A Phase I/II Trial of HER2 Vaccine–Primed Autologous T-Cell Infusions in Patients with Treatment Refractory HER2–Overexpressing Breast Cancer

This article is featured in Selected Articles from This Issue, p. 3251

High levels of intratumoral type I T cells, in particular CD8 T cells and Th1, are associated with tumor control (1). This is particularly true for HER2⁺ breast cancer. In a study of more than 100 HER2-overexpressing breast cancers, the presence of Tbet⁺ lymphocytes in peritumoral lymphoid structures after neoadjuvant chemotherapy was independently associated with improved relapse-free survival (P = 0.04; ref. 2). Trastuzumab therapy induces type I HER2-specific T cells that can be measured in the peripheral blood and high levels of these T cells are independently associated with the development of a pathologic complete response (CR) after neoadjuvant chemotherapy (OR, 8.82; P = 0.016; ref 3). Conversely, loss or depression of HER2-specific Th1 cells in the blood is associated with disease recurrence (4).

Only 40% of patients will have HER2-specific Th1 cells induced after treatment with trastuzumab (5). Immunity can be generated or augmented with active immunization targeting HER2. Vaccination can effectively elicit HER2-specific immunity even in patients with advanced stage disease. In a study of 22 patients with stage IV HER2-overexpressing breast cancers, HER2-specific type I T cells could be significantly boosted and maintained with immunization (5). Our vaccines are designed to elicit CD4 T-cell responses for several reasons: (i) class II interacting epitopes can be identified that bind promiscuously to multiple HLA-DR allowing the vaccine to be responsive in a broad population (6–8); (ii) HER2 vaccines solely immunizing with epitopes designed to elicit CD8 T cells generate only short-lived immunity (9); and (iii) tumor antigen–specific CD4⁺ Th1 cells home to the tumor and the Th1 cytokines secreted, such as IFNγ, enhance the function of local antigen-presenting cells and augment endogenous antigen presentation (10). Increased processing of native tumor antigens results in epitope spreading and the development of an immune response to the multiple immunogenic proteins expressed in the tumor (11). Finally, by providing a robust CD4⁺ Th1 T-cell response, tumor-specific CD8⁺ T cells will be elicited and proliferate endogenously (12) and immunologic memory generated. We found that the frequencies of HER2-specific T cells generated from the peripheral blood of vaccinated patients can be expanded greater than 25-fold higher ex vivo than the number of HER2-specific T cells expanded from the peripheral blood of vaccine-naïve patients (13). We questioned whether we could use vaccine-primed T cells for therapeutic expansion in patients who had disease progression after vaccination.

We conducted a prior phase I study in eight patients who previously received a HER2 vaccine, experienced disease progression, and now had treatment-refractory metastatic disease (14). We found HER2-specific T cells could be adequately expanded for infusion in 7 of 8 patients. We showed the infused HER2-specific T cells could persist in 4 of 6 patients for more than 70 days after the first infusion and three patients experienced a partial response (PR). On the basis of these data, we developed a regimen of rapid weekly immunization followed by leukapheresis to collect cells for T-cell expansion and performed adoptive transfer of those cells for the treatment of advanced stage HER2 metastatic breast cancer. Data from this phase I/II trial are presented here.

After written informed consent, patients were enrolled (NCT00791037). The study was approved by Fred Hutchinson Cancer Research Center (Seattle, WA)/University of Washington Cancer Consortium Institutional Review Board, and was conducted in accordance with the International Council for Harmonisation (ICH) Guideline for Good Clinical Practice. Major eligibility criteria included HER2⁺ stage IV breast cancer that was maximally treated with persistent measurable disease > 10 mm by any imaging. No chemotherapy and/or HER2-targeted therapy could be administered for at least 7 days prior to receiving T-cell infusions. Twenty-three eligible patients were enrolled. Four patients withdrew from the study after signing informed consent prior to receiving T-cell infusions (Supplementary Fig. S1). Nineteen patients, all female, received at least two infusions. The representativeness of study participants is shown in Supplementary Table S1.

