. Author manuscript; available in PMC: 2020 Aug 31.

Published in final edited form as: Nat Med. 2019 Apr 29;25(5):759–766. doi: 10.1038/s41591-019-0420-8

Rational design of anti-GITR-based combination immunotherapy

Roberta Zappasodi ^1,², Cynthia Sirard ³, Yanyun Li ^1,², Sadna Budhu ¹, Moshen Abu-Akeel ¹, Cailian Liu ¹, Xia Yang ¹, Hong Zhong ¹, Walter Newman ³, Jingjing Qi ^2,⁴, Philip Wong ^2,⁴, David Schaer ¹, Henry Koon ⁵, Vamsidhar Velcheti ⁶, Michael A Postow ^7,⁸, Margaret K Callahan ^2,^7,⁸, Jedd D Wolchok ^1,^2,^7,^8,^*,^†, Taha Merghoub ^1,^2,^7,^*,^†

PMCID: PMC7457830 NIHMSID: NIHMS1616665 PMID: 31036879

Modulating T-cell homeostatic mechanisms with checkpoint blockade can efficiently promote endogenous anti-tumor T-cell responses^1–11. However, many patients still do not benefit from checkpoint blockade¹², highlighting the need of targeting alternative immune pathways¹³. Glucocorticoid-induced TNFR-related protein (GITR) is an attractive target for immunotherapy, due to its capacity to promote effector T-cell (Teff) functions^14,15 and hamper regulatory T-cell (Treg) suppression^16–20. Based on the potent preclinical anti-tumor activity of agonist anti-GITR antibodies, reported by us and others^16,21,22, we initiated the first in-human phase-I trial of GITR agonism with the anti-GITR antibody TRX518 (NCT01239134). Here, we report the safety profile and immune effects of TRX518 monotherapy in advanced cancer patients and provide mechanistic preclinical evidence to rationally combine GITR agonism with checkpoint blockade in future clinical trials. We demonstrate that TRX518 reduces circulating and intratumoral Tregs to similar extents, providing an easily assessable biomarker of anti-GITR activity. Despite Treg reductions and increased Teff:Treg ratios, substantial clinical responses were not seen. Similarly, in mice with advanced tumors, GITR agonism was not sufficient to activate cytolytic CD8⁺ T cells due to persistent exhaustion. We demonstrate that T-cell reinvigoration with PD-1 blockade can overcome resistance of advanced tumors to anti-GITR monotherapy. These findings led us to start investigating TRX518 with PD-1 pathway blockade in patients with advanced refractory tumors (NCT02628574).

TRX518 is a fully humanized Fc-dysfunctional aglycosylated IgG1κ monoclonal antibody that triggers human GITR signaling²³. In this phase-I trial, TRX518 was tested as a monotherapy in 43 patients with refractory solid tumors after a median of 3 prior therapies (range 0–9), including immunotherapy in 12 patients (29.3%)(Supplementary Table 1-2). Patients were sequentially enrolled in 1 of 9 dosing cohorts to receive a single intravenous administration of TRX518 (0.0001–8.0 mg/kg) and were followed to determine TRX518 safety, pharmacokinetics, pharmacodynamics, and immune effects.

Based on the preferential expression of GITR on natural killer (NK) and T cells, in particular Tregs^24,25, and considering that the therapeutic activity of GITR agonist antibodies in mouse tumor models has been associated with reduction and functional modulation of intra-tumor Tregs and potentiation of anti-tumor CD8⁺ T-cell functions^{16,17,21,26,27}, we focused the pharmacodynamic analysis of TRX518 on these immune cell populations. We assessed modulation of T-cell subsets and NK cells in serial peripheral blood mononuclear cell (PBMC) samples collected up to 12 weeks after therapy from patients treated in the 7 highest dose cohorts (n=37; cohort 3–9; 0.005–8 mg/kg TRX518) (Extended Data Fig. 1a). NK, CD8⁺, or CD4⁺Foxp3⁻ Teff cells showed no significant and/or consistent modulation (Fig. 1a, Extended Data Fig. 1b,c); however, Tregs were frequently reduced, with few exceptions in lung cancer patients (Fig. 1b, Extended Data Fig. 1b-d). Treg reduction was not due to hampered Treg proliferation capacity, since the frequency of proliferating Ki67⁺ Tregs did not substantially decrease after TRX518 (Fig. 1b). We then assessed modulation of Tregs expressing GITR, which are a more specific target of TRX518. To measure GITR expression, we used an antibody that was not cross-blocked by TRX518 (Extended Data Fig. 2a). We found an overall dose-dependent trend for decreasing GITR⁺ Tregs with increasing TRX518 dose (Fig. 1b, Extended Data Fig. 1b,c and Extended Data Fig. 2b). GITR⁺CD8⁺ T cells and GITR⁺CD4⁺Foxp3⁻ Teff were less consistently modulated by TRX518 (Extended Data Fig. 2c), likely due to their significantly lower GITR expression levels (Expended Data Fig. 2d). In cohort-5 and cohort-7 patients, who showed the strongest decreases in GITR⁺ Teff (Extended Data Fig. 2c), we found that GITR⁺ Teff frequently down-regulated CD127 (IL7R) and occasionally CD25 (IL2RA) after TRX518; whereas GITR⁺ Tregs more frequently down-regulated CD25 after treatment, pointing to potentially distinct mechanisms regulating fitness and survival of activated GITR⁺ Teff and Tregs during GITR stimulation (Extended Data Fig. 3).

Figure 1. — Fold changes in CD3⁻CD56⁺ NK cells (percentage of single live cells), CD3⁺CD8⁺ T cells (percentage of CD3⁺), CD3⁺CD4⁺Foxp3⁻ Teff (percentage of CD3⁺) (a), and (b) fold changes in the indicated CD3⁺CD4⁺Foxp3⁺ Treg subsets (total Foxp3⁺ Tregs, percentage of CD3⁺CD4⁺; Ki67⁺ and GITR⁺ Tregs, percentage of Tregs) at the indicated time points after treatment with TRX518 relative to baseline (mean ± SEM in patients grouped by dose cohort for each time point). Cell subsets were gated upon exclusion of doublets, CD19⁺, CD14⁺ and dead cells. (c) Representative gating strategy of human eTregs and nTregs in peripheral blood (PB) and tumor (TM) and quantification of eTregs frequency (percentage of CD3⁺CD4⁺) in PB (melanoma, n=6; lung cancer, n=7) and TM (melanoma, n=10; lung cancer, n=10) from immunotherapy-naïve melanoma and non-small cell lung cancer (NSCLC) patients (mean ± SEM; two-sided unpaired t test; melanoma, p=0.009; NSCLC, p=0.006). (d) GITR expression levels (median fluorescence intensity, MFI) at baseline in CD45RA⁻Foxp3^hi Tregs (eTregs) and CD45RA⁺Foxp3^lo Tregs (nTregs) in PB from patients enrolled in cohort 8 and 9 where GITR MFI was evaluable (10 of 11 patients; two-sided paired t test; p=0.006). (e) Mean ± SEM fold change of eTregs and nTregs (percentage of CD3⁺CD4⁺) in patients grouped by dose cohort at the indicated time points after treatment with TRX518 relative to baseline. Wk, week; hrs, hours. ** = p<0.01.

