Human embryonic kidney 293 (HEK-293, CRL-1573) cells and HEK-293 cells stably transfected with the human chemokine receptor CCR9, or the empty vector (pCIneo) were a kind gift of A. Zaballos (CNB-CSIC, Madrid, Spain), cells were cultured as described (3). MOLT-4 (CRL-182) and Jurkat (TIB-152) human T-ALL cell lines were obtained from the American Type Culture Collection (ATCC). Cells were cultured in Dulbecco’s modified Eagle’s medium (Gibco) supplemented with 10% fetal bovine serum (FBS, Gibco), 2 mM l-glutamine, 50 U/ml penicillin, and 50 µg/ml streptomycin (complete medium). Neomycin-resistant stable HEK-293 transfectants were cultured in the presence of 1 mg/ml G418 (Sigma) and periodically tested for CCR9 expression (not shown). Recombinant human CCL25 and CXCL12 were purchased from Peprotech. We used the following antibodies: 3C3 (ATCC HB-12653), 112509, mouse mAb anti-hCCR9 (IgG2a; R&D) and M4, a serum pool generated by immunizing BALB/c mice with three intraperitoneal injections of 10⁷ MOLT-4 cells in PBS (days 1, 25, and 50); sera were collected on day 60.

Murine 91R and 92R anti-human CCR9 mAb were raised after immunization of BALB/c mice with a gene gun (Bio-Rad) particle-mediated DNA administration of the pCIneo plasmid bearing the human CCR9 cDNA, as previously described (34). Mouse sera were collected 7–10 days (d) after the last boost and tested for specific antibodies by flow cytometry using stably transfected hCCR9-HEK-293 cells, and pCIneo-HEK-293 cells as negative control. Selected mice were boosted intravenously with 10⁷ hCCR9-HEK293 cells 3 and 2 days prior to splenocyte fusion (35). Two weeks post-fusion, culture supernatants were screened by flow cytometry for CCR9-specific antibodies using hCCR9-HEK293 cells. Positive hybridomas were cloned, mAb purified from culture supernatants and antibody isotype determined by enzyme-linked immunosorbent assay (ELISA) (35).

For staining, 2 × 10⁵ cells/well were centrifuged in V-bottom 96-well plates and washed with phosphate-buffered saline, pH 7.4 (PBS) supplemented with 0.5% bovine serum albumin (BSA), 1% FBS, and 0.1% sodium azide (PBSst). Non-specific binding of the mAb to the cell surface was blocked by preincubating the cells with 40 µg/ml rat IgG (Sigma) in a 100 µl final volume (20 min, 4°C). Cells were incubated with the primary mAb (30 min, 4°C), washed, and the binding was revealed with a secondary FITC- or PE-goat F(ab’)2 anti-mouse IgG (H + L) antibody (Beckman Coulter; 30 min, 4°C). Samples were analyzed on an Epics XL or a Cytomics cytometer (Beckman Coulter). For competition analyses, cells were incubated with 50 µl of either the unlabeled antibody or an isotype-matched mAb (10 µg/ml, 40 min, 4°C), followed by 50 µl of an anti-CCR9 biotin-labeled antibody (0.5–2 µg/ml, 30 min, 4°C). After washing, FITC- or PE-conjugated streptavidin was added (30 min, 4°C). Cell staining was evaluated by flow cytometry.

Microtiter plates (Maxi-sorb, Nunc) were used to coat the hCCR9(2–22) synthetic peptide (1 µg/ml in PBS), overnight at 4°C. Afterward, the unoccupied protein-binding sites in the wells were blocked with 0.5% BSA in PBS. Previously titrated biotin-labeled mAb was mixed with unlabeled mAb (2 µg/ml in PBS–0.5% BSA), added to the plate and incubated 1 h, at room temperature. Antibodies bound to the plate were detected with peroxidase-labeled streptavidin (Sigma) and revealed with o-phenylenediamine dihydrochloride (4 mg/ml in 0.15 M sodium citrate buffer, pH 5.0; Sigma). The reaction was stopped with 3N sulfuric acid and the O.D. 492 nm determined. Antibodies were biotinylated with Hydrazide-LC-Biotin (ThermoFisher Scientific) following the supplier’s instructions.