The study was a phase I/II nonrandomized, single-arm trial administering escalating doses of ex vivo–expanded HER2-specific T cells after in vivo priming with a multiple peptide–based vaccine directed against the HER2 intracellular domain (ICD) administered with GM-CSF as an adjuvant. The vaccine consisting of peptides p776–790 (GVGSPYVSRLLGICL, p776), p927–941 (PAREIPDLLEKGERL, p927), and p1166–1180 (TLERPKTLSPGKNGV, p1166), 500 μg each, was administered with 100 μg GM-CSF for a total of three immunizations via the same regimen as published previously (5). Two weeks after the third vaccine, the patients underwent leukapheresis to collect cells for T-cell expansion with the immunizing peptides. After the T-cell expansion, the patients were to receive three escalating cell doses with limits of 5.0 × 10⁹ (dose 1), 5.0 × 10¹⁰ (dose 2), and 5.0 × 10¹¹ (dose 3) total cells over 7- to 10-day intervals. Cyclophosphamide (600 mg/m²) was only administered 48 hours before the first T-cell infusion as a preconditioning regimen (14). One month after the final infusion, patients received three booster immunizations at 2-month intervals to induce expansion and persistence of the antigen-specific T cells in vivo. Patients could not be treated with chemotherapy during the study, but trastuzumab, endocrine, and/or bisphosphonate therapy was allowed during immunizations as we have shown that these agents do not impact the generation or maintenance of HER2-specific immunity after vaccination (15). All patients had previously progressed on these agents. Toxicity was assessed at each visit and was graded as per CTCAE v 3.0. Multigated acquisition scans were obtained before leukapheresis and after all T-cell infusions. Peripheral blood mononuclear cells (PBMC) for immunologic analyses were collected prior to enrollment, prior to the first T-cell infusion, 1 month after the last T-cell infusion, and prior to each postinfusion vaccine administration.

The primary objective was to evaluate safety. Our previous study suggested HER2 T-cell infusions were associated with minimal toxicity (14); therefore, with low observed rates of dose-limiting toxicity (DLT; 10% or less) in 20 patients, we would have 90% confidence the true DLT rate is less than 0.27. The secondary objectives included the extent and persistence of HER2-specific T-cell immune response, development of epitope spreading, and antitumor effects of T-cell infusions. Patients received a CT scan 1 month after the last T-cell infusion to assess clinical response. Target lesion responses were described as CR: disappearance of all target lesions; PR: at least 30% decrease in the sum of the longest diameter (LD) of target lesions; progressive disease (PD): at least a 20% increase in the sum of the LD of the target lesions, taking the smallest LD recorded prior to receiving the first T-cell infusion as a reference; and, stable disease (SD): neither sufficient shrinkage to qualify for PR nor sufficient growth to qualify for PD.

HER2-specific T cells were generated from the aphaeresis product derived from immunized patients using our previously published methods (13, 14). Briefly, PBMCs were purified from leukapheresis using Ficoll–Hypaque density gradient centrifugation, aliquoted, and cryopreserved in plasmalyte-A (Baxter) with 10% DMSO (Sigma-Aldrich) and 10% human serum albumin (CSL Behring LLC) and stored in liquid nitrogen. At the time of T-cell expansion, the PBMCs were thawed, washed, and stimulated with the HER2 ICD peptides [p776–790 (p776), p927–941 (p927), and p1166–1180 (p1166); 10 μg/mL each; Genemed Synthesis Inc.] on day 0. IL2 (10 U/mL, Chiron/Novartis Pharmaceuticals) and IL12 (10 ng/mL; R&D Systems) were added into the culture on days 4 and 8. On day 12, the cultured T cells were stimulated with Dynabeads ClinExVivoTM CD3/CD28 (Invitrogen) and cultured with IL2 (30 U/mL) every 2 to 3 days for 12 days until infusion.

A 3-day IFNγ enzyme-linked immunosorbent spot (ELISpot) assay was used to measure the immune response within the T-cell product as described previously (13, 14). Antigens used in the ELISpot assays included the three stimulating HER2 ICD peptides (p776, p927, and p1166) and three peptides indicative of intramolecular epitope spreading [HER2 p328–345 (TQRCEKCSKPCARVCYGL, p328), p688–703 (RRLLQETELVEPLTPS, p688), and p971–984 (ELVSEFSRMARDPQ, p971) all at 10 μg/each; ref. 16]. Data are presented as the mean of IFNγ spots/10⁶ T cells from antigen wells subtracting the spots from no-antigen wells (Corrected (c) IFNγ spots/10⁶). Estimated total HER2-specific IFNγ-secreting T cells infused were calculated using the combined HER2 cIFNγ spots/10⁶ multiplied by the total number of T cells infused.

