Potent and selective antitumor activity of a T cell-engaging bispecific antibody targeting a membrane-proximal epitope of ROR1

Significance

Harnessing and enhancing the innate and adaptive immune system to fight cancer represents one of the most promising strategies in contemporary cancer therapy. Although bispecific antibodies (biAbs) that combine a T cell-engaging arm with a tumor cell-binding arm are particularly potent cancer immunotherapeutic agents, they rely on the identification of tumor antigens with highly restricted expression. The receptor tyrosine kinase ROR1 is expressed by numerous cancers and is largely absent from postnatal healthy cells and tissues. Here we show that T cell-engaging biAbs that target ROR1 are highly potent in in vitro, in vivo, and ex vivo models of cancer, in particular when targeting a conserved site on ROR1 close to the tumor cell membrane we precisely mapped by X-ray crystallography.

Abstract

T cell-engaging bispecific antibodies (biAbs) present a promising strategy for cancer immunotherapy, and numerous bispecific formats have been developed for retargeting cytolytic T cells toward tumor cells. To explore the therapeutic utility of T cell-engaging biAbs targeting the receptor tyrosine kinase ROR1, which is expressed by tumor cells of various hematologic and solid malignancies, we used a bispecific ROR1 × CD3 scFv-Fc format based on a heterodimeric and aglycosylated Fc domain designed for extended circulatory t_1/2 and diminished systemic T cell activation. A diverse panel of ROR1-targeting scFv derived from immune and naïve rabbit antibody repertoires was compared in this bispecific format for target-dependent T cell recruitment and activation. An ROR1-targeting scFv with a membrane-proximal epitope, R11, revealed potent and selective antitumor activity in vitro, in vivo, and ex vivo and emerged as a prime candidate for further preclinical and clinical studies. To elucidate the precise location and engagement of this membrane-proximal epitope, which is conserved between human and mouse ROR1, the 3D structure of scFv R11 in complex with the kringle domain of ROR1 was determined by X-ray crystallography at 1.6-Å resolution.

ROR1 is a receptor tyrosine kinase uniformly expressed on the cell surface of malignant B cells in chronic lymphocytic leukemia (CLL) (1–4) and mantle cell lymphoma (MCL) (5–7) but not on healthy B cells or, with some exceptions (8), other healthy cells and tissues of patients with cancer. In addition to leukemia and lymphoma, ROR1 is also expressed in subsets of carcinoma, sarcoma, and melanoma, i.e., in all major cancer categories (9). Thus, ROR1 is a highly attractive candidate for targeted cancer therapy. Ongoing phase I clinical trials are assessing the safety and activity of an mAb (ClinicalTrials.gov identifier NCT02222688) and chimeric antigen receptor (CAR)-engineered T cells (CAR-Ts; ClinicalTrials.gov identifier NCT02706392) targeting ROR1 in hematologic and solid malignancies.

By using phage display, we previously generated a panel of rabbit anti-human ROR1 mAbs from immune and naïve rabbit antibody repertoires (10, 11). These mAbs were shown to bind different epitopes on ROR1 with high specificity and affinity. As CAR-Ts, they mediated selective and potent cytotoxicity toward ROR1-expressing malignant cells (11, 12). One of these CAR-Ts based on rabbit anti-human ROR1 mAb R12 demonstrated safety and activity in nonhuman primates (13) and was translated to the ongoing phase I clinical trial.

With a mechanism of action (MOA) that is conceptually related to the MOA of CAR-Ts (14, 15), T cell-engaging bispecific antibodies (biAbs) are an alternative strategy for cancer immunotherapy. Like CAR-Ts, T cell-engaging biAbs utilize the power of T cells for tumor cell eradication, but, unlike CAR-Ts, are manufactured and administered similarly to conventional mAbs and are cleared from the patient with cancer. The concept of retargeting cytolytic T cells toward tumor cells by T cell-engaging biAbs has had substantial clinical success with a CD19 × CD3 biAb in bispecific T cell-engager (BiTE) format, blinatumomab, which received US Food and Drug Administration (FDA) approval for the treatment of refractory or relapsed B cell-precursor adult lymphoblastic leukemia in 2014 (16), and with numerous other formats to various targets and indications under investigation in clinical trials (17, 18). In view of their similar MOAs, CAR-Ts and T cell-engaging biAbs can be accompanied by potentially dangerous adverse events, particularly cytokine release syndrome and central nervous system toxicity (19). Although both on-target and off-target effects contribute to adverse events, targeting cell surface antigens that are restricted to tumor cells is generally thought to afford superior safety profiles compared with targeting more widely expressed cell surface antigens.

In addition to the restricted expression of ROR1 on tumor cells, its relatively large extracellular domain (ECD) consisting of an Ig, a frizzled (Fz), and a kringle (Kr) domain, together comprising ∼375 extracellular amino acids, provides an opportunity to compare the therapeutic utility of T cell-engaging biAbs that bind to membrane-distal and membrane-proximal epitopes of ROR1. Efficient formation of cytolytic synapses (20) between T cells and tumor cells is thought to entail an optimal spacing between T cell and tumor-cell membranes. This, in turn, depends on the spacing between the ROR1- and the CD3-engaging arm of the biAb as well as on the location of its epitopes on ROR1 and CD3. With a panel of rabbit anti-human ROR1 mAbs and an established humanized mouse anti-human CD3 mAb at hand, we sought to identify the best candidate for a heterodimeric and aglycosylated scFv-Fc format designed for extended circulatory t_1/2 and diminished systemic T cell activation. Rabbit mAb R11, which binds a conserved membrane proximal epitope on human and mouse ROR1 with similar affinity (10), emerged as the most potent ROR1-engaging arm. Its precise location and engagement was determined by cocrystallization of scFv R11 with the Kr domain of human ROR1. Collectively, our study encourages and enables the development of R11-based or R11 epitope-based T cell-engaging biAbs and other T cell-engaging cancer therapeutic agents.

Results

Design, Generation, and Validation of ROR1 × CD3 biAbs.