Considering that TRX518 specifically affected GITR⁺ Tregs and that GITR is an activation marker, we reasoned that activated functionally suppressive effector Tregs (eTregs), defined by high Foxp3 expression and low CD45RA levels (Fig. 1c)²⁸, should also be down-regulated by TRX518. We first confirmed that circulating CD4⁺CD45RA⁻Foxp3^hi eTregs with respect to naïve CD4⁺CD45RA⁺Foxp3^lo Tregs (nTregs) express significantly higher GITR levels (Fig. 1d). According to our hypothesis, we found that TRX518 more potently reduce eTregs than nTregs (Fig. 1e, Extended Data Fig. 1b,c). In addition, within the pool of GITR⁺ Tregs, the fraction of cells expressing high Foxp3 levels were more affected by TRX518 (Extended Data Fig. 4). These results indicated that TRX518 selectively targets highly activated functionally suppressive Tregs expressing GITR and elevated Foxp3 levels.

Using specific in vitro assays, we found that TRX518 counteracts Treg development and function and promotes T-cell co-stimulation (Extended Data Fig. 5a,b), as also reported for other human GITR agonist antibodies^29–31. Furthermore, we found that these effects are associated with loss of Treg phenotypic stability and enhanced cell death in particular in activated proliferating Tregs (Extended Data Fig. 5c-e). Importantly, as opposed to the physiologic GITR ligand, which induced cell death in both Teff and Tregs, TRX518 selectively increased cell death in Tregs (Extended Data Fig. 5e). These effects suggested that TRX518 favors induction of Treg cell death by promoting Treg hyper-activation and loss of phenotypic stability (Extended Data Fig. 5f). In line with these results, circulating eTregs in patients treated with TRX518 showed diminished survival potential, as indicated by BCL2L1 (encoding BCL-XL) down-regulation (Extended Data Fig. 5g).

Since eTregs are generally enriched at the tumor site^32,33, as verified in our experimental setting (Fig. 1c), we asked whether intra-tumor Tregs were also reduced upon TRX518. Of the 37 patients for whom we had peripheral Tregs FACS data, we also were generously provided by 8 patients with paired pre- and post-treatment biopsies with the post-treatment biopsy and peripheral blood sample for FACS analysis collected at the same time point. These included biopsies from 3 melanoma, 2 lung, 2 colorectal and 1 bladder cancer patients (Extended Data Fig. 1a). Consistent with Treg modulation in peripheral blood, intratumoral Tregs were frequently reduced after treatment, with the exception of 2 lung cancer patients, where Tregs were not down-regulated in either peripheral blood or tumor (Fig. 2). Treg reduction was not proportional to the quantity of tumor-infiltrating Tregs at baseline, as intratumoral Tregs were down-regulated in patients starting TRX518 with either low (0102–0005, 0002–0006 and 0102–0003) or higher (0273–00022, 0102–0002, 0002–0009) levels of Tregs (Fig. 2b). Importantly, in these patients, fold reduction in intratumoral and circulating Tregs directly correlated (Fig. 2c, Extended Data Fig. 6), suggesting that assessing modulations in peripheral Tregs after TRX518 may allow to predict changes in intratumoral Tregs.

Figure 2. — (a) Fold changes in circulating (PB) CD3⁺CD4⁺Foxp3⁺ Tregs (percentage of CD3⁺CD4⁺) at the indicated time points after TRX518 relative to baseline in 8 patients for whom pre- and post-tumor biopsies were also available for Treg analysis. Mean values of technical duplicates at each time point are shown for each patient. (b) CD4⁺Foxp3⁺ Treg number normalized on tumor area (μm²) in pre- and post-treatment biopsies by IF from the same patients shown in a. Post-treatment biopsy collection time point relative to treatment initiation is indicated for each patient. (c) Pearson correlation analysis between peripheral (PB) and intratumoral (TM) Treg fold changes at the same time point relative to baseline in 8 patients as in a and b [p (two-tailed)= 0.009]. Wk, week; hrs, hours.

Concordant down-regulation of Tregs in peripheral blood and tumor upon TRX518 was not sufficient to mediate substantial clinical responses in this patient population. Twenty-nine patients experienced disease progression and 4 achieved stable disease (Supplementary Data Table 3), with 2 being lost at week-12 follow up and 1 relapsing 18 weeks after treatment initiation (Supplementary Data Table 4).

We thus investigated possible mechanisms of resistance of advanced tumors to anti-GITR immunotherapy in mice. We modeled tumor sensitivity and refractoriness to GITR agonism by treating B16F10-melanoma bearing mice with the anti-GITR antibody DTA-1 4 days (curative regimen, early tumors) or 7 days (refractory regimen, advanced/established tumors) after tumor implantation (Fig. 3a). By performing a time-course analysis of T-cell infiltration up to 6 days after treatment (Fig. 3b schema) – the in vivo half-life of DTA-1²⁶ – we found that intra-tumor Tregs were significantly reduced and Teff:Treg ratio increased in both responding and refractory tumors (Fig. 3b, Extended Data Fig. 7a). However, in responding tumors treated at an early stage, Tregs completely failed to accumulate (Fig. 3b), suggesting that the presence of Tregs during tumor formation and progression could affect T-cell functionality. We confirmed the therapeutic advantage of abrogating intra-tumor Treg accumulation early during tumor formation by comparing day-4 versus day-7 Tregs depletion with diphtheria toxin (DT) injection in Foxp3-DT-receptor transgenic mice (Extended Data Fig. 7b). Interestingly, the kinetics and extent of Treg depletion and subsequent recovery did not differ between the two DT injection schedules (Extended Data Fig. 7b). We also found that in untreated 7-day-old tumors, compared to 4-day-old tumors, CD8⁺ T-cell:Treg ratios were reduced, Tregs expressed higher Foxp3 and CD25 levels, and CD8⁺ T cells down-regulated GITR expression and IFN-γ and TNFα production (Extended Data Fig. 7c). This CD8⁺ T-cell dysfunction could not be overcome if GITR stimulation and Treg reduction occurred in the setting of day-7 advanced tumors, but could be prevented with earlier anti-GITR treatment (Fig. 3c). Preventing intra-tumor Treg accumulation by treating mice 4 days after tumor implantation resulted in intra-tumor infiltration with CD8⁺ T cells expressing a non-exhausted profile and up-regulating activation and cytolytic markers, as opposed to CD8⁺ T cells from tumors treated at a more advanced stage (Fig. 3c). The majority of granzyme B⁺ CD8⁺ tumor infiltrating lymphocytes (TILs) from day-7-treated mice co-expressed PD-1, as opposed to CD8⁺ TILs from day-4-treated tumors that could express granzyme B without up-regulating PD-1 (Extended Data Fig. 7d). This result, together with a previous study reporting the association between lack of T-cell exhaustion and tumor regressions upon DTA-1 treatment in the anti-GITR-responsive MC38 mouse colon adenocarcinoma model²⁷, pointed to a critical role for T-cell exhaustion in anti-GITR therapeutic activity.