Migration assays were performed in transwell inserts (Costar) with a 5-µm pore diameter. MOLT-4 cells were re-suspended in RPMI with 1% BSA and 25 mM HEPES, pH 7.4 (10⁷ cells/ml), and 100 µl aliquots were loaded into the upper inserts. Samples of 0–300 nM human CCL25, prepared in 600 µl of the same medium, were placed in the lower wells. After 2 h incubation at 37°C, 5% CO₂, inserts were removed and the number of cells that had migrated from the transwell insert to the well were counted on an EPICS XL flow cytometer. To analyze the antibody-blocking activity of CCL25-induced migration, MOLT-4 cells were preincubated with different amounts of anti-CCR9 mAb or irrelevant isotype-matched mAb before being loaded into the transwell. For these experiments, 200 nM human CCL25 was used as chemoattractant.

Linear peptides were synthesized in the Proteomic facility of the CNB, with an automated multiple-peptide synthesizer (AMS 422, Abimed) using a solid-phase procedure and standard Fmoc-chemistry. The synthesis of multiple peptides was performed simultaneously, on a cellulose membrane, by sequential conjugation of membrane-protected amino acids (aa), from their carboxy terminal ends. The application of the activated aa was carried out using the Auto-Spot Robot (ASP222, Abimed) (36). Two independent membranes were prepared with the same set of peptides (each 12 aa long, 10 aa overlap), spanning the complete hCCR9-A isoform (369 aa). In addition, a similar synthesis was carried out where to peptides corresponding to aa 8–19 of hCCR9-A, aa in positions 11–16, each one of them was substituted for each of the remaining proteinogenic aa. Membranes were blocked by incubation with 1% BSA in PBS for 1 h at room temperature, washed, and then incubated for 2 h with anti-CCR9 or isotype control antibodies (in PBS containing 1% BSA and 0.05% Tween-20). After three additional washes, membranes were incubated with a peroxidase-labeled goat anti-mouse IgG antibody (Sigma), for 1 h at room temperature, developed with a chemiluminescence system (GE Healthcare), and exposed to standard X-ray film. The densitometric quantification of the signal obtained at each spot was performed with the ImageJ software.

Surface Plasmon Resonance experiments were carried out in a biosensor Biacore 3000 (Biacore, GE Healthcare), using HBS-EP (10 mM HEPES, 0.15 M NaCl, 3 mM EDTA, 0.005% Surfactant P20, pH 7.4) as running buffer. At the end of each binding cycle, the sensor surface was regenerated with 10 mM glycine–HCl, pH 1.5, allowing resonance signals to return to baseline values. A synthetic peptide corresponding to the aa in positions 2–22 of the human CCR9 isoform A (hCCR9A, identifier: P51686-1), with an additional cystein residue in its N terminus (C-TPTDFTSPIPNMADDYGSEST) was immobilized on a carboxymethylated dextran CM5 sensor chip by an amine coupling reaction, as recommended by the supplier. A reference surface was generated in the same manner, except that all carboxyl groups were blocked in the absence of ligand. For kinetic analyses, immobilized antigen at low density was used to minimize mass transport effects and analyte rebinding. Anti-CCR9 mAbs were used as soluble analytes in HBS-EP buffer at concentrations ranking from 0.41 to 33 nM. The interaction analyses were carried out at 25°C with a flow rate of 30 µl/min. Data were collected for 90 s of association and 180 s of dissociation. For competition analyses, a biotin-labeled peptide corresponding to aa 2–19 of hCCR9A (biotin-K-TPTDFTSPIPNMADDYGS) was captured on a streptavidin-coated chip (SA). Anti-CCR9 mAbs at a constant concentration (10 nM) were mixed with competitor synthetic peptides at different concentrations (0.01–1 µM). The antibody–peptide mixtures were preincubated for 30 min before they were injected into the biosensor. Sensograms were overlaid, aligned, and analyzed with BIA evaluation Software 4.1. The K_D was determined by fitting the data using a bivalent analyte model. All data set were processed using a double-referencing method (37).

BALB/c Rag2^−/− mice (Taconic Bioscience) were bred in the CNB animal facility and used at ages ranging from 8 to 22 weeks. For in vivo experiments, MOLT-4 cells (2 × 10⁶) were inoculated sub-cutaneously (s.c.) in the flank of Rag2^−/− mice on day 0. In these experimental conditions, 80–90% of the cell inoculations gave rise to tumors. The animals carrying the MOLT-4 cells were divided into four groups, which were inoculated intraperitoneally (i.p.) with anti-hCCR9 91R, its isotype control (IgG2b), anti-hCCR9 92R, or its isotype control (IgG2a) on days 1, 8, 15, and 22 (4 mg/kg on day 1 and 8; 2 mg/kg on days 15 and 22). Tumor size was measured with a Vernier caliper (Mitutoyo) and tumor volume (mm³) calculated as V = [axial diameter length, mm] × [(rotational diameter, mm)²/2]. Mice were sacrificed and tumors were weighted and processed for histology. Tumor burden is expressed as percent tumor weight relative to that of isotype control-treated mice.