By using the established knobs-into-holes technology (21, 22), we designed IgG1-mimicking ROR1 × CD3 biAbs in scFv-Fc format. These comprised (i) the variable light chain (V_L) and variable heavy chain (V_H) domains of a rabbit anti-human ROR1 mAb linked with a (Gly₄Ser)₃ polypeptide linker; (ii) (Gly₄Ser)₃-linked V_H and V_L domains of humanized mouse anti-human CD3 mAb v9 (23), which was derived from mouse anti-human CD3 mAb UCHT1 (24); and (iii) a human IgG1 Fc module with knobs-into-holes mutations for heterodimerization and with an Asn297Ala mutation to render the biAb aglycosylated (25) (Fig. 1A). To confirm preferential heterodimerization, we combined the scFv-Fc knobs cassette-encoding vector with a vector encoding the complementary Fc holes cassette without scFv module. Analysis by SDS/PAGE revealed that >90% of Protein A-purified antibody assembled as heterodimer. Based on seven rabbit anti-human ROR1 mAbs with diverse epitope specificity and affinity (10, 11) (SI Appendix, Table S1), a series of ROR1 × CD3 biAbs was constructed that included R12 × v9, R11 × v9, XBR1-402 × v9, ERR1-403 × v9, ERR1-TOP43 × v9, ERR1-306 × v9, and ERR1-324 × v9. The two polypeptide chains of the heterodimeric scFv-Fc were encoded on two separate mammalian cell-expression vectors based on pCEP4 and combined through transient cotransfection into HEK 293 Phoenix cells. Nonreducing and reducing SDS/PAGE revealed the expected ∼100-kDa and ∼50-kDa bands, respectively, after purification by Protein A affinity chromatography with a yield of ∼10 mg/L (Fig. 1B and SI Appendix, Fig. S1A). Further analysis by size-exclusion chromatography (SEC) revealed that the seven ROR1 × CD3 biAbs eluted mainly as a single peak with <10% aggregates (Fig. 1C and SI Appendix, Fig. S1B). Sequential Protein A affinity chromatography and SEC was used to purify monomeric biAbs for the following functional studies.

Fig. 1.

Design and generation of bispecific ROR1 × CD3 scFv-Fc. (A) Schematic diagram of ROR1 × CD3 biAb in scFv-Fc format combining a rabbit anti-human ROR1 scFv with the humanized mouse anti-human CD3 scFv v9 via a mutated Fc domain of human IgG1. The Fc domain contains one mutation for aglycosylation in the C_H2 domain (N297A) and two or four mutations in the C_H3 knob (S354C and T366W) and C_H3 hole (Y349C, T366S, L368A, and Y407V) domains, respectively, for heterodimerization. (B) SDS/PAGE and Coomassie blue staining analysis of purified representative R12 × v9 scFv-Fc showing the expected bands at ∼100 kDa under nonreducing (nr) and ∼50 kDa under reducing (r) conditions. (C) SEC analysis of R12 × v9 scFv-Fc eluting as major peak at 12.37 mL. The 10.38-mL minor higher molecular weight peak indicates the level of aggregation.

To study the dual binding specificity of ROR1 × CD3 biAbs, a flow cytometry assay was performed by using CD3-expressing human T cell line Jurkat, a stable K562 cell line ectopically expressing ROR1 (K562/ROR1), and MCL cell line JeKo-1, which expresses ROR1 endogenously. As shown in Fig. 2, and consistent with the pairing of the same CD3-engaging arm with seven different ROR1-engaging arms, all ROR1 × CD3 biAbs revealed similar binding to Jurkat cells but different binding to K562/ROR1 and JeKo-1 cells by flow cytometry. No binding to ROR1-negative parental K562 cells was detected for any of the biAbs (Fig. 2). To further confirm the dual binding specificity of the biAbs, we also cloned, expressed, and purified the corresponding eight homodimeric anti-ROR1 and anti-CD3 scFv-Fc. Flow cytometry analysis of R12, R11, XBR1-402, ERR1-403, ERR1-TOP43, ERR1-306, and ERR1-324 scFv-Fc showed binding to JeKo-1 but not Jurkat cells (SI Appendix, Fig. S2). By contrast, the v9 scFv-Fc bound only to Jurkat cells (SI Appendix, Fig. S2). These data suggest that the ROR1 × CD3 biAbs retained the integrity and specificity of the parental mAbs.

Fig. 2.

Cell surface binding of bispecific ROR1 × CD3 scFv-Fc. The indicated bispecific ROR1 × CD3 biAbs were analyzed for binding to human CD3-positive T cell line Jurkat and human ROR1-positive cell lines K562/ROR1 and JeKo-1 by flow cytometry at a concentration of 5 μg/mL followed by APC-conjugated goat anti-human IgG pAbs. Parental K562 is a ROR1-negative control cell line. Open histograms show the background binding signal of the secondary pAbs.

In Vitro T Cell Recruitment and Activation Mediated by ROR1 × CD3 biAbs.

We next examined the functionality of the seven ROR1 × CD3 biAbs in terms of target-dependent T cell recruitment and activation in vitro. Primary human T cells were isolated and expanded from healthy donor peripheral blood mononuclear cells (PBMCs) by using anti-CD3/anti-CD28 beads. The ex vivo-expanded human T cells served as effector cells and JeKo-1, K562/ROR1, and parental K562 cells as target cells. To study the ability of ROR1 × CD3 biAbs to recruit effector cells to target cells, we examined their cross-linking. T cells were stained red and mixed with green-stained K562/ROR1 or K562 cells in the presence of 1 μg/mL ROR1 × CD3 biAbs for 1 h, gently washed, and fixed with paraformaldehyde. Double-stained events detected and quantified by flow cytometry indicated cross-linked cell aggregates. As shown in Fig. 3 A and B, all seven ROR1 × CD3 biAbs exhibited highly efficient cross-linking (∼50%) between T cells and K562/ROR1 cells. A negative control scFv-Fc in which the ROR1-engaging arm was replaced with catalytic mAb h38C2 (26), which does not have a natural antigen, revealed only background cross-linking (<10%). Likewise, only background cross-linking was observed when using ROR1-negative parental K562 cells (SI Appendix, Fig. S3).

Fig. 3.

T cell engagement mediated by bispecific ROR1 × CD3 scFv-Fc. (A) Cross-linking of 5 × 10⁴ primary human T cells (stained with CellTrace Far Red) and 5 × 10⁴ K562/ROR1 cells (stained with CellTrace CFSE) in the presence of 1 µg/mL ROR1 × CD3 and control scFv-Fc. Double-stained events were detected by flow cytometry. (B) Quantification of double-stained events from three independent triplicates (mean ± SD). (C) The cytotoxicity of ROR1 × CD3 scFv-Fc was tested with ex vivo expanded primary human T cells and K562/ROR1 (C), JeKo-1 (D), or K562 (E) cells at an effector-to-target ratio of 10:1 and measured after 16 h. Shown are mean ± SD values from three independent triplicates.