To measure HER2 immunity in PBMCs, the cells were stimulated using HER2 antigens described above at different time points before and after infusion. Data are presented as a calculated 1/frequency of IFNγ-secreting cells to individual antigen in 10⁶ PBMCs. Data are also presented as cIFNγ spots/10⁶ PBMCs to HER2 peptides. Patients were considered to have augmented HER2-specific immunity postinfusion if the maximal cIFNγ response post T-cell infusion was greater than 2 standard deviations above the baseline level (P < 0.05; ref. 17).

Cryopreserved T-cell products were thawed and cultured in complete RPMI medium with IL2 (10 ng/mL, BioLegend) at a density of 1 × 10⁶/mL in a 37°C, 5% CO₂ incubator. After overnight recovery, viable T cells were enriched using Ficoll-Paque Plus (GE Healthcare). CD4⁺ and CD8⁺ T-cell subsets were separated using anti-CD4 or anti-CD8 microbeads (Miltenyi Biotec) and resuspended in fresh complete RPMI medium at a density of 1 × 10⁶/mL. Approximately 100 μL of T-cell suspension was seeded into a well of 96-well flat-bottom plate precoated with anti-human CD3 (clone OKT3, 10 μg/mL in PBS at 4°C) with a supplement of soluble anti-human CD28 (clone CD28.2) at a final concentration of 5 μg/mL. After culture at 37°C, 5% CO₂ for 24 hours, the cells were stained with PE-conjugated anti-CD4 or Alexa Fluor 647–conjugated anti-CD8 antibody at room temperature for 10 minutes, rinsed once with PBS, and resuspended in complete RPMI medium at a density of 1 × 10⁶/mL. Approximately 30 μL of cell suspension was loaded into the 32-plex antibody loaded IsoCode Chip (IsoPlexis) and incubated at 37°C, 5% CO₂ for additional 16 hours. Protein secretion from approximately 1,000 T cells was then captured. The polyfunctional T cells that cosecreted 2+ cytokines per cell were analyzed by the IsoSpeak software across five functional groups: (i) effector: Granzyme B, TNFα, IFNγ, MIP-1α, Perforin, TNFβ; (ii) stimulatory: GM-CSF, IL2, IL5, IL7, IL8, IL9, IL12, IL15, IL21; (iii) chemoattractive: CCL11, IP-10, MIP-1β, RANTES; (iv) regulatory: IL4, IL10, IL13, IL22, sCD137, sCD40L, TGFβ1; and (v) inflammatory: IL6, IL17A, IL17F, MCP-1, MCP-4, IL1β. The polyfunctional strength Index (PSI) of T-cell products was computed using a prespecified formula, defined as the percentage of polyfunctional cells, multiplied by mean fluorescence intensity (MFI) of the proteins secreted by those cells (18–21).

Cryopreserved PBMCs from pretreatment and 1-month after the last T-cell infusion were thawed and washed according to previously published methods (22). The washed cells (10⁶) were incubated with 10% normal mouse serum in PBS at room temperature for 30 minutes to block nonspecific binding. After washing, the cells were stained with fluorochrome-conjugated mAbs, including CD3 FITC (clone HIT3a), CD4 APC (RPA-T4), CD8 PE-Cy7 (HIT8a), CD45RA PerCP/Cy5.5 (HI100), CCR7 (CD197) PE (G043H7), CD69 APC Cy7 (FN50), PD-1 APC (EH12.2H7), and HLA-DR PE-Cy7 (L243) for T-cell phenotype analysis. T-regulatory cell and myeloid-derived suppressor cell (MDSC) analyses were performed using our previously published methods (14, 23). All antibodies were purchased from BioLegend. Data acquisition was performed on a FACSCanto flow cytometer (BD Biosciences) and analyzed using a FlowJo software (RRID:SCR_008520). Data are expressed as the percentage of CD45RA⁻ CCR7⁺ central memory, CD45RA⁻ CCR7⁻ effector memory, CD45RA⁺ naïve, CD69⁺, PD-1⁺, or HLA-DR⁺ cells among CD3⁺ CD4⁺ and CD3⁺ CD8⁺ cells.

Statistical analysis was performed using GraphPad Prism V8 (RRID:SCR_002798). The differences of the magnitude of immune responses in the different time course of treatment were determined by two-way ANOVA with a Dunnett multiple comparison test. The differences of parameters between two groups of patients were analyzed using a two-tailed Student t test if data were normally distributed, and Mann–Whitney test if data were skewed. The differences of parameters between pre- and posttreatment in patients were analyzed using paired t test when data were normally distributed, and Wilcoxon matched-pairs test if data were skewed. Kaplan–Meier curves were generated to show the probability of overall survival (OS), which was defined as the time elapsed between the enrollment and death. Median survival time was defined as the time when the survival probability dropped to 50%. Furthermore, to explore the difference between survival curves of responders (PR+SD) compared with nonresponders (PD) to the study treatment, log-rank (Mantel–Cox) test was conducted. Statistical significance was defined as P < 0.05.