NOD.Cg-Prkdc^scid Il2rg^tm1Wjl/SzJ stock # 005557 (NSG mice, Jackson Laboratories, ME, USA) were bred in the CIB animal facility and used for similar experiments as described in the above paragraph. Animals were injected subcutaneously on day 0 with MOLT-4 cells (1 × 10⁶) in the flank. Two groups were inoculated intra-peritoneally with 4 mg/kg on day 2 and day 7 with anti-hCCR9 92R or its isotype control (IgG2b). All mice were sacrificed at the same time and tumors were weighted and processed for histology.

MOLT-4 cells (10⁵ cells in 100 µl) were plated on each well of a 96-well V-bottom plate. The cells were incubated with the indicated concentrations of 92R, 91R (anti-hCCR9) or isotype-matched control mAb (30 min, 37°C), centrifuged, and washed. Active or 56°C heat-inactivated baby rabbit complement (25%; AbD Serotec) was added in serum-free Dulbecco’s modified Eagle’s medium with 1% BSA (1 h, 37°C). The complement in M4 serum was also heat inactivated (56°C, 30 min). The number of non-viable cells was evaluated by flow cytometry after staining the cells with the viability exclusion marker 7-aminoactinomycin (7-AAD; BD Biosciences; 10 min, 4°C); each condition was analyzed in triplicate. Specific lysis was calculated as: 100 × (% dead cells with active complement − % dead cells with inactive complement)/(100% − % dead cells with inactive complement).

Murine NK cells were isolated from spleens from BALB/c mice using the Auto Macs Pro negative selection system (Miltenyi). After purification, cells were analyzed for mCD3, mCD45 and mCD49b expression by flow cytometry. The purity of NK cells, defined as CD3⁻CD45⁺CD49b⁺, in all the preparations was at least 90%. Cells were cultured for 6–7 days in RPMI 1640 (Lonza), supplemented with 10% FBS, and 1,000 U/ml murine recombinant IL-2 (Peprotech). For cytotoxicity assays, target MOLT-4 cells labeled with Cell Trace CFSE (Invitrogen) were preincubated (30 min) with the indicated mAb concentrations. NK and target cells were cocultured (4 h) at a 20:1 ratio in RPMI–10% FBS, then stained with 7-AAD (10 min, 4°C) and analyzed by flow cytometry. Gating on 7-AAD-positive cells within the CFSE⁺ population indicated the proportion of dead target cells. Specific killing was calculated as: 100 × [(% dead target cells in sample − % spontaneous dead target cells)/(100 − % spontaneous dead target cells)]. Target cells incubated without effector cells were used to assess spontaneous cell death.

Statistical analyses were performed using GraphPad Prism 4 software. Statistical significance was established at P < 0.05 as evaluated by Student’s t-test, unless otherwise indicated. Results are shown as mean ± SEM.

The murine anti-hCCR9 mAb 92R (IgG2a) was generated after gene gun immunization with the full-length hCCR9-A cDNA coding sequence, inserted in a eukaryotic expression vector (pCINeo), using the same strategy as described for 91R mAb (34). Specificity of the binding of 92R to hCCR9-A protein was assessed by flow cytometry analyses of HEK-293 cells stably transfected with the construct used for immunization, using as negative control the same cells transfected with the empty pCINeo plasmid (Figure 1A). In addition, we demonstrated that 92R, similarly to 91R also stained cells expressing the endogenous CCR9 protein in the MOLT-4 T-ALL cell line, but failed to stain the Jurkat T-ALL cells that do not express CCR9 on their cell surface (Figure 1B).

Furthermore, competition analyses demonstrated that 92R competes with itself and 91R for binding to the CCR9⁺ cells MOLT-4, but not with the 3C3 anti-hCCR9 mAb in flow cytometry. Similarly, 91R competes with itself and 92R but not with 3C3 mAb, as demonstrated by flow cytometry (Figure 2A). These data were corroborated by ELISA assays, where binding of biotinylated 91R to the hCCR9 (aa 2–22) synthetic peptide was competed by unlabeled 92R, but not by the isotype control antibody IgG2a. Similarly, binding of biotinylated 92R to the same peptide was competed by unlabeled 91R, but not by the isotype control antibody IgG2b (Figure 2B).

