEM every 3–4 days from days 0–25 after initial treatment (B and D). Data are mean ± SEM across 10 mice. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 by Friedman’s ANOVA multiple comparisons. (E) Immunoblot analysis of FGFR2-PHGDH–overexpressing BaF3 cells xenograft tumors harvested 5 hours after the final round of bpAb-B/C, bpAb-B/D, or IgG1 administration at 25 days after initial treatment. (F) Immunoblot analysis of ICC13-7 xenograft tumors collected 5 hours after the final round of antibody administration on day 38 after initial treatment. (G) Representative images of H&E and IHC staining for proliferation marker Ki-67 in ICC13-7 xenograft tumor samples on the final day of treatment. Scale bars, 100 μm. (H) Quantification of the percent of Ki-67–positive nuclei normalized to the total number of nuclei (nuclei counterstain). Data are from 2 biological replicates per treatment group with at least 14 representative images for analysis per group. Data are presented in a superplot where each color represents data points from the same biological sample. Black dots indicate the average values for each biological sample, while black lines represent the overall average for all data points. All data are mean ± SEM. One independent experiment was performed.

The biparatopic antibodies also showed prominent in vivo efficacy against xenograft tumors formed by the patient-derived, ICC13-7 cholangiocarcinoma model. While the parental antibodies had only marginal effects on tumor growth, the biparatopics were highly effective at both 10 and 30 mg/kg dose concentrations. Notably, bpAb-B/C showed greatest potency, resulting in tumor stasis at 38 days after treatment (Figure 5, C and D), comparable with the efficacies of clinically used FGFR inhibitors (28, 31). Importantly, bpAb-B/C and bpAb-B/D treatment in both in vivo models led to a marked decrease in total FGFR2 levels and reductions in p-FGFR, p-FRS2, and p-ERK compared with IgG1 control (Figure 5, E and F, and Supplemental Figure 5, C and D). By contrast, the parental antibodies showed limited effect on total FGFR2 levels or on downstream signaling (Supplemental Figure 5, E and F). Consistent with the tumor growth inhibition data, bpAb-B/C and bpAb-B/D markedly decreased tumor cell proliferation (Ki-67 staining) compared with parental antibodies or IgG1 control (Figure 5, G and H). None of the antibody treatments affected mouse body weight (Supplemental Figure 5, G and H). Assessment of antibody tumor distribution by IHC staining showed that bpAb-B/C and bpAb-B/D localized to the cell membrane and exhibited diffuse staining throughout ICC13-7 xenografts (Supplemental Figure 5I), suggesting that biparatopic antibodies penetrate tumor effectively.

To investigate the potential involvement of immune effector functions mediated by biparatopic antibodies in ICC13-7 xenografts, we performed IHC staining for mouse NKp46, a marker for NK cell–mediated antibody dependent cell-mediated cytotoxicity (ADCC) activation (32) and found no significant changes (Supplemental Figure 5, J and K). Similarly, RNA-seq analysis revealed minimal changes in murine gene expression across treatments except for the bpAb-B/C at 10 mg/kg treatment group with only 4 immune-related genes upregulated (Supplemental Figure 5L). We further analyzed the immune system–related gene sets and found no significantly differentially expressed genes observed among treatment groups (Supplemental Figure 5, N–Q). In all cases, tumor growths of matching bpAb-B/C– and bpAb-B/D–treated xenografts were substantially inhibited (Supplemental Figure 5M). Additionally, these antibodies were not potent inducers of NK cell killing of cancer cells (Supplemental Figure 5R), nor robust inducers of NFAT reporters via CD16 (ADC) or CD32a (antibody dependent cellular phagocytosis) in engineered Jurkat cells (Supplemental Figure 5, S and T). Together, these results demonstrate that bpAb-B/C and bpAb-B/D have improved antitumor activity compared with their parental antibodies in vivo, likely driven by receptor downregulation.