Figure 3. — (a) B16 models of response and refractoriness to anti-GITR therapy. Differences in tumor growth in B16-bearing mice treated with a single administration of the anti-GITR antibody DTA-1 on day 4 (optimal curative condition, αGITR D4) or day 7 (suboptimal refractory condition, αGITR D7) after tumor implantation or the matched isotype IgG control. Data are mean ± SEM averaged shortest and longest tumor diameters of 8 mice/group from 1 of 3 independent experiments (homoscedastic two-sided multiple t test with Bonferroni-Sidak correction; day 24, p=0.00004). (b) Time course analysis of the indicated intra-tumor T-cell ratios and frequencies in the different treatment groups (black, isotype IgG control; red, αGITR D4; purple, αGITR D7). TILs were quantified by flow cytometry (FC) at baseline (Pre-Tx), and 3 and 6 days after optimal (Post-D4 Tx) or suboptimal (Post-D7 Tx) anti-GITR therapy or control treatment. Data are mean ± SEM of 4–10 independent experiments (CD4⁺Teff and Tregs: control day 4, n=4; control day 7, n=8; control day 10, n=10; control day 13, n=4; D4 αGITR day 7, n=6; D4 αGITR day 10, n=5; D7 αGITR day 10, n=7; D7 αGITR day 13, n=4; CD8⁺ T cells: control day 4, n=4; control day 7, n=8; control day 10, n=8; control day 13, n=4; D4 αGITR day 7, n=4; D4 αGITR day 10, n=5; D7 αGITR day 10, n=5; D7 αGITR day 13, n=4)(two-sided unpaired t test). (c) qPCR analysis of the indicated genes in CD8⁺ TILs isolated 6 days after αGITR D4, αGITR D7 or control IgG treatments from B16-bearing mice as indicated in the schema. Heatmap with unsupervised Ward hierarchical clustering of CD8⁺ TILs pooled from 5 mice/treatment and tested in triplicate in 1 of 2 independent experiments. * = p<0.05, ** = p<0.01, *** = p<0.001, **** = p<0.0001.

We thus reasoned that, if resistance to suboptimal day-7 anti-GITR treatment was due to CD8⁺ T-cell dysfunction, T-cell functional reinvigoration with PD-1 blockade should overcome this hurdle and make anti-GITR therapy effective against more advanced tumors. In agreement with this hypothesis, anti-PD-1 in combination with suboptimal anti-GITR (starting at day 7 after tumor implantation) controlled B16 growth as efficiently as optimal anti-GITR (administered 4 days after tumor implantation), achieving 50% complete tumor remissions associated with the development of long-lasting anti-tumor immunological memory (Fig. 4a). Importantly, the same combination was also able to regress tumors in mice bearing the poorly immunogenic and highly metastatic mammary carcinoma 4T1 (Fig. 4b). Mechanistically, we found that CD8⁺ TILs from mice treated with the combination on day 7 were phenotypically more similar to CD8⁺ TILs from mice treated with optimal day-4 anti-GITR than CD8⁺ TILs from mice treated with suboptimal day-7 anti-GITR or anti-PD-1, indicating T-cell reinvigoration/rejuvenation upon anti-GITR+anti-PD-1 (Fig. 4c,d). Specifically, the combination regimen counteracted CD8⁺ TIL up-regulation of Lag3, Pdcd1 or Tigit exhaustion-associated genes and promoted expression of a set of activation markers, including Klrg1 and Il2ra, in contrast to each monotherapy administered on day 7, yet similar to optimal day-4 anti-GITR (Fig. 4d). CD8⁺ TILs from day-7 anti-GITR+anti-PD-1-treated mice also overexpress the follicular helper T-cell markers Bcl6 and Icos (Fig. 4d), which have been recently found to define stem-cell-like memory CD8⁺ T cells in chronic viral infection^34,35. Overall, these phenotypic differences were coupled with increased clonality of day-7 CD8⁺ TILs from anti-GITR+anti-PD-1-treated mice (Extended Data Fig. 8), suggesting expansion of tumor-specific cytotoxic T cells. Accordingly, CD8⁺ TILs from anti-GITR+anti-PD-1-treated mice displayed a significantly enhanced tumor lytic capacity (Fig. 4c,e), even slightly superior to the cytotoxicity of CD8⁺ TILs from day-4 anti-GITR-treated mice (Fig. 4e), which was in agreement with a trend toward better survival upon treatment with anti-GITR+anti-PD-1 (Fig. 4a).

Figure 4. — (a) C57BL/6J mice were implanted with B16 cells and 7 days later treated with a single administration of anti-GITR monotherapy (suboptimal refractory condition, αGITR D7) or in combination with anti-PD-1 administered 4 times 3 days apart (αGITR+αPD-1 D7). In parallel, separated groups of mice were treated with anti-PD-1 monotherapy starting on day 7 (αPD-1 D7), optimal anti-GITR administered on day 4 (αGITR D4) or the matched isotype controls. Mean ± SEM averaged shortest and longest tumor diameters in the indicated number of mice per group from 1 of 2 independent experiments (left; 2-way ANOVA with Bonferroni correction), and overall survival (time to death or sacrifice) after the first tumor implantation (middle; log-rank test; two-sided p=0.004 and p<0.0001) and following a second tumor challenge in mice that rejected the first tumor (right) in 2 pooled experiments with the indicated number of mice per group. Number of tumor-free mice in each condition is reported. (b) Balb/c mice were implanted with 4T1 cells and 7 days later treated with αGITR, αPD-1, αGITR+αPD-1 or isotype controls and monitored for tumor growth and survival. Mean ± SEM volume (left; homoscedastic two-sided multiple t test with Bonferroni-Sidak correction; αPD-1 D7 *vs.* αGITR+αPD-1 D7, p=0.00007; αGITR D7 *vs.* αGITR+αPD-1 D7, p=0.00003) and overall survival (time to death or sacrifice; right) in the indicated number of mice per group. Number of tumor-free mice in each condition is reported. (c-e) CD8⁺ TILs were purified from B16 tumors 6 days after αGITR D4, αGITR D7, αPD-1 D7, αGITR+αPD-1 D7 or control treatments (c) and tested for expression of the indicated genes (d) and anti-tumor cytotoxicity in an *ex-vivo* 3D killing assay (e). (d) Heatmap with unsupervised Ward hierarchical clustering according to gene expression profiles of CD8⁺ TILs pooled from 5 mice/treatment and tested in triplicate in 1 of 2 independent experiments. Boxes highlight relevant genes similarly modulated in CD8⁺ TILs from mice treated with αGITR D4 and αGITR+αPD-1 D7. (e) Average ± SD percent of killed B16 cells upon co-culture with CD8⁺ TILs from animals treated as indicated in 1 of 2 independent experiments (n=3 except for D4 IgG where n=2; two-sided unpaired t test). * = p<0.05, ** = p<0.01, *** = p<0.001, **** = p<0.0001.

Taken as a whole, these results suggest that Teff exposure to Tregs in the tumor microenvironment contributes to the imprinting of T-cell exhaustion and demonstrate that Treg elimination alone is not sufficient to reinvigorate anti-tumor T-cell function in advanced tumors. Consistent with this concept, we found that maximal intra-tumor Treg downregulation upon GITR stimulation with TRX518, such as in patient 0002–0009 (Fig. 2b), was associated with persistent high frequency of PD-1-expressing CD8⁺ TILs, including GzmB⁺ effector CD8⁺ TILs (Extended Data Fig. 9).