To analyze the capability of the ROR1 × CD3 biAbs in mediating cytotoxicity in vitro, specific lysis of target cells after 16-h incubation with concentrations ranging from 2 pg/mL to 15 μg/mL at an effector-to-target ratio of 10:1 was assessed. K562/ROR1 and JeKo-1 cells but not K562 cells were killed in the presence of all seven ROR1 × CD3 biAbs, indicating that target cell lysis was strictly dependent on ROR1 (Fig. 3 C–E). All biAbs had higher activity against K562/ROR1 cells, which express substantially higher levels of ROR1 compared with JeKo-1 cells (Fig. 2). Negative control biAb, h38C2 × v9, was inactive. One ROR1 × CD3 biAb, R11 × v9, was significantly and consistently more potent than the other ROR1 × CD3 biAbs and killed K562/ROR1 and JeKo-1 cells with EC₅₀ values of 22 ng/mL (0.2 nM) and 209 ng/mL (2 nM), respectively (Fig. 3 C and D).

We next studied the ROR1 × CD3 biAbs for inducing T cell activation. The biAbs induced T cell activation only in the presence of K562/ROR1 cells but not K562 cells (Fig. 4). As analyzed by flow cytometry, the early T cell activation marker CD69 was up-regulated after 16 h incubation with 1 μg/mL ROR1 × CD3 biAbs compared with negative control biAb, h38C2 × v9 (Fig. 4A). As analyzed by ELISA, the release of type 1 cytokines IFN-γ, TNF-α, and IL-2 was also strictly dependent on the presence of ROR1-expressing target cells and ROR1 × CD3 biAbs (Fig. 4 B–D). Whereas all seven ROR1 × CD3 biAbs induced high IFN-γ secretion, R11 × v9 induced significantly higher levels of TNF-α and IL-2 secretion.

Fig. 4.

T cell activation mediated by bispecific ROR1 × CD3 scFv-Fc. Ex vivo expanded primary human T cells were incubated with 1 μg/mL of the indicated biAbs in the presence of K562/ROR1 or K562 cells at an effector-to-target ratio of 10:1 for 16 h. (A) Percentage of activated T cells based on CD69 expression. Cytokine release measured by ELISA for IFN-γ (B), TNF-α (C), and IL-2 (D). Shown are mean ± SD values for independent triplicates. One-way ANOVA was used to analyze significant differences between ROR1 × CD3 (colored) and control scFv-Fc (black; *P < 0.05, **P < 0.01, and ***P < 0.001).

In Vivo Activity of ROR1 × CD3 biAbs.

To investigate whether the in vitro T cell recruitment and activation mediated by ROR1 × CD3 biAbs would translate into in vivo activity, we used a xenograft mouse model of systemic human MCL. For this, 0.5 × 10⁶ JeKo-1 cells stably transfected with firefly luciferase (JeKo-1/ffluc) (12) were injected i.v. (tail vein) into five cohorts of eight female NOD-scid-IL2Rγ^null (NSG) mice per cohort, and, after 1 wk, when the tumor was disseminated, mice were injected i.v. (tail vein) with 5 × 10⁶ ex vivo expanded human T cells. One hour later, 10 μg of the ROR1 × CD3 biAbs R11 × v9 (cohort 2) and R12 × v9 (cohort 3), 10 μg of a positive control CD19 × CD3 biAb (cohort 4), and 10 μg of a negative control HER2 × CD3 biAb (cohort 5) were administered i.v. (tail vein). The CD19 × CD3 biAb shared the same heterodimeric and aglycosylated scFv-Fc format and the same CD3-binding arm with the ROR1 × CD3 biAbs. Its CD19-binding arm was derived from human anti-human CD19 mAb 21D4 (Medarex; US Patent 8,097,703). Cohort 1 received only vehicle (PBS solution). All five cohorts were treated with a total of three doses of T cells (every 8 d) and six doses of biAbs (every 4 d). Mice were preconditioned with 250 μL human serum 24 h before every dose. To assess the response to the treatment, in vivo bioluminescence imaging was performed before the first dose and then every 4 d until day 39, when the signals were saturated in the control cohorts. Cohorts 1 (no biAb) and 5 (HER2 × CD3 biAb) revealed aggressive tumor growth (Fig. 5). In cohort 2, which received R11 × v9, we observed significant tumor growth retardation and tumor eradication starting on day 14 after one dose of T cells and two doses of biAb, comparable to the CD19 × CD3 biAb in cohort 4 (Fig. 5 A and C). By contrast, R12 × v9 in cohort 3 revealed only weak activity compared with the negative control cohorts. As shown in Fig. 5B, no weight loss was observed during the treatment in the responding cohorts, including the R11-based ROR1 × CD3 biAb, which cross-reacts with mouse ROR1 (SI Appendix, Table S1). Weight loss in the nonresponding or weakly responding cohorts was noticeable with increasing tumor burden. The corresponding Kaplan–Meier survival curves showed that all treatment groups except for cohort 5 (HER2 × CD3 biAb) survived significantly longer than cohort 1 (no biAb). This included both ROR1 × CD3 biAb cohorts (R11 × v9, P < 0.001; R12 × v9; P < 0.05) and the CD19 × CD3 biAb cohort (P < 0.001; Fig. 5D). Although all mice with this aggressive xenograft eventually exhibited relapse, survival was longest in the CD19 × CD3 biAb cohort. As anticipated from the in vivo bioluminescence imaging, mice treated with the R11-based ROR1 × CD3 biAb significantly outlived mice treated with the R12-based ROR1 × CD3 biAb (P < 0.05).

Fig. 5.

In vivo efficacy of bispecific ROR1 × CD3 scFv-Fc. Five cohorts of mice (n = 8) were inoculated with 0.5 × 10⁶ JeKo-1/ffluc cells via i.v. (tail vein) injection. After 7 d, 5 × 10⁶ ex vivo expanded primary human T cells and 10 μg of the indicated biAbs or vehicle alone were administered by the same route. The mice received a total of three doses of T cells every 8 d and a total of six doses of biAbs or vehicle alone every 4 d. (A) Bioluminescence images of all 40 mice from day 6 (before treatment) and day 34 (after treatment) are shown. (B) The weight of all 40 mice was recorded on the indicated days (mean ± SD). (C) Starting on day 6, all 40 mice were imaged every 4–5 d and their radiance was recorded (mean ± SD). Significant differences between cohorts treated with biAbs (colored graphs) and vehicle alone (black graph) were calculated by using a two-tailed and heteroscedastic t test (**P < 0.01 and ***P < 0.001). Red arrows indicate the three T cell doses and black arrows the six biAb or vehicle-alone doses. (D) Corresponding Kaplan–Meier survival curves with P values [log-rank (Mantel–Cox) test] comparing survival between cohorts treated with biAbs (colored graphs) and vehicle alone (black graph; *P < 0.05 and ***P < 0.001).