Biparatopic antibodies promote receptor internalization and lysosomal degradation. We next explored the potential mechanism for FGFR2 downregulation by the biparatopic antibodies. To determine whether bpAb-B/C and bpAb-B/D promote FGFR2-fusion internalization, we treated FGFR2-PHGDH–expressing BaF3 with bpAb-B/C, bpAb-B/D, or IgG control and then transferred cells to 4°C to block or 37°C to induce internalization. Surface levels of FGFR2 were analyzed by flow cytometry (Figure 6, A and B). Cells treated with bpAb-B/C and bpAb-B/D showed increased internalization from 60 to 960 minutes (from approximately 6% to 80% shift in surface FGFR2) (Figure 6B). The internalization assay was repeated in ICC13-7 cells treated with bpAb-B/C, bpAb-B/D, respective parental antibodies, or IgG control. ICC13-7 cells treated with bpAb-B/C and bpAb-B/D had a significant decrease in surface FGFR2 compared with cells treated with parental antibodies B, C, or D, or IgG1, suggesting that bpAb-B/C and bpAb-B/D enhanced FGFR2 receptor internalization (Figure 6C). Next, we labeled biparatopic and parental antibodies with a Fab fragment conjugated to a pH-sensitive fluorophore (33) and assessed lysosome-mediated induction of fluorescence in FGFR2-PHDGH, FGFR2-AHCYL1, and FGFR2-BICC1–expressing NIH3T3 cells (Figure 6D). Treatment with bpAb-B/C and bpAb-B/D resulted in marked increases in the fluorescent signal compared with the parental antibodies (Figure 6, E–H). Labelling of lysosomes with lysotracker (green) and biparatopic antibodies with Fab-Fluor (red) demonstrated colocalization of the 2 signals, confirming the presence of the antibodies in the lysosomes (Supplemental Figure 6A). Consistent with results in FGFR2 fusion–expressing NIH3T3 cells, treatment of the ICC13-7 cholangiocarcinoma cell line with bpAb-B/C and bpAb-B/D led to increases in fluorescent signals compared with parental antibodies (Figure 6I). In addition, bpAb-B/C and bpAb-B/D showed enhanced receptor internalization and degradation compared with parental antibodies as well as parental antibody mixtures, confirming the unique mechanism of action of biparatopic antibodies beyond antibody combinations (Supplemental Figure 6C).

The biparatopic antibodies promote receptor internalization and lysosomal degradation. (A) Flow cytometry histograms of surface FGFR2-PHGDH in BaF3 cells at 4°C (blue) and 37°C (red) upon treatment with bpAb-B/C or bpAb-B/D from 60–960 minutes. (B) Quantification of the histograms demonstrating the percentage of internalized FGFR2 at 60, 120, 180, 240, and 960 minutes after bpAb-B/C or bpAb-B/D incubation. (C) Quantification of histograms showing percent internalized FGFR2 in ICC13-7 cell line at 4°C and 37°C after 5 hours of treatment with parental antibody B, D, C or biparatopic antibodies bpAb-B/C or bpAb-B/D (n = 3). Data are mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 by 1-way ANOVA multiple comparisons. Data are representative of 1 out of 2 independent experiments. (D) Illustrations of Fabfluor-pH antibody labeling assay. The pH sensitive dye-based system exploits the acidic environment of the lysosomes to quantify internalization of the labeled antibody. Fluorescent signals that indicate the internalization/degradation events were tracked using Incucyte. (E) Representative images of detected fluorophore in NIH3T3 cells expressing FGFR2-PHGDH treated with parental antibody B, D, C, or biparatopic antibody bpAb-B/C and bpAb-B/D at 15 hours after incubation. Scale bars: 300 μm. (F–H) Quantification of internalization/degradation signals in FGFR2-AHCYL1 (F), FGFR2-BICC1 (G), and FGFR2-PHGDH (H) expressing NIH3T3 cells treated with parental antibodies B, D, C, or biparatopic antibody bpAb-B/C and bpAb-B/D from 24 hours after incubation. Data are representative of 1 out of 2 independent experiments. (I) Quantification of internalization/degradation signals in ICC13-7 cells treated with parental antibodies B, D, C, or biparatopic antibody bpAb-B/C and bpAb-B/D at 4 hours after incubation. Data are representative of 1 out of 2 independent experiments. (J) Immunoblot of ICC13-7 cells treated with IgG1, bpAb-B/C,or bpAb-B/D antibodies alone or cotreated with bafilomycin A1 (BafA1) for 24 hours. BafA1 was preincubated for 1 hour prior to antibody treatments. Data are representative of 1 independent experiment.