In addition, 92R fails, similarly to 91R (34), to inhibit CCL25-induced migration of CCR9⁺ MOLT-4 cells, unlike the anti-CCR9 mAb 3C3, described to inhibit the CCL25:CCR9 interaction. Indeed, we determined the migration response of MOLT-4 cells to different CCL25 concentrations, observing that it reached the maximum level between 200 and 250 nM CCL25 (Figure 3A). Then, using 200 nM CCL25, the percentage of migrating cells in the absence or presence of different antibodies anti-hCCR9 or their isotype controls was determined. The results show a migration inhibition only in the presence of 3C3 mAb, but neither 92R nor 91R (nor their isotype controls) inhibited this migration (Figure 3B).

Since 92R and 91R are able to cross-compete with each other for the binding of hCCR9 on the cell surface (Figure 2A), and both of them bind the same epitope, comprised by aa 2–22 of hCCR9-A (Figure 2B), we aimed to identify the energetically critical aa for the interaction “hotspots” within the epitope. For this reason, pepscan analyses were carried out using 180 overlapping synthetic peptides, covering the entire hCCR9 sequence. These experiments allowed to identify the hotspot residues within the epitope, energetically critical for high-affinity binding of each antibody. Indeed, both antibodies gave positive signals with the same peptides 3–6 (Figure 4A), allowing to identify aa 11–16 from hCCR9-A as the functionally critical residues on the epitope recognized by 92R and 91R (Figure 4B). It is worth to note that although the peptides identified by these mAb are the same, the signals obtained with 91R and 92R are somehow different from each other (Figure 4A). Subsequently, each one of the aa from hotspots (sequence PNMADD) was substituted for one of the 19 other aa, allowing to ascertain the relative relevance of each one of the energetically critical aa for the binding of 91R and 92R to the hCCR9 synthetic peptides (Figure 4C). For both 91R and 92R mAbs, turned out that in these assays, the binding of these antibodies to the hotspots are strictly dependent of the presence of an N residue in position 12. The only replacement allowed at position 16 is a D for an E, indicating the relevance of a negative charge at this position. A14 could be replaced by hydrophilic uncharged aa (N, Q, or S) with a significant reduction of signal intensity. The few allowed changes for P11, unlike for A14, are more evident for 91R. Conversely, M13 and D15 allow more changes, with appreciable signal differences between 92R and 91R. For both mAb, the substitution of D15 for K or P is not allowed. The substitution of D15 for R is allowed for 91R but 92R loses its binding capacity. Taken together, these data support the notion that 91R and 92R are different antibodies, since the allowed changes on each position were different. This was further corroborated by sequence analyses of the variable regions from the light and heavy chain cDNAs (Figure 5). Indeed, there are five aa differences on the heavy chain framework and ten on the light chain framework. In addition, on the heavy chain, there is an aa change in CDR1, and on the light chain, there are two aa changes in CDR1, one in CDR2 and another one in CDR3 (Figure 5).

Surface Plasmon resonance analyses using the synthetic surface-bound peptide hCCR9 (2–22) as molecule representative of CCR9, allowed to determine the apparent affinity constants for these antibodies, which was estimated as K_D 3.6 nM for 91R and K_D 8.9 nM for 92R (Figures 6A,B). Competition experiments showed that 1 µM of peptide hCCR9 (2–22) inhibited most of the binding of 92R to the surface and fully inhibited the 91R binding; whereas peptide hCCR9 (13–30), corresponding to the N-terminus of the B isoform, failed to inhibit mAb binding. The competition with the same concentration of the peptide hCCR9 (10–30), which also contains the hotspots, gave, however, a slightly different competition pattern between 92R and 91R, since for 92R, the sensograms were similar when the competitors were hCCR9(10–30) or hCCR9(2–22), whereas a 20% of the 91R signal was still detected at the end of the association phase in the presence of hCCR9(10–30) peptide (Figures 6C,D). Taken together, these data show that the two antibodies 91R and 92R are able to bind differentially to the same hCCR9 epitope with high affinity.