An increase in intra-tumor Teff:Treg ratio is generally considered to be a surrogate marker of therapeutic anti-tumor responses³⁶. Our findings clearly indicate that in the setting of advanced tumors this concept is open to further iteration, as Treg removal from an established immunosuppressive tumor microenvironment cannot improve tumor control unless the T-cell exhaustion process is concurrently blocked (Extended Data Fig. 10). This points to the need for combining Treg-inhibiting/depleting immunotherapies with strategies able to counteract exhaustion, such as PD-1 blockade, to regress advanced tumors (Extended Data Fig. 10). PD-1 blockade monotherapy can also benefit from such an approach, as it has been reported that anti-PD-1 therapeutic activity is inversely correlated with tumor burden³⁷, indicating that the immunosuppressive tumor microenvironment may also limit the anti-tumor effects of T-cell reinvigoration with anti-PD-1. Our findings indicate that GITR agonism is a particularly suitable immunotherapy to combine with PD-1 blockade because, in addition to counteracting Tregs, it can also contribute to CD8⁺ T-cell re-activation together with anti-PD-1 in the setting of advanced tumors. Here, we find that the Fc-dysfunctional anti-GITR TRX518 preferentially induces loss of activated proliferating Tregs, thus contributing to the decrease in peripheral and intratumoral Tregs, in particular activated potentially pathogenic tumor-induced eTregs, in TRX518-treated patients. The possibility to exploit GITR functional modulation to selectively affect human Tregs without the need for engaging Fc-mediated mechanisms has the great advantage to avoid depletion of activated GITR⁺ CD8⁺ and CD4⁺ Teff that may be key to effective anti-tumor responses, specifically in the setting of PD-1 blockade. Based on our preclinical observations and TRX518 immune effects and safety profile in this first phase-I study, we have designed a clinical trial testing repeated administrations of TRX518 in combination with PD-1 pathway blockade in patients with advanced refractory solid tumors (NCT02628574). Three of the first patients enrolled in this study have demonstrated clinical responses (1 CR, 1 PR and 1 durable SD), including one patient that had progressed on prior anti-PD-1 therapy.

To our knowledge, this is the first time that GITR agonism is reported to decrease peripheral Tregs in cancer patients. These findings are supported by recent observations in humanized mice²⁹ where GITR agonism was found to affect human Tregs in peripheral blood, as opposed to mouse Tregs that are preferentially targeted in the tumor microenvironment by anti-GITR^21,26. This further strengthens the rationale and value of monitoring peripheral Tregs as a reliable surrogate biomarker of anti-GITR activity, to assess in combination with relevant peripheral biomarkers arising from studies with checkpoint blockade^37,38, in particular in patients treated with anti-GITR+anti-PD-(L)1.

Methods

Patient characteristics

43 patients with advanced solid malignancies requiring therapy were enrolled sequentially into 1 of 9 dosing cohorts (up to 7 patients/cohort) to receive a single intravenous dose of TRX518 ranging from 0.0001 mg/kg to 8.0 mg/kg (Supplementary Table 1). To be eligible, patients had to be ≥18 years old and have solid tumors which had relapsed or progressed following at least one prior systemic therapy for advanced disease. Patients were not permitted to have had immunomodulatory therapy for at least 42 days prior to study entry and could not have a history of autoimmune disorders or unresolved immune related adverse events following prior biologic therapy. The median age was 57 years (range 28–80) (Supplementary Table 1). More males than females (58.1% males versus 41.9% females) were enrolled in the study and the majority of patients were white (76.7%) (Supplementary Table 1). The study population was heterogenous with a variety of solid tumors represented, the most frequent being melanoma (n=10, 23.3%), lung cancer (n=9, 20.9%) and colorectal cancer (n=7, 16.3%) (Supplementary Table 2). Overall, enrolled patients had received a median of 3 prior therapies (range 0–9), including immunotherapy in 12 cases (27.9%).

Monitoring of adverse events (AEs) was conducted from the date of informed consent throughout the duration of study participation. Toxicities were graded according to the NCI CTCAE Scale (Version 4.0). Efficacy assessments were performed during screening and post-TRX518 (weeks 12 and 18) and evaluated using the Immune-related Response Criteria (irRC)³⁹. The majority (40 [93.0%] patients) experienced at least 1 treatment-emergent AE (TEAE) during the study. The most common TEAEs overall include fatigue (13 [30.2%] patients overall), cough (11 [25.6%] patients overall), nausea (9 [20.9%] patients overall), abdominal pain (8 [18.6%] patients overall), anorexia and vomiting (7 [16.3%] patients overall, each), and dyspnea (6 [14.0%] patients overall). A total of 15 (34.9%) patients experienced at least 1 TEAE of Grade ≥3, with the most common Grade ≥3 events being abdominal pain and neoplasm progression (3 [7.0%] patients each). Grade 4 events included 1 patient each with blood bilirubin increased and dyspnea. Three patients experienced events of neoplasm progression that resulted in death (Grade 5).

Overall, 16 (37.2%) patients experienced a treatment-related AE, the most common of which was fatigue (reported to be treatment-related in 5 [11.6%] patients overall). A total of 15 (34.9%) patients experienced one or more serious TEAEs during Part A of the study. Serious TEAEs that were reported in more than one patient overall included neoplasm progression (3 [7.0%] patients overall), abdominal pain (2 [4.7%] patients overall), and dyspnea (2 [4.7%] patients overall). There were no dose limiting toxicities or related serious adverse events. No safety trends in laboratory values, electrocardiograms, vital signs or physical examinations were identified.

PBMC samples were collected at baseline and at different time points up to 18 weeks after treatment. Flow cytometry analyses were performed on PBMC samples collected at baseline and up to 12 weeks after treatment from 37 patients, including 6 melanoma, 7 lung and 7 colorectal cancer patients and 17 patients with 11 other solid tumors (Extended Data Fig. 1a). Pre- and post-treatment biopsies from 3 melanoma, 2 colorectal, 2 lung and 1 bladder cancer patients enrolled in cohort 6–9 were available for immunofluorescent (IF) staining analysis (Extended Data Fig. 1a).

All procedures involved in this clinical trial were in compliance with the ethical regulations. All patients signed an approved informed consent before any study related procedures. Patient samples were collected according to the study protocol approved by IRBs at the participating sites MSKCC, Cleveland Clinic and University Hospitals.

In vitro assays with TRX518

MLR assays were performed by culturing donor-derived PBMCs (1×10⁶ cells/ml) with γ-irradiated (30 Gy) allogeneic PBMCs from a different healthy donor (1×10⁶ cells/ml) in the presence of IL-2 (100 IU/ml) and IL-15 (10 ng/mL). TRX518 or isotype control (10 μg/ml) was added to the cultures and the relative abundance of Foxp3⁺CD4⁺ Tregs and CD45RA⁻Foxp3^hiCD4⁺ eTregs was evaluated 7 days later by flow cytometry. Suppression assays were performed by incubating human Tregs, or Teff as control, immunomagnetically purified from donor PBMCs (human CD4⁺CD25⁺ Regulatory T Cell Isolation kit, Miltenyi Biotec) with CFSE-labeled CD8⁺ T cells immunomagnetically purified (human CD8 microbeads, Miltenyi Biotec) from the same donor (1:1 ratio) stimulated with plate-bound anti-CD3 (1 μg/ml) and TRX518 or isotype control (10 μg/ml) and soluble anti-CD28 (0.1 μg/ml) for 72 hours. CFSE dilution (proliferation) and CD25 expression (activation) in target CD8⁺ T cells, and Tbet and Foxp3 expression in CD4⁺ T cells (Tregs and Teff as control) from these co-cultures were then quantified by flow cytometry. Where indicated, Tregs were labeled with the CellTraceViolet dye (CTV, Invitrogen) to concurrently quantify cell death in proliferating and non-proliferating Tregs in the different treatment conditions by flow cytometry.