To investigate whether the observed increase in FGFR2 internalization is triggered by the intermolecular binding of antibodies creating a large complex, as shown in previous work (17, 34), we performed size exclusion chromatography coupled with multiangle light scattering (SEC-MALS), to determine the mass of the antibody and its complexes. Upon increasing the ratio of antigen (FGFR2 ECD) to the biparatopic antibody bpAb-B/C (ECD:Ab) from 1:1, 3:1, and 5:1, SEC-MALS data showed absolute masses consistent with higher-order complexes (Supplemental Figure 6B, see predicted complexes). These results suggest that the bpAb-B/C biparatopic antibodies bind to FGFR2 receptors in trans, likely creating larger antibody-receptor complexes and leading to more rapid internalization.

To determine whether the internalization and receptor downregulation are mediated by lysosomal degradation, we suppressed lysosome acidification and catabolism using the vacuolar-type H+–ATPase inhibitor bafilomycin A1 (BafA1). BafA1 treatment rescued bpAb-B/C- or bpAb-B/D-induced FGFR2-OPTN downregulation in ICC13-7 compared with IgG1-treated control (Figure 6J and Supplemental Figure 6D). Together, these data demonstrate that bpAb-B/C- and bpAb-B/D-induce FGFR2-fusion internalization, trafficking, and lysosomal-mediated degradation to decrease FGFR2 fusion–driven activity and growth. Notably, this mode of action induced by the biparatopic antibodies as shown in our work and others (17, 35–37), does not require cotargeting of lysosome-targeting receptors, membrane E3 ligases, or autophagy signaling molecules, as seen in the development of LYTAC, AbTAC, or AUTAC systems (38).

Biparatopic antibodies potentiate the efficacy of FGFR inhibitors. Given the specificity of FGFR2 antibodies and the potency of FGFR1-3 kinase inhibitors, combining 2 distinct treatment modalities might result in cooperativity specific to FGFR2 while sparing FGFR1 and 3, leading to more potent FGFR2 inhibition. To test whether bpAb-B/C and bpAb-B/D synergize with FGFRi, FGFR2-PHGDH–expressing BaF3 cells were treated in a titration matrix of bpAb-B/C or bpAb-B/D in combinations with approved FGFRi infigratinib, futibatinib, and pemigatinib. The Bliss model was then applied to determine the degree of synergy (39). Bliss scores of 0–10 generally indicate additive interactions, while scores greater than 10 demonstrate synergistic interactions. In the absence of FGF10, combination of bpAb-B/D with infigratinib, pemigatinib, or futibatinib as well as combination of bpAb-B/C with futibatinib or pemigatinib moderately enhanced growth inhibition (Figure 7, A and B). Synergy between bpAb-B/C and infigratinib in a ligand-independent setting was striking, with a Bliss score of greater than 20 (Figure 7, B and C). In the presence of FGF10, cotreatments of bpAb-B/C or bpAb-B/D with infigratinib, futibatinib, and pemigatinib all enhanced growth suppression compared with treatment with single agents (Figure 7, A–C). In accordance with the dose-response, all Bliss values were well above 10 in the ligand-dependent context (Figure 7C). These data highlight the potential of the biparatopic antibodies to boost the activity of FGFR inhibitors both in the presence and absence of ligand.