The antitumor potential of 92R mAb was assessed in immunodeficient (Rag2^−/−, BALB/c) mice, after subcutaneous injection of 2 × 10⁶ CCR9⁺ cells from the human T-ALL cell line MOLT-4. On days 1, 8, 15, and 22 after tumor cell injection, the animals were treated with either 92R, 91R, or their isotype controls (IgG2a or IgG2b, respectively) at 4 mg/kg (days 1 and 8), or 2 mg/kg (days 15 and 22). The size of developing tumors was measured until day 78, when mice were sacrificed. The differences between tumor size were significant on 91R-treated mice as compared to isotype-treated mice from day 60 (P < 0.05) (Figures 7A,C). On 92R, we failed to detect tumors, although they grew in their control isotype-treated mice (Figures 7A,D). At the time of sacrifice, tumors were removed and weighted. The mean tumor weight for IgG2b isotype-treated control was 1.23 ± 0.33 g and for IgG2a, isotype-treated control was 1.25 ± 0.39 g, whereas for 91R-treated mice was 0.11 ± 0.06 g (Figure 7B) and were absent in animals treated with 92R (Figures 7B,D). As a positive control, 91R was used, where we found a reduction of 91% on tumor burden, in agreement with previously published data (34). As mentioned above, for 92R, no tumors were detected after sacrificing the animals, although 6/9 animals treated with its isotype control (IgG2a) had tumors (Figure 7D). These data support that 92R efficiently blocked the in vivo progression of acute T cell leukemia xenografts and suggest the possibility of using this antibody for therapeutic purposes in human CCR9⁺ tumors.

Complement-dependent cytotoxicity and ADCC are two of the main mechanisms in vivo for tumor cell elimination by therapeutic antibodies. For this reason, we tested the in vitro ability of 92R to induce lysis of MOLT-4 leukemia cells by either complement fixation, or by NK cell mediated cell cytotoxicity triggered by binding of 92R mAb to the NK cell surface Fc receptors.

For the CDC experiments, specific death of the MOLT-4 cells was evaluated by flow cytometry analyses of 7-AAD incorporation. Both 92R and 91R were able to promote complement-dependent cell lysis (46 ± 1 and 49 ± 2%, respectively) with a much higher efficiency than the commercial anti-CCR9 antibody 112509 (7.1 ± 0.9%) (Figures 8A,B). As positive control, we used a mouse sera (M4) against MOLT-4, which gave a specific lysis of 60 ± 0.8%. The minimal concentration of 92R mAb that gave a detectable specific cell lysis was 0.4 µg/ml.

For ADCC experiments, MOLT-4 target cells labeled with the green fluorescent dye CFSE and precoated with either 92R, or isotype control (IgG2a, negative control) were combined with previously in vitro activated mouse NK cells (4 h at 37°C). The incorporation of 7-AAD by the CFSE-labeled cells allowed to determine specific NK-mediated killing of MOLT-4 cells by flow cytometry. On these experiments, 91R mAb was used as positive control. Both 92R and 91R were able to promote NK-dependent cell lysis of the cells (57.7 ± 1.1 and 54.9 ± 4.1, respectively) with a much higher efficiency than the commercial anti-CCR9 antibody 112509 used as a control (20.5 ± 0.9%) (Figures 8C,D). As an additional positive control, we used the mouse serum M4 raised against MOLT-4, which gave a specific lysis of 91.1 ± 1.1%. The 92R mAb concentration needed for a detectable response was 0.04 µg/ml, whereas it was negligible in the absence of antibody. 91R was used as positive control for ADCC, obtaining similar results to the previously published (34).

Taken together, these results suggest a role for CDC and ADCC in the in vivo reduction of tumor growth observed in the xenograft model. To directly determine the in vivo role of CDC and ADCC in the antitumoral potential of 92R mAb, we generated subcutaneous xenografts by injection of 1 × 10⁶ CCR9⁺ cells from the human T-ALL cell line MOLT-4 in NSG mice. We used this particular mouse strain because its complement is not functional (due to a mutation in the C5 gene), and their NK activity is highly diminished (Jackson Labs, Bar Harbor, MD, USA). Therefore, if 92R was able to inhibit tumor growth in this model, the mechanism(s) of in vivo tumor reduction would be different from CDC and ADCC. On days 2 and 9 after tumor cell injection, the animals were treated with either 92R or its isotype control (IgG2a) at 4 mg/kg. On day 31, mice were sacrificed (Figure 9A), tumors removed, and weighted. The mean weight of the tumors was smaller in 92R-treated mice (0.256 ± 0.22 g) as compared to control isotype-treated mice (0.776 ± 0.53 g) (Figure 9B); therefore, the total tumor burden, measured as the mean of the tumor weights for each group, was reduced by 67% on animals treated with 92R, as compared to control isotype-treated animals, which correlated with the pictures taken of the tumors (Figure 9C). These data support the notion that mechanisms distinct from CDC and ADCC may play a role on the inhibition of acute leukemia tumor growth mediated by 92R in vivo in xenografts generated by human CCR9⁺ tumors. However, in vitro experiments analyzing whether 92R could mediate the inhibition of tumor growth through apoptosis (Figure S1 in Supplementary Material) or phagocytosis (Figure S2 in Supplementary Material) gave negative results.