We next carried out a pharmacokinetic (PK) study with R11 × v9 and R12 × v9 scFv-Fc in mice to examine their circulatory t_1/2 values. Five female CD-1 mice for each biAb group were injected i.v. (tail vein) with 6 mg/kg of the biAb. Blood samples were withdrawn at indicated time points (SI Appendix, Fig. S4) over a period of 2 wk, and plasma was prepared. The biAb concentration in the plasma was measured with a sandwich ELISA using immobilized ROR1 ECD for capture and peroxidase-conjugated goat anti-human IgG pAbs for detection. Analysis of the PK parameters by two-compartment modeling revealed t_1/2 values (mean ± SD) of R11 × v9 and R12 × v9, respectively, of 155 ± 23 h (6.46 ± 0.96 d) and 152 ± 52 h (6.33 ± 2.17 d; SI Appendix, Table S2). For comparison, the t_1/2 of glycosylated and aglycosylated [³⁵S]Met-labeled chimeric mouse/human IgG1 in BALB/c mice was determined to be 6.5 ± 0.5 d (27).

Ex Vivo Activity of ROR1 × CD3 biAbs.

We next tested the potency of ROR1 × CD3 biAbs against primary malignant B cells from 13 patients with untreated CLL. CLL PBMCs with very low autologous effector-to-target ratios typical for patients with CLL (SI Appendix, Table S3) were incubated with 6.6 nM R11 × v9 and R12 × v9 scFv-Fc. For comparison, we included equimolar concentrations of one-armed R11 and R12 scFv-Fc without the CD3-binding arm as well as the negative control HER2 × CD3 biAb. After 11 d, an average of 38%, 23%, and 13% CLL PBMCs were killed in the presence of R11 × v9, R12 × v9 scFv-Fc, and HER2 × CD3 biAb, respectively (Fig. 6A). Averages of 4.9% and 9.3% CLL PBMCs were killed in the presence of the one-armed R11 and R12 scFv-Fc, suggesting that the cytotoxicity is mediated by T cells rather than by other effector cells in the CLL PBMCs. However, high variability in the response to ROR1 × CD3 biAbs was noted among the 13 CLL PBMCs, with only R11 × v9, but not R12 × v9, revealing a significant difference vs. its one-armed counterpart (Fig. 6A). Nonetheless, there was a positive correlation between effector-to-target ratios and cytotoxicity for R11 × v9 and R12 × v9 scFv-Fc but not for the negative control HER2 × CD3 biAb (Fig. 6B).

Fig. 6.

Primary CLL cell killing mediated by ROR1 × CD3 scFv-Fc. (A) Percent CLL-specific killing by treatment condition after 11 d in culture calculated by the following formula: (% untreated CLL viability − % treated CLL viability)/(% untreated CLL viability) × 100. Each dot represents one sample and treatment condition. Color of data points represent patient-matched samples and correspond with colors in SI Appendix, Table S3. Mean ± SD values are shown. Asterisks indicate statistical significance by Dunn’s multiple comparisons test (**P < 0.01). (B) Spearman’s correlation between initial effector-to-target ratios in PBMC samples used and CLL cell-specific killing by R11 × v9 scFv-Fc (green), R12 × v9 scFv-Fc (blue), and HER2 × CD3 biAbs (red) after 11 d in culture.

In Vitro Activity of ROR1 × CD3 biAbs Toward Carcinoma Cell Lines.

To demonstrate the broader therapeutic utility of the ROR1 × CD3 biAbs beyond ROR1-expressing hematologic malignancies, we examined breast adenocarcinoma cell line MDA-MB-231 and renal cell adenocarcinoma cell line 786-O as ROR1-expressing target cells in the in vitro cytotoxicity assay. Prior analysis by flow cytometry confirmed that both carcinoma cell lines are recognized by all seven ROR1 × CD3 biAbs. Among these, the R12-, XBR1-402-, and ERR1-TOP43–based ROR1 × CD3 biAbs, which are the highest-affinity binders and have overlapping membrane-distal epitopes (SI Appendix, Table S1) (11), revealed the strongest binding to both carcinoma cell lines (Fig. 7A). By using the same cytotoxicity assay with ex vivo expanded human T cells as before (Fig. 3 C–E), all seven ROR1 × CD3 biAbs revealed dose-dependent killing of MDA-MB-231 and 786-O cells (Fig. 7 B and C). Negative control biAb, h38C2 × v9, was again inactive. Notably, despite its weaker binding, R11 × v9 significantly outperformed all other ROR1 × CD3 biAbs for both MDA-MB-231 and 786-O cells, revealing EC₅₀ values of 100 ng/mL (1 nM) and 120 ng/mL (1.2 nM), respectively (Fig. 7 B and C). Collectively, these data suggest that the membrane-proximal epitope targeted by R11 is a preferred site for T cell-engaging biAbs in scFv-Fc format.

Fig. 7.

Bispecific ROR1 × CD3 scFv-Fc targeting breast cancer and renal cell cancer cell lines. (A) The indicated bispecific ROR1 × CD3 biAbs were analyzed for binding to human ROR1-positive triple-negative breast adenocarcinoma cell lines MDA-MB-231 and renal cell adenocarcinoma cell line 786-O by flow cytometry at a concentration of 5 μg/mL followed by Alexa Fluor 647-conjugated goat anti-human IgG pAbs. Open histograms show the background binding signal of the secondary pAbs. The cytotoxicity of ROR1 × CD3 scFv-Fc was tested with ex vivo-expanded primary human T cells and MDA-MB-231 (B) or 786-O (C) cells at an effector-to-target ratio of 10:1 and measured after 16 h. Shown are mean ± SD values from independent triplicates.

Crystallization of R11 scFv in Complex with the Kr Domain of Human ROR1.