Combinations of biparatopic antibodies with FGFR inhibitors. (A and B) Biparatopic antibody B/D (A) or B/C (B) with Infigratinib, Futibatinib, or Pemigatinib combination dose response matrices in the presence of absence of FGF10. 1 = 100% viability and 0= 0% viability after indicated treatment. (C) Heatmap showing Bliss scores calculated from dose response matrices using SynergyFinder (39) application for drug combination analysis. (D and E) Viability of NIH3T3 cells stably expressed FGFR2-AHCYL1 with V565I or V565F mutations treated with bpAb-B/D, bpAb-B/C, or IgG1 (n = 3). (F) Immunoblot analysis of NIH3T3 cells stably expressing FGFR2-AHCYL1 with V565I or V565F treatment with bpAb-B/D, bpAb-B/C, or IgG1 for 5 hours (n = 3). (G and H) Quantification of internalization/degradation signals in FGFR2-AHCYL1 with V565I or V565F–expressing NIH3T3 cells treated with biparatopic antibody bpAb-B/C, bpAb-B/D, or IgG1 from 0–38 hours after incubation. (I) Viability of CCLP-1 cells stably expressed FGFR2–PHGDH fusion with V565F mutation upon treatment with IgG1, bpAb-B/D, or bpAb-B/C alone or in combination with Infigratinib (percentage compared with IgG1 treated control) (n = 3). (J) Immunoblot analysis of CCLP-1 cell line expressing FGFR2-PHGDH with V565F mutation upon treatment with IgG1, bpAb-B/C, bpAb-B/D, IgG1+Infigratinib, bpAb-B/C + Infigratinib, or bpAb-B/D + Infigratinib for 5 hours. (K) Deletion mutations derived from 4 different patients and the respective FGFR2 ECD. (L) Viability of 4 patient-derived N-terminus oncogenic mutants upon treatments with IgG1, bpAb-B/C, or bpAb-B/D as indicated (percentage viability compared with IgG1) (n = 3). (M) Immunoblot of NIH-3T3 cells bearing an FGFR2 H167_N173 in-frame deletion allele (patient 2) after treatment with IgG, bpAb-B/C, bpAb-B/D, or the relevant parental antibodies for 5 hours. All data are mean ± SEM. Data are representative of 2 independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 by 1-way ANOVA with multiple comparisons.

Diverse secondary FGFR2 kinase domain mutations drive clinical resistance to each of each FGFR TKI studied to date (3, 40, 41). Given the intracellular location of the kinase domain, we hypothesized that the biparatopic antibodies might remain active against these mutations. To test this hypothesis, we selected the gatekeeper mutations V565I and V565F, which are common mechanisms of resistance to the approved FGFR inhibitors. NIH3T3 cells that stably expressed FGFR2-AHCYL1 with a V565I or V565F mutation were resistant to infigratinib (Supplemental Figure 7A) but were sensitive to bpAb-B/C and bpAb-B/D, showing inhibition of both growth (Figure 7, D and E) and downstream signaling, as evidenced by levels of p-FGFR, p-FRS2, and p-ERK1/2 (Figure 7F and Supplemental Figure 7B). Moreover, bpAb-B/C or bpAb-B/D–induced lysosomal degradation of the FGFR2 fusion in these cells as assayed by anti-Fc Fab fragment conjugated pH-sensitive fluorophore (Figure 7, G and H), similar to that observed in NIH3T3 cells expressing the initial FGFR2 fusions (Figure 6, F–H). Given the complexity of resistance mechanisms in patient tumors, which may implicate multiple oncogenes and bypass mechanisms, we modeled the efficacy of our antibodies in the FGFR1-dependent cholangiocarcinoma cell line, CCLP-1, stably transduced to express the FGFR2-PHGDH-WT or FGFR2-PHGDH-V565F alleles (Supplemental Figure 7, C and D). CCLP-1 parental cells as well as CCLP-1 cells expressing FGFR2-PHGDH WT were sensitive (IC50 < 2 nM), while FGFR2-PHGDH V565F cells were resistant (IC50 > 2,000 nM) to infigratinib (Supplemental Figure 7E). To determine the dose of infigratinib to use in combination studies (in order to suppress the concurrent FGFR1 activity), we determined the infigratinib concentration that sensitized cells expressing FGFR2-PHGDH-WT but not FGFR2-PHGDH-V565F (0.15 μM). Treatment with bpAb-B/C or bpAb-B/D in combination with infigratinib significantly suppressed growth of V565F resistant mutants and resensitized the CCLP-1 resistant cells to infigratinib, indicating robust suppression of the introduced FGFR2 resistance allele (Figure 7I). In addition, cotreatments of infigratinib and bpAb-B/C or bpAb-B/D decreased levels of FGFR2, p-FGFR, p-FRS2, and p-ERK1/2 (Figure 7J and Supplemental Figure 7F). These results support the use of bpAb-B/C and bpAb-B/D to overcome secondary FGFR2 kinase domain mutations.