Caspase activation assays were performed by incubating CD8⁺, CD4⁺CD25⁻ Teff and CD4⁺CD25^hi Tregs immunomagnetically purified (human CD4⁺CD25⁺ Regulatory T Cell Isolation kit, Miltenyi Biotec) from healthy donor PBMC samples with plate-bound anti-CD3 (1 μg/ml), TRX518 or isotype control or recombinant human GITR ligand (GenScript) (10 μg/ml) and soluble anti-CD28 (0.1 μg/ml) in the presence of a fluorogenic caspase 3/7 substrate (1μM, Invitrogen) for 16 and 32 hours. Cell were then washed and stained with anti-human CD4, anti-human CD8 and 7AAD immediately before flow cytometry acquisition.

Mice

All mouse procedures were performed in accordance with institutional protocol guidelines at MSKCC. Wild type C57BL/6J and Balb/c mice were obtained from the Jackson Laboratory. DTR-Foxp3 transgenic mice were generously provided by Dr. Alexander Rudensky and backcrossed to C57BL/6J at MSKCC. Same sex, same age mice were used in each experiment.

Tumor cell lines

HEK293 NF-kB Reporter (Luc) cells were purchased from BPS Bioscience (San Diego, CA). 400,000 cells were transfected with 1μg of full length GITR in pcDNA3.1(+) using Lipofectamine 2000 (Life Technologies) and Optimem Serum Free Medium (Life Technologies) in 6 well plates. Cells were cloned twice by limiting dilution. The B16F10 mouse melanoma cell line (referred to as B16) was originally obtained from I. Fidler (M. D. Anderson Cancer Center, Houston, TX) and cultured in RPMI 1640 medium supplemented with 10% inactivated FBS, 1× nonessential amino acids and 2 mM l-glutamine. The 4T1 mammary carcinoma cell line was purchased from ATCC and cultured in RPMI 1640 medium supplemented with 10% inactivated FBS, 1× nonessential amino acids and 2 mM l-glutamine. Cell lines were routinely screened to avoid mycoplasma contamination and maintained in a humidified chamber with 5% CO2 at 37°C for up to 1 week after thawing before injection in mice.

In vivo mouse tumor injection and treatments

C57BL/6J were implanted with B16F10 melanoma cells intradermally (0.75×10^5 cells/mouse, for tumor-growth and survival analyses) or subcutaneously in Matrigel (Matrigel Matrix Growth Factor Reduced, Becton Dickinson) (2×10^5 cells/mouse, for immune-cell infiltrate analyses). Balb/c mice were implanted with 4T1 mammary carcinoma cells (100,000 cells/mouse) in the mammary fat pad. αGITR (clone DTA-1, BioXcell, 1mg/injection), or the matched isotype IgG (BioXcell), was administered intraperitoneally (i.p.) on day 4 (optimal curative treatment) or day 7 (suboptimal refractory treatment) after tumor implantation. Treatment with αPD-1 (clone RMP1–14, BioXcell, 250μg/injection) or the matched isotype IgG (BioXcell) was started 7 days after tumor implantation for 4 i.p. administrations 3 days apart. Tumor-free mice approximately 100 days after the first tumor implantation were inoculated intradermally with a double amount of B16 cells (1.5×10^5) in the opposite flank and monitored for tumor growth to test development of anti-tumor immunological memory. A single dose of diphtheria toxin (DT, 1μg/mouse) was administered i.p. on day 4 or day 7 after tumor implantation to induce short-term Treg depletion similar to the anti-GITR antibody DTA-1. Tumor growth was monitored by caliper twice a week. Survival was defined as time to death or time to sacrifice for those animals that had to be euthanized because sick and/or their tumors reached the size limits.

Flow cytometry analyses

B16 tumors were dissociated after 30 min incubation with Liberase TL and DNAse I (Roche) to obtain single-cell suspensions. When tumor mass exceeded 0.1 gr, immune-cell infiltrates were enriched by Percoll (GE Healthcare) gradient centrifugation. Surface staining of mouse cells was performed after Fcγ receptor blockade with an anti-mouse CD16/CD32 antibody (clone 2.4G2; BD Biosciences) for 15 min on ice by using panels of appropriately diluted fluorochrome-conjugated antibodies (from BD Biosciences, eBioscience or Invitrogen) against the following mouse proteins in different combinations: CD45 (clone 30-F11), CD4 (clone RM4–5), CD8a (clone 5H10 or 53–6.7), PD-1 (clone RMP1–30), GITR (clone DTA-1), CD25 (clone PC61.5) and an eFluor506 fixable viability dye. For intracellular staining, mouse cells were fixed and permeabilized (Foxp3 fixation/permeabilization buffer, eBioscience) and incubated with appropriately diluted FITC-labeled anti-mouse Foxp3 (clone FJK-16s, eBioscience) and PECF594-labeled anti-granzyme B (clone GB11, BD Biosciences) antibodies. For intracellular cytokine staining, mouse tumor immune infiltrates were re-stimulated with 500 ng/ml PMA and 1 μg/ml ionomycin in complete RPMI 1640 supplemented with 1 mM sodium pyruvate and 50 μM β-mercaptoethanol at 37°C. After 1 hour, 1× GolgiStop and 1× GolgiPlug (BD Biosciences) were added to the cultures and incubated for additional 4–5 hours at 37°C. Surface staining was performed after blocking Fcγ receptors by incubating cells with PE-Texas Red-labeled CD8, APCCy-labeled anti-CD45 and an eFluor506-labeled fixable viability dye. After 30 min incubation, cells were washed, fixed and permeabilized with the fixation/permeabilization buffer (eBioscience) and stained for 45 min with V450-labeled anti-IFNγ (clone XMG1.2, BD Biosciences) and APC-labeled anti-TNFα (clone MP6-XT22, BD Biosciences) antibodies.

Surface staining of human cells was performed in the presence of the FcγR Blocking Reagent (Miltenyi Biotec) with proper dilutions of fluorochrome-conjugated antibodies (from BD Biosciences, eBioscience or Tonbo) against the following human proteins in different combinations: CD45RA (clone H100), CCR7 (clone 150503), CD127 (clone HIL7RM21), CD3 (clone UCHT1), CD4 (clone S3.5), CD8 (clone RPA-T8), CD56 (clone CMSSB), GITR (clone eBioAITR), CD19 (clone HIB19), CD14 (clone M5E2), CD25 (M-A251) and a Near IR-labeled fixable viability dye. For intracellular staining, human cells were fixed and permeabilized (Foxp3 fixation/permeabilization buffer, eBioscience) and then incubated with proper dilutions of fluorochrome-conjugated antibodies against the following intracellular proteins: Foxp3 (clone PCH101, eBioscience or clone 259D/C7, BD Biosciences), ki67 (clone B56, BD Biosciences), and Tbet (clone eBio4B10, eBioscience) antibodies.

Analysis of competition between TRX518 and the FACS anti-human GITR eBIOAITR antibody was performed by incubating human GITR-transfected HEK293 with a saturating concentration of TRX518 (10 μg/ml) or a matched isotype control for 30 min on ice followed by staining with a PE-labeled anti-human GITR (eBioAITR) or the matched isotype IgG and an AlexaFluor647-conjugated anti-human IgG (Invitrogen) for 30’ on ice. DAPI was added to stained samples right before acquisition.

Samples were acquired on an LSRII or Fortessa flow cytometer (BD Biosciences) using BD FACSDiva software (BD Biosciences) and data analyzed with FlowJo 10.2 software (Tree Star Inc.).

FACS-sorting of human eTregs was performed on a FACSAria II cell sorter (BD Biosciences) upon PBMC incubation with the FcγR Blocking Reagent (Miltenyi Biotec) and staining with FITC-labeled anti-CD4, PE-labeled anti-CD45RA and APCCy-labeled anti-CD25 antibodies, and DAPI immediately before acquisition.