The datasets used and/or analyzed during this study are available from the corresponding author on reasonable request.

Expansion of HER2 vaccine–primed T cells after weekly vaccination in treatment-refractory patients is feasible and infusions are associated with limited toxicity. The patient characteristics are shown in Table 1. Over half of the patients had four or more previous lines of therapy and two or more sites of metastatic disease. Fourteen patients received all three planned T-cell infusions and five patients received two infusions. Three of the five patients did not have enough lymphocytes in the PBMCs derived from the leukapheresis to generate three expansions, the product from one patient was contaminated with bacteria and was not infused, and the final patient received an infusion of trastuzumab within the 7 days of the planned infusion. The median fold increase in CD3⁺ cells among the 19 subjects was 6.4 (range 0.1–17.4) with some level of expansion occurring in all patients but one (Fig. 1A). The median number of T cells infused was 2,731 × 10⁶ (range 14–5,000) in dose 1, 7,420 (56–33640) in dose 2, and 13,678 (272–14534) in dose 3. The median total number of T cells infused was 17,275 × 10⁶ (range 343–57,120 × 10⁶) with a median of CD3⁺ 99% (range 90%–100%), CD4⁺ 44% (range 3%–82%), and CD8⁺ T cells 48% (range 17%–96%). The median HER2-specific T cells infused, defined as the sum of T cells responding to both stimulating peptides and epitope spreading–associated peptides assessed in an ELISpot using material from each product prior to infusion, among the 19 patients was 36.3 million (range 0.2–417 × 10⁶), among SD+PR was 60 (0.2–262), and PD was 11 (1–416; Fig. 1B). Thirteen percent of expanded T-cell products were polyfunctional; 8% cosecreted 2+ proteins, 3% 3+ proteins, 1% 4+ proteins, and 1% 5+ proteins (Fig. 1C). If polyfunctionality was calculated from cytokine-producing single cells, 43% of these were polyfunctional. The enhanced PSI of the T-cell products was comprised primarily of effector and stimulatory functions with granzyme B, IFNγ, and TNFα secretion occurring in >5% (range 7%–12%) of the cells (Fig. 1D). The chemoattractive and regulatory groups were minimally represented with the exception of sCD137 secretion that occurred in 13% of cells.

A total of 556 adverse events (AE) were collected during the trial (Table 2; Supplementary Table S2). Of all AEs, 96% were grades 1 or 2. There were 3% grade 3 AEs, 10 related to changes in hematologic indices that resolved spontaneously, two related to gastrointestinal symptoms induced by cyclophosphamide, and one headache. There was a 1% grade 4 AE rate all related to hematologic indices (lymphocyte and absolute neutrophil count), which resolved spontaneously. The most common related AE was injection-site reaction after vaccination (13%) or those related to cyclophosphamide administration (Table 2; Supplementary Table S2).

The majority of patients had increased HER2-specific T cells in the peripheral blood after infusion, which is reflective of both immunizing and intramolecular spreading epitopes. One month after the last T-cell infusion, the majority of patients had augmented immunity to p776 (56%), p927 (65%), or p1166 (59%), respectively (Fig. 2A). The median antigen-specific T-cell response to HER2 p776, p927, and p1166 at study entry was a frequency of less than one antigen-specific cell in 53,865 (range 1:2,183–1:1,000,000), 600,000 (1:918–1:1,000,000), and 1,000,000 (1:1,550–1,000,000) PBMCs, respectively. The maximal response postinfusions was a median frequency of one in 32,955 (1:478–1:1,000,000), 6,630 (1:1,033–1:300,000), and 11,215 (1:837 to 1:1,000,000) PBMCs, respectively (Fig. 2B). Overall, 82% (14/17) of patients significantly augmented the immune response to at least one of the immunizing epitopes (all post vs. pre T-cell infusion, P < 0.05).