In addition to FGFR2 rearrangements, a recent study revealed that activating in-frame FGFR2 ECD deletions occur in approximately 3% of patients with ICC. Patients with these FGFR2 ECD deletions responded well to FGFRi treatments, suggesting that these ECD mutations are oncogenic drivers (42). Since these mutations are located in the ECD, it is possible that they might lack sensitivity to our biparatopic antibodies. To determine whether bpAb-B/C or bpAb-B/D have activity against oncogenic FGFR2 ECD in-frame–deletion mutations, we engineered NIH3T3 cells to stably express 4 patient-derived FGFR2 ECD-deletion mutations (Figure 7K). Compared with NIH3T3 cells expressing FGFR2-WT, cells expressing deletion mutations had increased transformation capacities and receptor dimerization as analyzed by soft-agar assay and NanoBiT assays, respectively (Supplemental Figure 7, G–K). In addition, the ECD mutants had elevated FGFR2 downstream phosphorylation; p-FGFR, p-FRS2, and p-ERK1/2, which was blocked by infigratinib, confirming their FGFR2 dependency (Supplemental Figure 7, L and M). While bpAb-B/C or bpAb-B/D had moderate activities against patient 1– and 3–derived mutants, both bpAb-B/C and bpAb-B/D effectively inhibited growth of patient-2 and -4 variants (Figure 7L). These results correlated with the decrease in levels of FGFR2, p-FGFR, p-FRS2, and p-ERK1/2 for the H167_N173Del (patient 2) variant (Figure 7M and Supplemental Figure 7O). Importantly, levels of FGFR2 decreased upon bpAb-B/C and bpAb-B/D treatments, suggesting that receptor internalization and degradation mediate the observed growth inhibition (Figure 7M and Supplemental Figure 7O). Crucially, mutations found in patients 1–4 are predicted to alter the 3-dimensional structure of FGFR2 D2 and D3 domains (42) and may consequently affect the binding affinities of bpAb-B/C and bpAb-B/D with D1 and D2 binding arms. Nevertheless, the fact that bpAb-B/C and bpAb-B/D remain effective against patient 2 and 4 variants suggest that as long as the binding avidities of D1 and D2 binders are sufficient to establish intermolecular interaction and trigger internalization, the bpAb-B/C and bpAb-B/D should be effective. These data demonstrate that bpAb-B/C and bpAb-B/D have activities against intracellular kinase domain mutations and specific patient-derived FGFR2 ECD oncogenic deletions. Together with the observed synergy, these data support the notion of combining FGFR1-3 inhibitors with FGFR2 biparatopic antibodies.