To elucidate the precise location of the R11 epitope, scFv R11 was crystallized in complex with the Kr domain of human ROR1 [Protein Data Bank (PBD) ID code 6BA5]. The complex crystallized as P1 space group with four scFv:Kr complexes in the asymmetric unit (SI Appendix, Table S4). Overall, the rmsd of atomic positions of each complex was <0.47 Å, indicating little variation among the independent structures. The Kr domain revealed a typical kringle domain folding that appears in various extracellular proteins (SI Appendix, Fig. S5). Notably, it showed higher structural homology with the Kr domains of secreted proteins compared with the recently reported first 3D structure of a Kr domain in the ECD of a transmembrane protein (28) (PDB ID code 5FWW). For example, the rmsd with human plasminogen Kr domain 4 (PDB ID code 1KRN) is only 0.66 Å. However, the shallow surface pocket known to host a free lysine or lysine analog in many Kr domains (29–31) is constricted in the ROR1 Kr domain as a result of an inward conformation of loop 5 and partial occupation by the side chain of Lys369 (SI Appendix, Fig. S5).

In the scFv:Kr complex, the N-terminal portion of β-strand A of the V_H domain undergoes domain swapping such that the first 6 aa integrate into the BED β-sheet of the neighboring V_H domain (Fig. 8A). The domain swapping results in crystallographic as well as noncrystallographic twofold symmetry between two neighboring complexes and is likely a crystallization artifact, as the presence of C_H1 and C_κ would block this dimerization. Also, it does not influence the binding of the ROR1 Kr domain, and purification by SEC before crystallization revealed the monomeric scFv:Kr complex as major peak.

Fig. 8.

Crystal structure of scFv R11 in complex with the Kr domain of ROR1. The 3D structure of scFv R11 in complex with the Kr domain of ROR1 was determined by X-ray crystallography at 1.6-Å resolution (PBD ID code 6BA5). (A) Ribbon model of the scFv:Kr complex showing twofold noncrystallographic symmetry. The scFv is shown in dark blue (V_H) and cyan (V_L), the Kr domain in orange. The domain swapping β-strand A of the V_H domain is shown in green. (B) Cartoon showing the location of the R11 scFv epitope on the Kr domain of ROR1. The dashed area corresponds to the crystal structure of the scFv:Kr complex shown on the right. Note that only the V_H domain (dark blue) interacts with the Kr domain. (C) Interaction of the three CDRs of the V_H domain (dark blue) with the Kr domain (orange surface model). The two enlarged areas show the salt bridges formed by HCDR1 (Asp30-Lys382) and HCDR3 (Asp102-Arg332), respectively. All interacting amino acid residues are listed in SI Appendix, Table S5.

The total buried surface area between scFv R11 and ROR1 Kr domain is 634 Ų, accounting for 6% and 13% of the total surface areas of scFv and Kr domain, respectively. Although the interaction involves multiple salt bridges, hydrogen bonds, and hydrophobic interactions, it is mediated solely by the V_H domain (Fig. 8B). No amino acid residue of the V_κ domain is involved in epitope recognition. Arg332 and Lys382 of the Kr domain form salt bridges with Asp102 (HCDR3) and Asp30 (HCDR1), respectively, of the V_H domain, enabling a strong interaction between the antibody and the antigen that is further stabilized by nine hydrogen bonds involving all three CDRs of the V_H domain (SI Appendix, Table S5). Furthermore, multiple residues of the CDRs make van der Waals interactions to establish a favorable shape complementarity with the Kr domain (Fig. 8C). These include favorable contacts of Asp27 (HCDR1) and Tyr100 (HCDR3) with Lys369 and Arg332, respectively, of the Kr domain, which delimits the paratope, and Tyr31 and Pro32 of HCDR1 with Phe381 of the Kr domain.

The amino acid sequence identities of the Kr domains of human ROR1 vs. human ROR2 and human ROR1 vs. mouse ROR1 are 62% and 99%, respectively (32, 33) (SI Appendix, Fig. S6). Fab R11 binds human ROR1 and mouse ROR1 with essentially the same association rate constant (k_on), dissociation rate constant (k_off), and K_d (2.7 vs. 3 nM) (10), but it does not cross-react with human ROR2. SI Appendix, Fig. S6, shows an amino acid sequence alignment of the three Kr domains with highlighted residues to indicate the epitope. These residues are highly diverse between human ROR1 and human ROR2 and completely conserved between human ROR1 and mouse ROR1, explaining the reactivity of mAb R11.

Discussion

Harnessing and enhancing the innate and adaptive immune system to fight cancer represents one of the most promising strategies in contemporary cancer therapy. The observation that T cell immunity plays a critical role in the immune rejection of many cancers is a key premise for cancer immunotherapy (34). A critical step of T cell-mediated eradication of tumor cells is the formation of cytolytic synapses (20) between T cells and tumor cells. This step can be mediated by biAbs combining a T cell-binding and activating arm with a tumor cell-binding arm. Numerous formats of T cell-engaging biAbs have been described and translated from basic to preclinical to clinical investigations (18). Thus far, however, only one T cell-engaging biAb, the CD19 × CD3 BiTE blinatumomab (16), has been approved by the FDA and marketed. Here we describe the generation and characterization of a panel of ROR1 × CD3 biAbs in a heterodimeric and aglycosylated scFv-Fc format that confers a circulatory t_1/2 of nearly 1 wk and eludes systemic T cell activation. Built on these attributes, our panel of ROR1 × CD3 biAbs that was based on seven different rabbit anti-human ROR1 mAbs (10, 11) potently and selectively killed ROR1-expressing MCL, breast cancer, and kidney cancer cells in the presence of primary T cells in vitro. A ROR1 × CD3 biAb that was based on a rabbit anti-human ROR1 mAb binding to a membrane-proximal epitope in the Kr domain was significantly more active in vitro and dramatically more active in vivo compared with ROR1 × CD3 biAbs with membrane-distal epitopes. Moreover, this ROR1 × CD3 biAb with membrane-proximal epitope significantly mediated the ex vivo killing of primary malignant B cells by autologous T cells at the very low effector-to-target ratios found in patients with CLL. Cocrystallization of the scFv in complex with the Kr domain revealed a discontinuous epitope that is conserved between human and mouse ROR1 and located in proximity to the transmembrane segment.