The median peptide-specific T-cell response to the three peptides, which represented intramolecular epitope spreading within HER2, p328, p688, and p971, before study enrollment was a frequency of less than one antigen-specific cell in 171,429 (range 1:1,299–1:1,000,000), 600,000 (1:3,040–1:1,000,000), and 44,690 (1:1,856–1:1,000,000) PBMCs, respectively. The maximal response postinfusion was a median frequency of one in 25,532 (1:638–1:1,000,000), 9,534 (1:1,133–1:1,000,000), and 25,307 (1:1,309–1,000,000), respectively (Fig. 2B). Patients had augmented immunity to p328 (64%), p688 (50%), or p971 (56%; Fig. 2A). Overall, 81% (13/16) of patients demonstrated enhanced intramolecular epitope spreading (all post vs. pre T-cell infusion, P < 0.05).

Of the 19 treated patients, 17 were evaluable for assessment of clinical response. Two of the patients did not receive a CT scan 4 weeks after the last T-cell infusion. No patient had a CR. One patient had a PR (6%) that was durable for 8 months after the end of infusions and we could document infused T cells homing to numerous sites of metastatic disease in this patient (24). This patient received trastuzumab during vaccination. Eight patients experienced SD (47%) for a disease control rate of 53%. Of these eight patients, five received trastuzumab and three received both trastuzumab and endocrine therapy during the vaccination phase of the study. Of note, although patients received trastuzumab and/or endocrine therapy during the vaccination phase, each individual had progressed disease on the therapy prior to enrollment in this study. The average time to progression in these patients was 6 months (range 2–18 months) from the time of the first infusion. Eight patients had PD (47%). Five of these patients received trastuzumab during the vaccination stage of the study. The median OS for 19 patients was 28.6 months, and for 17 patients evaluable for assessment of clinical response was 35.7 months. In an exploratory analysis, we show that the median survival for those with PD was 20.5 months, and for those with either PR or SD was 45.0 months (P = 0.008; Fig. 3A; Supplementary Fig. S2). There was no significant difference in survival between two infusions and three infusions (P = 0.88).

The responding patients (PR+SD) significantly generated or enhanced the IFNγ response to the three vaccinating epitopes 1-month postinfusion as compared with the prevaccine baseline (mean ± SE of cSPOT/10⁶ PBMCs: pre, 134 ± 71; 1-month postinfusion, 927 ± 529; P = 0.012), but the patients with PD did not show a significant increase at that time point (P = 0.26; Fig. 3B). Enhanced HER2-specific immunity was maintained at higher levels in responding patients at 3 months (mean ± SE, 591 ± 421) and 5 months post T-cell infusion (541 ± 411) compared with those with PD (Fig. 3B). Responding patients received a median of three booster vaccines after T-cell infusion, and patients with PD received a median of two postinfusion vaccines.

In an exploratory analysis, we evaluated immune parameters at the time of enrollment to identify candidate biomarkers that may predict patients most likely to respond to therapy. The level of CD4⁺ CD69⁺ T cells prior to treatment for the responding patients was a median of 6.6% (range 1.4%–13.5%) of all CD4 T cells as compared with a median of 2.5% (range 0.5%–5.4%) in nonresponders (P = 0.059; Fig. 4A). The same association was observed with CD8⁺ CD69⁺ T cells that we measured at a median of 9.2% (range 4.0%–14.2%) of all CD8 T cells in responders, but were found at a median of 3.7% (range 3.0%–5.0%) in nonresponders (P = 0.018; Fig. 4B). In addition, central memory (CD45RA⁻ CCR7⁺) CD4⁺ T cells were significantly increased at baseline in responding patients: 70.0% (range 43.6%–87.0%) of CD4 T cells as compared with nonresponders, 40.3% (range 24.6%–50.7%; P = 0.004; Fig. 4C). There was no association of the pretreatment levels of central memory CD8⁺ T cells, PD-1⁺ CD4 or CD8 T cells, T-regulatory cells, or MDSCs with clinical outcome (all P > 0.05; Supplementary Table S3).

We also assessed whether there were changes in immune cell phenotypes that occurred with treatment that were associated with clinical outcome. The percentages of effector memory (EM) T cells, CD45RA⁻, CCR7⁻, CD4, and CD8, significantly increased in the responding patients (Figs. 4D and E). In responding patients, EM constituted a median of 17.5% (range 5.6%–30.4%) of CD4 T cells at baseline and were significantly increased 1 month after the end of T-cell infusion (median 23.3%; range 13.5%–42.2%; P = 0.031; Fig. 4D). No such increase was observed in CD4 EM in patients with PD (P = 0.313). Similarly, levels of CD8 EM were also increased in responding patients (Fig. 4E). EM constituted a median of 23.7% (range 8.4%–63.0%) of CD8 T cells at baseline and were significantly increased 1 month after the end of T-cell infusion (median 31.5%; range 11.0%–67.8%; P = 0.016; Fig. 4D). Increases were not detected in patients with PD (P = 0.125). Although levels of PD-1⁺, CD4, and CD8 T cells were similar among all patients at baseline, those who had a response to therapy reduced the number of PD-1⁺ CD8 T-cells in the peripheral blood as compared with nonresponding patients (Fig. 4F). In responders, PD-1⁺ CD8⁺ T-cells were detected at a median level of 17.4% (range 7.6%–77.9%) at baseline and decreased to a level of 14.3% (range 2.1%–70.4%) after therapy (P = 0.016). There was no such change in nonresponders. There were no significant associations with clinical outcome for changes in PD-1⁺ CD4, CD69⁺ T cells, T-regulatory cells, or MDSCs (all P > 0.05; Supplementary Table S4).