IF staining and analyses

IF staining was performed by the Molecular Cytology Core Facility at MSKCC using the Discovery XT processor (Ventana Medical Systems) as previously described⁴⁰. Briefly, tissue sections were deparaffinized with EZPrep buffer (Ventana Medical Systems), treated with CC1 buffer (Ventana Medical Systems) for antigen retrieval, and then blocked for 30 min with Background Buster solution (Innovex), followed by 8 min incubation with avidin-biotin blocking kit (Ventana Medical Systems). IF staining of Foxp3 and CD4 was performed by incubating blocked tissue sections with a mouse anti-human Foxp3 antibody (clone 236A/E7, 5 μg/ml, Abcam) for 4 hours, followed by 60 min incubation with biotinylated horse anti-mouse IgG (1:200, Vector labs). Signals were revealed by Streptavidin-HRP D (part of DABMap kit, Ventana Medical Systems) followed by incubation with AlexaFluor488 Tyramide (Invitrogen), according to the manufacturer’s instructions. Next, tissue sections were incubated with a rabbit anti-human CD4 antibody (clone SP35, 0.5 μg/ml, Ventana) for 5 hours, followed by 60 min incubation with a biotinylated goat anti-rabbit IgG (1:200, Vector Labs). Signals were revealed by Streptavidin-HRP D (part of DABMap kit, Ventana Medical Systems) followed by incubation with AlexaFluor546 Tyramide (Invitrogen), according to the manufacturer’s instructions. IF staining of PD-1, GzmB and CD8 was performed by incubating blocked tissue sections with a mouse anti-human PD-1 antibody (clone NAT105, 3.37 μg/ml, Ventana) for 4 hours, followed by 60 min incubation with biotinylated horse anti-mouse IgG (1:200, Vector labs). Signals were revealed by Streptavidin-HRP D (part of DABMap kit, Ventana Medical Systems) followed by incubation with AlexaFluor488 Tyramide (Invitrogen), according to the manufacturer’s instructions. Next, tissue sections were incubated with a rabbit anti-human GzmB polyclonal antibody (0.3 μg/ml, Ventana) for 5 hours, followed by 60 min incubation with a biotinylated goat anti-rabbit IgG (1:200, Vector Labs). Signals were revealed by Streptavidin-HRP D (part of DABMap kit, Ventana Medical Systems) followed by incubation with Tyramide Alexa568 (Invitrogen), according to the manufacturer’s instructions. Lastly, sections were incubated with rabbit anti-human CD8 antibody (clone SP57, 0.07 μg/ml, Ventana) for 5 hours, followed by 60 min incubation with biotinylated goat anti-rabbit IgG (1:200, Vector labs). Signals were revealed by Streptavidin-HRP D (part of DABMap kit, Ventana Medical Systems) followed by incubation with Tyramide Alexa647 (Invitrogen), according to the manufacturer’s instructions. Slides were counterstained with DAPI (Sigma Aldrich, 5 μg/ml) for 10 min and cover-slipped with Mowiol. Stained slides were digitally scanned using Pannoramic Flash 250 (3DHistech, Hungary) using 20×/0.8NA Zeiss objective and custom filters for the Alexa fluorophores. Relevant tissue regions were drawn on the scanned images as to retain tissue areas with immune infiltrates without artifacts or necrosis using CaseViewer software (3DHistech, Hungary), and the raw data from those regions were exported into TIFF files. TIFF images were analyzed using the FIJI/ImageJ software (NIH) by creating custom macro to count the positive cells. Specifically, nuclei were segmented using DAPI channel and positivity for each staining for each cell was assessed individually. Then double positive cells were tallied.

3D killing assay

A 3D collagen–fibrin gel culture system previously described⁴¹ was employed to measure cytotoxic activity of CD8⁺ TILs. Briefly, CD8⁺ T cells were immunomagnetically purified (CD8 microbeads, Miltenyi Biotec) from tumor immune infiltrates enriched on a Percoll gradient and co-embedded with B16F10 into collagen–fibrin gels (10:1 ratio, 1×10^5 CD8⁺ TILs: 0.1×10^5 B16 cells) and incubated for 24 hours. Gels were then lysed, and tumor cells were diluted and plated in 6-well plates for colony formation. After 7 days, plates were fixed with 3.7% formaldehyde and stained with 2% methylene blue before counting colonies as described⁴¹.

Real time quantitative PCR

Total RNA was extracted by using TRIZOL reagent (Invitrogen) and reverse-transcribed into cDNA using the High Capacity cDNA Transcription kit (Applied Biosystems). Expression of the indicated transcripts was quantified with the Fluidigm BiomarkTM system by using the appropriate FAM-MGB-conjugated TaqMan primer probes (Applied Biosystem) upon target gene pre-amplification according to the manufacturer’s protocol. Gene expression was normalized relative to glyceraldehyde-3-phosphate dehydrogenase (GΑPDH). Data were analyzed by applying the 2^(−dCt) calculation method. Heatmaps and unsupervised hierarchical clustering were generated in the R statistical environment (R development Core Team, 2008; ISBN 3–900051-07–0) (version 3.1.3) using hclust with Euclidean distance and Ward linkage.

Next-generation T-cell receptor (TCR) sequencing

150 ng of RNA from purified mouse CD8⁺ TILs were reverse-transcribed into cDNA using the High Capacity cDNA Transcription kit (Applied Biosystems) and used for survey level deep sequencing analysis of mouse TCR β-chain CDR3 regions by ImmunoSEQ (Adaptive Biotechnology). Data were analyzed by using the ImmunoSEQ analyzer toolset (Adaptive Biotechnology).

Statistical analyses

Two-sided Student’s t test and 2-way ANOVA (with Bonferroni’s multiple comparisons test) were used to detect statistically significant differences between groups. P values for survival analyses were calculated with log-rank (Mantel-Cox) test. Pearson correlation test was used to analyze dependency between variables. Statistical analyses were performed on the Prism 7.0a software (GraphPad Software) version for Macintosh Pro personal computer. Detailed information of the statistical test and number of observations/replicates used in each experiment are appropriately reported in each figure legend. Significance was defined as follows: * = p<0.05, ** = p<0.01, *** = p<0.001, **** = p<0.0001.

Reporting Summary

Further information on research design is available in the Nature Research Life Sciences Reporting Summary linked to this article.

Data availability

The TCR sequencing data that support the findings are available on the immuneACCESS Analyzer portal, https://clients.adaptivebiotech.com/pub/zappasodi-2019-natmed. All other relevant data are available upon reasonable request.

Extended Data

Extended Data Figure 3 — Changes in frequency of GITR⁺CD4⁺ T-cell subsets and their expression of cytokine receptors after TRX518. (a,b) Frequency of GITR⁺ Teff (blue) and GITR⁺ Tregs (red) (percentages of Teff or Tregs respectively) (top) and expression of IL7 receptor (CD127) and IL2 receptor alpha (CD25) on these cell subsets (bottom) in peripheral blood at baseline and after treatment with TRX518 at the indicated time points in cohort-5 (a) and cohort-7 (b) patients. CD127 and CD25 expression in GITR-negative Foxp3⁻CD4⁺ T cells (GITR^neg Teff, black) is also reported as control. Mean ± SD values of PBMC samples tested in duplicate at each time point. MFI, median fluorescence intensity.