A recent study reported ROR1 × CD3 biAbs in BiTE format (35). The ROR1-binding arm was derived from two different rat anti-human ROR1 mAbs binding to an epitope in the Ig and Fz domain, respectively. The CD3-binding arm was derived from mouse anti-human CD3 mAb OKT3 (36), which shares an overlapping epitope on CD3δε with mouse anti-human CD3 mAb UCHT1 (37) and its humanized variant v9 (23, 38, 39), which we used for our ROR1 × CD3 biAbs in scFv-Fc format. Interestingly, the BiTE targeting the Fz domain was found to be significantly more potent than the BiTE targeting the Ig domain (35). Although a 3D structure of the ROR1 ECD has not been reported yet, it was assumed that the BiTE epitope in the Fz domain is in closer proximity to the cell membrane than the BiTE epitope in the Ig domain, and that this closer proximity may augment the formation of cytolytic synapses between T cells and tumor cells. An increased potency of T cell-engaging biAbs that bind to membrane-proximal epitopes on tumor cell antigens has also been reported for BiTE-based MCSP × CD3 in melanoma (40) and for IgG-based FcRH5 × CD3 biAbs in multiple myeloma (41). By using confocal microscopy, the latter study provided evidence that biAbs with membrane-proximal epitopes augment the formation of cytolytic synapses by promoting target clustering and exclusion of CD45 phosphatase from the cell–cell interphase, which triggers efficient ZAP70 translocation. It was shown that this process replicates the TCR-MHC/peptide-driven formation of cytolytic synapses (41).

Epitope location on ROR1 is also critical for the activity of CAR-T cells. An R12-based CAR with membrane-distal epitope was found to be more potent, with a shorter spacer between scFv and transmembrane segment, whereas a longer spacer was critical for the activity of an R11-based CAR with membrane-proximal epitope (12). These findings were confirmed with an XBR1-402–based CAR, which shares an overlapping epitope with R12, and an XBR2-401–based CAR, which binds to the Kr domain of human ROR2 (11). Collectively, the screening of a panel of mAbs with diverse epitopes is critical for identifying suitable candidates for T cell engagement by biAbs and CAR-T cells. Alternatively, if such panels are not available, customizing biAb and CAR formats may be required to incite activity. By determining the crystal structure of the R11 scFv in complex with the Kr domain of ROR1 at 1.6-Å resolution, we precisely mapped the epitope to a discontinuous epitope located near the C-terminus of the Kr domain, 20 aa upstream of the predicted transmembrane segment in the primary structure of ROR1 (32). Notably, the crystal structure revealed that only V_H of scFv R11 makes contacts, covering ∼13% of the surface area of the Kr domain. In addition to single-domain antibodies (42), our study encourages the development of R11 V_H-mimicking non-Ig scaffolds (43), such as affibodies (44) and designed ankyrin repeat proteins (45), to explore alternative building blocks for ROR1 × CD3 engagement.

Despite being smaller, the scFv-Fc format (∼100 kDa) mimics the natural IgG1 format (∼150 kDa) in using neonatal Fc receptor (FcRn)-mediated recycling for extended circulatory t_1/2. A recent study measured circulatory t_1/2 values in mice of ∼200 h for a chimeric mouse/human IgG1, ∼100 h for a chimeric mouse/human scFv-Fc, and ∼0.5 h for a mouse scFv, with all three sharing identical mouse V_L and V_H amino acid sequences (46). In approximate agreement, our PK study revealed a circulatory t_1/2 of ∼150 h for the ROR1 × CD3 scFv-Fc. This prolonged presence in the blood permitted a twice-weekly dosing in our in vivo experiment that likely can be further reduced to once weekly. By contrast, BiTEs, which are ∼50 kDa (scFv)₂ without Fc domain and have a circulatory t_1/2 of ∼2 h in humans (47), require continuous administration via, e.g., a portable minipump and i.v. port catheter. Although, in the case of blinatumomab, this setting has the advantage that treatment can be interrupted to address adverse events, particularly cytokine release syndrome and neurologic events (48), an increasing number of biAb formats in preclinical and clinical studies include an Fc domain to avoid continuous i.v. infusion (18). We chose an aglycosylated Fc domain to retain FcRn-mediated recycling but elude binding to Fcγ receptors CD16, CD32, and CD64 (49). Although this design excludes antibody-dependent cellular cytotoxicity and antibody-dependent cellular phagocytosis as contributing effector functions, it ensures that T cells are not activated systemically by Fcγ receptor-bound scFv-Fc that cross-link CD3 (50). The use of an aglycosylated Fc did not impact expression yields in mammalian cells, purification yields using standard Protein A affinity chromatography, monodispersity, or solubility of the scFv-Fc, affirming its suitability for manufacturing.

Despite limited insights into its role in cancer biology, the receptor tyrosine kinase ROR1 is an attractive target for antibody-mediated cancer therapy because of its highly restricted expression in a range of hematologic and solid malignancies (9). Postnatal healthy cells and tissues are largely negative for cell-surface ROR1 despite some exceptions (6–8, 51). Although solid malignancies have remained challenging indications for antibody therapeutic agents in general, targeting ROR1 in hematologic malignancies such as CLL, MCL, and subsets of acute lymphoblastic leukemia could circumvent concomitant depletion of healthy B cells by the FDA-approved CD19-targeting antibody therapeutic agents. Two early clinical trials are currently investigating ROR1 as a target for mAb and CAR-T cell therapy, respectively. ROR1 × CD3 biAbs, which combine off-the-shelf availability of mAbs with T cell-engaging potency of CAR-T cells, could provide a best-of-both-worlds option for patients with ROR1-expressing cancers. In view of its potency in vitro and in vivo, its recent humanization (52), and the precise mapping of its paratope and epitope by X-ray crystallography, mAb R11 has emerged as a prime candidate for further preclinical and clinical studies of ROR1 × CD3 biAbs.

Materials and Methods

Cell Lines and Primary Cells.

The K562, JeKo-1, and Jurkat cell lines were obtained from the American Type Culture Collection (ATCC) and cultured in RPMI-1640 supplemented with l-glutamine, 100 U/mL penicillin/streptomycin, and 10% (vol/vol) FCS (all from Thermo Fisher Scientific). The K562/ROR1 and JeKo-1/ffluc cell lines (6, 12) were provided by Stanley R. Riddell (The Fred Hutchinson Cancer Research Center, Seattle, WA) and Michael Hudecek (University of Würzburg, Würzburg, Germany), respectively. HEK 293 Phoenix (ATCC), MDA-MB-231 (ATCC), and 786-O cells (an NCI-60 panel cell line obtained from The Scripps Research Institute’s Cell-Based High-Throughput Screening Core) were grown in DMEM supplemented with l-glutamine, 100U/mL penicillin/streptomycin, and 10% (vol/vol) FCS. PBMCs were isolated from healthy donor buffy coats by using Lymphoprep (Axis-Shield) and cultured in X-VIVO 20 medium (Lonza) with 5% (vol/vol) off-the-clot human serum AB (Gemini Bio-Products) and 100 U/mL IL-2 (Cell Sciences). Primary T cells were expanded from PBMCs as previously described (31) by using Dynabeads ClinExVivo CD3/CD28 (Thermo Fisher Scientific). Cryopreserved treatment-naïve CLL PBMCs were derived from a natural history study at the National Institutes of Health (NIH) Clinical Center registered under ID code NCT00923507 at https://clinicaltrials.gov/. The study was approved by the institutional review boards of the NIH Clinical Center and The Scripps Research Institute. Informed consent was obtained in accordance with the Declaration of Helsinki. Clinical and laboratory data were collected from electronic medical records.

Cloning, Expression, and Purification of ROR1 × CD3 biAbs.

All amino acid sequences have been previously published or patented. The variable domain encoding cDNA sequences of the rabbit anti-human ROR1 mAbs and the humanized mouse anti-human CD3 mAb v9 were PCR-amplified from phagemids (10, 11) and previously described DART-encoding plasmids (53), respectively. A (Gly₄Ser)₃ linker encoding sequence was used to fuse V_L and V_H by overlap-extension PCR. The Fc fragment with a N297A mutation and either knob mutations, S354C and T366W, or hole mutations, Y349C, T366S, L368A, and Y407V, were synthesized as gBlocks (Integrated DNA Technologies). Anti–ROR1-Fc knob and anti–CD3-Fc hole-encoding fragments were assembled by overlap-extension PCR, respectively, and AscI/XhoI-cloned into mammalian cell expression vector pCEP4 and transiently cotransfected into HEK 293 Phoenix cells (ATCC) with polyethylenimine (Sigma-Aldrich) as described (54). Supernatants were collected three times over a 9-d period, followed by 1-mL Protein A HiTrap HP columns in conjunction with an ÄKTA FPLC instrument (both from GE Healthcare Life Sciences). Subsequent preparative and analytic (10 μg biAb) SEC was performed with a Superdex 200 10/300 GL column (GE Healthcare Life Sciences) in conjunction with an ÄKTA FPLC instrument using PBS solution at a flow rate of 0.5 mL/min. The purity of the biAbs was confirmed by SDS/PAGE followed by Coomassie blue staining, and their concentration was determined by measuring the absorbance at 280 nm. One-armed R11 and R12 scFv-Fc were cloned, expressed, and purified as described for the ROR1 × CD3 biAbs by replacing the anti–CD3-Fc hole with an empty Fc hole.

Flow Cytometry.

Flow cytometry was performed on a FACSCanto system (BD Biosciences), and data were analyzed with WinMDI 2.9 software. APC or Alexa Fluor 647-conjugated goat anti-human IgG pAbs were purchased from Jackson ImmunoResearch Laboratories. Alexa Fluor 647-conjugated mouse anti-human CD69 mAb was purchased from BioLegend. For the cross-linking assay, target and effector cells were labeled with CellTrace CFSE and CellTrace Far Red (both from Thermo Fisher Scientific), respectively, according to the manufacturer’s protocol. The labeled target and effector cells at a 1:1 ratio were then incubated with 1 µg/mL biAbs in 100 µL final volume. Following incubation for 1 h at room temperature, the cells were gently washed, fixed with 1% (wt/vol) paraformaldehyde, and analyzed by flow cytometry as described earlier.

In Vitro Cytotoxicity Assay.

Cytotoxicity was measured by using CytoTox-Glo (Promega) following the manufacturer’s protocol with minor modifications. Primary T cells expanded from healthy donor PBMCs as described here earlier were used as effector cells and K562/ROR1, JeKo-1, K562, MDA-MB-231, or 786-O cells were used as target cells. The cells were incubated at an effector-to-target ratio of 10:1 in X-VIVO 20 medium (Lonza) with 5% (vol/vol) off-the-clot human serum AB. The target cells (1 × 10⁴) were first incubated with the biAbs before adding the effector cells (1 × 10⁵) in a final volume of 100 µL per well in a 96-well tissue culture plate. The plates were incubated for 16 h at 37 °C with biAb concentrations ranging from 2 pg/mL to 15 µg/mL. After centrifugation, 50 µL of the supernatant was transferred into a 96-well clear-bottom white-walled plate (Costar 3610; Corning) containing 25 µL per well CytoTox-Glo. After 15 min at room temperature, the plate was read by using a SpectraMax M5 instrument with SoftMax Pro software. The same supernatants (diluted 20-fold) used for the CytoTox-Glo assay were also used to determine IFN-γ, TNF-α, and IL-2 secretion with the human IFN-γ, human TNF-α, and human IL-2 ELISA Ready-SET-Go! reagent sets (eBioscience), respectively, following the manufacturer’s protocols.

Mouse Xenograft Studies.

Forty 6-wk old NSG mice (The Jackson Laboratory) were each given 0.5 × 10⁶ JeKo-1/ffluc cells via i.v. (tail vein) injection on day 0. On day 7, each mouse was i.v. injected (tail vein) with 5 × 10⁶ primary T cells expanded from healthy donor PBMCs as described earlier and, 1 h later, with 10 µg biAbs or PBS solution alone. The mice received a total of three doses of expanded primary T cells every 8 d and a total of six doses of biAbs or PBS solution alone every 4 d. Every 4–5 d, tumor growth was monitored by bioluminescent imaging 5 min after i.p. injections with 150 mg/kg d-luciferin (Biosynth). For this, mice were anesthetized with isoflurane and imaged by using an Xenogen IVIS imaging system (Caliper) 6, 8, 10, 12, and 14 min after luciferin injection in small binning mode at an acquisition time of 10 s to 1 min to obtain unsaturated images. Luciferase activity was analyzed by using Living Image Software (Caliper), and the photon flux was analyzed within regions of interest that encompassed the entire body of each individual mouse. The weight of the mice was measured every 4–5 d. All procedures were approved by the institutional animal care and use committee of The Scripps Research Institute and were performed according to the NIH Guide for the Care and Use of Laboratory Animals.

PK Study.