Extended Data Figure 4 — Effects of TRX518 on Foxp3^low relative to Foxp3^hi GITR⁺ Tregs. (a) Representative gating strategy of Foxp3⁺ Tregs (Foxp3⁺CD4⁺), GITR⁺ Tregs (GITR⁺Foxp3⁺CD4⁺), Foxp3^lowGITR⁺ Tregs and Foxp3^hiGITR⁺ Tregs. Cell subsets were gated upon exclusion of doublets, CD19⁺, CD14⁺ and dead cells. (b) Fold changes in total Foxp3⁺GITR⁺ Tregs (as in Fig. 1b), Foxp3^low and Foxp3^hi GITR⁺ Tregs (percentage of Tregs) at the indicated time points after treatment with TRX518 relative to baseline. (c) Frequency of Foxp3^low and Foxp3^hi cells among GITR⁺ Tregs at baseline and after treatment with TRX518 at the indicated time points. (b-c) Mean ± SD values in one representative patient per dose cohort tested in duplicate.

Extended Data Figure 5 — Mechanisms leading to Treg loss upon treatment with TRX518. (a) Quantification of Foxp3⁺CD4⁺ Tregs, CD8⁺ T cells and CD4⁺Foxp3⁻ Teff in 7-day mixed leucocyte reaction cultures with PBMCs from 3 different donors treated with 10 μg/ml soluble TRX518 or isotype control. Each data point is average of 2–3 technical replicates; paired data are isotype- and TRX518-treated samples from the same donor (two-sided ratio paired t test; Tregs absolute #, p=0.0037; Tregs % of live cells, p=0.0364; Tregs % of CD4⁺, p=0.0200; n.s., not significant). (b-d) Standard proliferation/suppression assays with anti-CD3/anti-CD28-stimulated CFSE-labeled autologous CD8⁺ T cells cultured alone or with CD4⁺CD25^hi Tregs (1:1 ratio), or CD4⁺CD25⁻ Teff as control, immunomagnetically purified from donor-derived PBMCs, in the presence of 10 μg/ml plate-bound TRX518 or the isotype control. (b) Frequency of proliferating activated CFSE^low (top) and CD25⁺ (bottom) CD8⁺ T cells in each condition after 72-hour incubation in 1 of 2 independent experiments with different donors are shown (mean ± SD; CD8 alone, n=3; CD8:Teff, n=3; CD8:Tregs, n=2). (c) Proportion of CD4⁺ T cells (Teff or Tregs) expressing Foxp3 and/or Tbet (left), and proportion of dead Tregs expressing Foxp3 and/or Tbet (right) in the indicated T-cell co-cultures from 2 biologically independent donor-derived PBMC samples after 72-hour incubation (mean ± SD, n=3). (d) Frequency of dead cells in proliferating [CellTraceViolet(CTV)^low] and non-proliferating (CTV^hi) Teff and Tregs in the same cultures as in c. (e) Proportion of cells stained with a caspase 3/7 fluorogenic substrate (casp) and/or 7AAD viability dye in monocultures of immunomagnetically purified CD4⁺CD25^hi Tregs, CD8⁺ and CD4⁺CD25⁻ Teff activated with anti-CD3/-CD28 in the presence of 10 μg/ml plate-bound TRX518, isotype control (IgG), or a trimeric recombinant human GITRL (rhGITRL) after 16 and 32-hour incubation (mean ± SD, n=3 replicates). (f) Schematic representation of the sequence of events leading to Treg loss upon GITR stimulation. (g) Expression of the anti-apoptotic gene BCL2L1 (BCL-XL) in eTregs sorted from 2 melanoma patients at baseline and at the indicated time points after treatment (mean ± SD, n=3). FACS plot depicts the gating strategy to sort viable eTregs based on CD25 and CD45RA expression in CD4⁺ T cells. (b-e,g) Two-sided unpaired t test, * = p<0.05, ** = p<0.01, ***= p<0.001, **** = p<0.0001. n.a., not assessable.

Extended Data Figure 6 — Representative Treg modulations in peripheral blood and tumor upon treatment with TRX518. (a) Flow cytometry plots of Foxp3, CD25 and CD45RA expression in circulating CD14⁻CD19⁻dead⁻ CD3⁺CD4⁺ T cells from a melanoma (0102–0003, left) and a lung cancer patient (0002–0004, right) from cohort 7 (2 mg/kg TRX518) at baseline (Pre-Tx) and after treatment (Post-Tx) at the same time point as tumor biopsies were collected, as representative examples of patients showing Treg decreases or increases after TRX518 as shown in Fig. 2a. Gates showing frequencies of total Foxp3⁺ Tregs and Foxp3^hiCD45RA⁻ eTregs are depicted. (b) IF staining of Foxp3 (pink), CD4 (green) and DAPI (blue) on tumor tissue sections from the same patients and at the same time points shown in a, as representative examples of results shown in Fig. 2b for 8 patients (scale bar, 50 μm; 40× original magnification; inset, 60× original magnification). White arrows indicate Foxp3⁺CD4⁺ Tregs. PB, peripheral blood; TM, tumor.

Extended Data Figure 7 — GITR stimulation cannot overcome T-cell exhaustion in the setting of an immunosuppressive advanced tumor microenvironment. (a) B16-bearing mice were treated with a single administration of the anti-GITR antibody DTA-1 (αGITR) or the matched isotype IgG control 4 (D4) or 7 (D7) days after tumor injection and 6 days later flow cytometry analyses of tumor infiltrating lymphocytes (TILs) were performed. Data are mean ± SEM from 1 representative experiment out of those averaged in Fig. 3b (two-sided unpaired t test; αGITR, n=4; IgG, n=5). (b) B16-bearing Foxp3- diphtheria-toxin-receptor transgenic mice were treated with diphtheria toxin (DT) to deplete Tregs 4 or 7 days after tumor implantation as indicated in the schema. A separate group of mice was injected with PBS as control. Mean ± SEM averaged shortest and longest tumor diameters (n=5; 2-way ANOVA with Bonferroni correction; p<0.0001) and frequency of circulating Foxp3⁺ Tregs at the indicated time points after treatment (n=5; unpaired t test) from 1 representative of 2 independent experiments. (c) Frequencies, ratios and phenotype of CD8⁺ TILs and intra-tumor Foxp3⁺ Tregs (TM-Tregs) from B16-bearing untreated mice 4 (D4) or 7 (D7) days after tumor implantation. Data are mean ± SEM of 5 mice/group from 1 representative of 2–4 independent experiments depending on the parameter (two-sided unpaired t test). (d) Flow cytometry analysis of granzyme B (GzmB) and PD-1 expression in CD8⁺ TILs and PD-1 on GzmB⁺CD8⁺ TILs from B16 tumors harvested 6 days after optimal (αGITR 6 D4) or suboptimal (αGITR D7) anti-GITR therapy or control treatments (IgG). Data are mean ± SEM of 4 or 5 mice in αGITR- and IgG-treated groups respectively from 1 representative of 2 independent experiments (unpaired t test). * = p<0.05, ** = p<0.01, *** = p<0.001, **** = p<0.0001; n.s., not significant.

Extended Data Figure 8 — Modulation of intratumoral T-cell clonality. CD8⁺ TILs pooled from 5 mice/group 6 days after αGITR D4, αGITR D7, αPD-1 D7, αGITR+αPD-1 D7 or isotype control treatments as in Fig. 4c were processed for TCR β-chain CDR3 sequencing analysis (ImmunoSEQ, Adaptive Biotechnology). (a) Overview of TCR β-chain CDR3 clonality results and (b) frequencies of the indicated top rearrangements (productive frequency by amino acid sequences) in each treatment condition. Values for clonality range from 0 (polyclonal samples) to 1 (monoclonal or oligoclonal samples). Productive Clonality is calculated by normalizing Productive Entropy using the total number of unique Productive Rearrangements and subtracting the result from 1. Accordingly, the higher is the frequency of top rearrangements within a sample the higher is the clonality.

Extended Data Figure 9 — PD-1 expression on human CD8 TILs during TRX518 treatment. Representative IF staining of PD-1 (green), GzmB (red), CD8 (white) and DAPI (blue) on tumor tissue sections from patient 0002–0009 (cohort 8, colon cancer) before (Pre-Tx) and after (Post-Tx) TRX518 treatment (scale bar, 20 μm; 40× original magnification). White arrows indicate PD-1⁺GzmB⁺CD8⁺ T cells (top). Box and whiskers (min to max and median) plots showing frequencies of PD-1-expressing CD8⁺ TILs and PD-1-expressing GzmB⁺CD8⁺ TILs and absolute numbers of PD-1⁺CD8⁺ and PD-1⁺GzmB⁺CD8⁺ TILs normalized relative to tumor area (μm²) (Pre-Tx, n=10 optical fields; Post-Tx, n=3 optical fields) (bottom).

Extended Data Figure 10 — Model of effects of anti-GITR therapy in combination with PD-1 blockade in the setting of advanced tumors. Advanced tumors are generally characterized by abundant Treg infiltration and dysfunctional CD8⁺ T cells (1). Administration of anti-GITR in this setting can reduce intra-tumor Tregs promoting increases in CD8⁺ T-cell:Treg ratios (2–3). This however may be insufficient to cause effective anti-tumor T-cell responses because T cells remain dysfunctional (3). Changes in tumor-infiltrating T-cell quality is also key to ensure induction of anti-tumor immunity. This may be achieved by concurrent treatment with PD-1 blockade (4) that, leading to T-cell reinvigoration/rejuvenation (5), allows for optimal anti-tumor T-cell activation in the absence of Tregs (6).

Supplementary Material

Zappasodi et al _Tables

NIHMS1616665-supplement-Zappasodi_et_al__Tables.pdf^{(121.1KB, pdf)}

Acknowledgments

We thank the Immune Monitoring, Flow Cytometry and Molecular Cytology Core Facilities at MSKCC for technical assistance. This research was funded in part through the NIH/NCI Cancer Center Support Grant P30 CA008748, NIH/NCI R01 CA056821, the Swim Across America, Ludwig Institute for Cancer Research, Parker Institute for Cancer Immunotherapy and Breast Cancer Research Foundation. R.Z. was supported by the Parker Institute for Cancer Immunotherapy scholar award.

Competing Interests

R.Z. is inventor on patent applications related to work on GITR, PD-1 and CTLA-4. R.Z. is consultant for Leap Therapeutics. C.S. and W.N. are employed by Leap Therapeutics and report stock option ownership in Leap Therapeutics. P.W. is consultant for Leap Therapeutics. V.V. reports consultant/ advisory role for Genentech, Bristol Myers Squibb, Merck, AstraZenca, Celgene, Alkermes, Nektar Therapeutics, Reddy labs, Foundation Medicine, Taekeda Oncology, and institutional research grants from Genentech, Bristol Myers Squibb, Merck, Astrazenca, Leap therapeutics, Alkermes, Altor Biosciences. M.A.P. is consultant for Bristol Myers Squibb, Merck, Array BioPharma, Novartis, Incyte, NewLink Genetics, Aduro. M.A.P. reports grants from RGenix, Infinity, Bristol Myers Squibb, Merck, Array BioPharma, Novartis and honoraria from Bristol Myers Squibb and Merk. M.K.C. reports grants from and employment of a family member by Bristol-Myers Squibb; personal fees for advisory/consulting role from AstraZeneca/MedImmune, Incyte, Moderna and Merck. T.M. is consultant for Leap Therapeutics, Immunos Therapeutics and Pfizer, and co-founder of Imvaq therapeutics. T.M. has equity in Imvaq therapeutics. T.M. reports grants from Bristol-Myers Squibb, Surface Oncology, Kyn Therapeutics, Infinity Pharmaceuticals, Peregrine Pharmeceuticals, Adaptive Biotechnologies, Leap Therapeutics, Aprea. T.M. is inventor on patent applications related to work on oncolytic viral therapy, alphavirus-based vaccines, neo-antigen modeling, CD40, GITR, OX40, PD-1 and CTLA-4. J.D.W. is consultant for: Adaptive Biotech; Advaxis; Amgen; Apricity; Array BioPharma; Ascentage Pharma; Astellas; Bayer; Beigene; Bristol Myers Squibb; Celgene; Chugai; Elucida; Eli Lilly; F Star; Genentech; Imvaq; Janssen; Kleo Pharma; Linneaus; MedImmune; Merck Pharmaceuticals; Neon Therapuetics; Ono; Polaris Pharma; Polynoma; Psioxus; Puretech; Recepta; Trieza; Sellas Life Sciences; Serametrix; Surface Oncology; Syndax. J.D.W. reports grants from: Bristol Myers Squibb; Medimmune; Merck Pharmaceuticals; Genentech. J.D.W. has equity in: Potenza Therapeutics; Tizona Pharmaceuticals; Adaptive Biotechnologies; Elucida; Imvaq; Beigene; Trieza; Linneaus. J.D.W. is inventor on patent applications related to work on DNA vaccines in companion animals with cancer, assays for suppressive myeloid cells in blood, oncolytic viral therapy, alphavirus-based vaccines, neo-antigen modeling, CD40, GITR, OX40, PD-1 and CTLA-4. The other authors declare no competing interests.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Zappasodi et al _Tables

NIHMS1616665-supplement-Zappasodi_et_al__Tables.pdf^{(121.1KB, pdf)}

PERMALINK

Rational design of anti-GITR-based combination immunotherapy

Roberta Zappasodi

Cynthia Sirard

Yanyun Li

Sadna Budhu

Moshen Abu-Akeel

Cailian Liu

Xia Yang

Hong Zhong

Walter Newman

Jingjing Qi

Philip Wong

David Schaer

Henry Koon

Vamsidhar Velcheti

Michael A Postow

Margaret K Callahan

Jedd D Wolchok

Taha Merghoub

Figure 1. Circulating Tregs are preferentially affected by TRX518.

Figure 2. Correlation between peripheral and intratumoral Treg modulations upon TRX518.

Figure 3. Dysfunctional profile of CD8+ TILs after suboptimal anti-GITR therapy.

Figure 4. T-cell reinvigoration with PD-1 blockade potentiates activity of anti-GITR therapy against advanced tumors.

Methods

Patient characteristics

In vitro assays with TRX518

Mice

Tumor cell lines

In vivo mouse tumor injection and treatments

Flow cytometry analyses

IF staining and analyses

3D killing assay

Real time quantitative PCR

Next-generation T-cell receptor (TCR) sequencing

Statistical analyses

Reporting Summary

Data availability

Extended Data

Extended Data Figure 1.

Extended Data Figure 2.

Extended Data Figure 3.

Extended Data Figure 4.

Extended Data Figure 5.

Extended Data Figure 6.

Extended Data Figure 7.

Extended Data Figure 8.

Extended Data Figure 9.

Extended Data Figure 10.

Supplementary Material

Acknowledgments

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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Links to NCBI Databases

Figure 3. Dysfunctional profile of CD8⁺ TILs after suboptimal anti-GITR therapy.