Five female CD-1 mice (∼25 g; Charles River Laboratories) were injected i.v. (tail vein) with R11 × v9 or R12 × v9 at 6 mg/kg. Blood was collected at 5 min and 12, 24, 48, 72, 108, 168, 240, and 336 h aftetr injection with heparinized capillary tubes. Plasma was obtained by centrifuging the samples at 2,000 × g for 5 min in a microcentrifuge and stored at −80 °C until analysis. The concentrations of biAbs in the plasma samples were measured by ELISA. For this, each well of a 96-well Costar 3690 plate (Corning) was incubated with 200 ng recombinant human ROR1 ECD (10) in 25 μL carbonate/bicarbonate buffer (pH 9.6) at 37 °C for 1 h. After blocking with 150 μL 3% (wt/vol) BSA/PBS solution for 1 h at 37 °C, the plasma samples were added. Peroxidase AffiniPure F(ab′)₂ Fragment Goat Anti-Human IgG pAbs (Jackson ImmunoResearch Laboratories) were used for detection. The concentration of the biAbs in the plasma samples was extrapolated from a four-variable fit of the standard curve. PK parameters were analyzed by using Phoenix WinNonlin PK/PD Modeling and Analysis software (Pharsight).

Ex Vivo Cytotoxicity Assay.

Cryopreserved PBMCs were thawed and plated at 3 × 10⁶ cells per milliliter in 24-well plates in RPMI 1640 (Gibco), 10% FBS, 1% penicillin/streptomycin, 50 μM β-mercaptoethanol (Sigma-Aldrich). A total of 60 IU/mL of IL-2 was added to cultures 6–12 h after initial plating. Cells were then incubated with bispecific or one-armed scFv-Fc at 6.6 nM and harvested on day 11. Untreated wells containing cells and medium alone were left as background controls. Cell viability was assessed with LSRFortessa (BD Biosciences) by using LIVE/DEAD fixable violet stain (Thermo Fisher Scientific). To analyze the cell viability, CLL cells were identified as CD8⁻/CD4⁻/CD5⁺/CD20⁺ and T cells as CD8⁺ or CD4⁺ by staining with commercial mAbs (BD Biosciences): CD4 (RPA-T4), CD8 (HIT8a), CD20 (L27), and CD5 (L17F12). Specific killing was calculated by the following formula: (% untreated CLL viability − % treated CLL viability)/(% untreated CLL viability) × 100. Effector T cell to target CLL cell ratios were determined based on frequencies of live cells.

Statistical Analyses.

Statistical analyses were performed by using Prism software (GraphPad). The in vitro data shown in Fig. 4 were subjected to one-way ANOVA, and the in vivo data shown in Fig. 5C were analyzed with a two-tailed and heteroscedastic t test with a CI of 95%. Statistical analysis of survival (Fig. 5D) was done by log-rank (Mantel–Cox) testing. The ex vivo data shown in Fig. 6A were analyzed with a Dunn’s multiple comparisons test. Results with a P value <0.05 were considered significant.

Crystallization and Structure Determination.

Cloning, expression, and purification.

Leaderless R11 scFv and human ROR1 Kr domain (amino acids 319–391) encoding cDNA sequences were PCR-amplified from previously described phagemids and plasmids (10) and cloned as polycistronic assembly that also included a leaderless Escherichia coli disulfide bond isomerase C (DsbC), which is a chaperone that aids proper disulfide bond formation (55), into E. coli expression vector pET-15b (Novagen). The resulting expression cassette included a ribosome binding site (RBS) with a start codon, an N-terminal (His)₆ tag, a thrombin cleavage site, the Kr domain (flanked by NdeI/XhoI), a second RBS with a start codon followed by the scFv (flanked by XhoI/XhoI), and a third RBS with a start codon followed by DsbC (flanked by XhoI/BamHI). The prepared plasmid was transformed into E. coli strain Rosetta-Gami 2(DE3) (Novagen). Following induction of protein expression by 0.3 mM isopropyl-d-thiogalactoside, the bacterial cultures were grown in Luria-Bertani (LB) medium containing ampicillin, tetracycline, and chloramphenicol at 20 °C and 220 rpm in an incubator shaker for 18 h.

Protein purification.

The proteins were purified by immobilized metal ion chromatography followed by SEC in conjunction with an ÄKTA FPLC instrument. In brief, bacterial pellets were harvested by centrifugation and resuspended in sonication buffer [20 mM Hepes, pH 8.0, 300 mM NaCl, 15 mM imidazole, 10% (vol/vol) glycerol], sonicated in an ice-water bath, and centrifuged for 25 min at 53,300 × g. The supernatant was loaded on a custom-packed 10-mL HIS-Select column (Sigma-Aldrich) and washed with sonication buffer. The scFv:Kr complex was eluted with a linear gradient of 15–500 mM imidazole and treated overnight at 4 °C with thrombin (Sigma-Aldrich) to remove the N-terminal (His)₆ tag on the scFv. The clipped scFv:Kr complex was then purified further by SEC on a Superdex 200 26/60 column (GE Healthcare Life Sciences) equilibrated with 50 mM NaCl, 10 mM Hepes (pH 7.4). Fractions containing the complex were combined and concentrated to ∼15 mg/mL with a 10-kDa molecular weight cut-off Amicon ultrafiltration unit (Millipore).

Crystallization and structure determination.

Crystals of the purified scFv:Kr complex were grown by vapor diffusion at room temperature by using 1.5 µL of 15 mg/mL protein and an equal volume of precipitant containing 0.2 M ammonium fluoride in 20% (wt/vol) PEG 3350, and were fully grown within 2 d. The crystals were flash-frozen in liquid nitrogen by using nylon loops after removing excess mother liquor. A diffraction data set with Bragg spacings to 1.6 Å was collected on an ADSC Quantum 315r detector at the 24-ID-E beamline at the Advanced Photon Source, Argonne National Laboratory. Data were processed with iMOSFLM software (56). The structure of the scFv:Kr complex was solved by the molecular replacement method by using BALBES with 2ZNW as the search model (57). Crystallographic refinement was performed by using a combination of PHENIX 1.12 (58) and BUSTER 2.9 (59). Manual rebuilding and adjustment of the structure were carried out by using the graphics program Coot (60). Data processing and refinement statistics are shown in SI Appendix, Table S4. Molecular figures were created by using PyMOL software (Schrödinger). Buried surface area was calculated using PISA (61), and structure validation was carried out with MolProbity (62). The crystal structure was deposited in PDB under ID code 6BA5.