Antigen specific vaccination can be used as a priming tool to increase and activate T cells in vivo for optimal ex vivo expansion (25). We have previously reported a T-cell expansion method utilizing IL2 and IL12 that favors proliferation of type I CD4⁺ T cells (26). We applied this T-cell culture method along with HER2-specific vaccination to rapidly generate HER2-specific autologous T cells for therapeutic infusion. Data presented here demonstrate that it is possible to utilize vaccination as means to prime T cells for ex vivo expansion in patients with advanced stage treatment-refractory metastatic breast cancer, that epitope spreading can be elicited and augmented, and that infused antigen-specific T cells can persist in vivo.

Vaccine primed T cells have been used with some success in the treatment of hematologic malignancies. A recent phase I/II trial in mantle cell lymphoma–treated patients with T cells expanded in culture after vaccination with irradiated CpG-activated tumor cells and demonstrated promising clinical outcomes (27). T cells were infused after remission was induced with immunochemotherapy. Vaccination generated tumor-specific memory CD8 T cells in 40% of patients and the presence of these cells was associated with a longer time to progression. Application of this approach to the treatment of patients with advanced solid tumors has been less well studied, in part, due to availability of vaccines particularly for tissue types where tumor cells are difficult to obtain. In one trial, patients with advanced ovarian cancer were treated with dendritic cells (DC) pulsed with autologous tumor lysates as the priming immunization and PBMCs were collected (28). After T-cell expansion on tumor lysates in vitro, patients underwent lymphodepletion with cyclophosphamide and subsequent T-cell infusion. Of six patients treated, two demonstrated PR and two experienced disease stabilization (28). An earlier trial in advanced renal cell carcinoma (RCC) also utilized autologous tumor cells for vaccination (29). In this phase II study, after immunization, lymph nodes draining the vaccine site were removed and used as a source of T cells. Cells derived from the lymph nodes were nonspecifically expanded with an anti-CD3 mAb and IL2 treatment then infused. Thirty-four patients were treated with nine responses; four CR, and five PR for an overall response rate (ORR) of 27%. Notably, although all patients were heavily pretreated with multiple lines of chemotherapy, most patients could be effectively immunized and generate detectable and even high levels of tumor-reactive T cells after immunization. The HER2-specific vaccine used in this study is immunogenic in patients with advanced solid tumors and allows extension of this approach to tumors such as breast cancer, where the collection of autologous tumor cells for vaccination is not possible in many cases (16).

A unique aspect of this study is the generation or augmentation of intramolecular epitope spreading in the majority of treated patients. Epitope spreading, which is a broadening of the immune response to other antigens in the tumor, represents activation of the tumor immune environment to support enhanced cross-priming (30). Infusion of high number of IFNγ-secreting HER2-specific CD4 T cells could have contributed to the significant incidence of epitope spreading during this study. We were able to track infused T cells in one patient using concurrent single-photon emission CT (SPECT)/PET-CT imaging (24). Twenty-four hours after infusion, adoptively transferred T cells could be detected at all sites of metastatic disease. Furthermore, concurrent PET-CT imaging demonstrated a fluorodeoxyglucose flare at most sites of disease at 48 hours. These data suggest HER2-specific Th1 can home to tumor sites and potentially activate antigen-presenting cells at various locations to increase tumor inflammation and cross-priming facilitating the development of immunity to other tumor-associated antigens. The generation of epitope spreading during treatment with a cancer immunotherapy has been associated with favorable clinical outcomes (30). Preclinical studies have shown that while adoptively transferred T cells or chimeric antigen receptor (CAR) T cells can elicit rapid antitumor responses, the endogenous immunity generated, as indicated by epitope spreading, prevents the development of antigen-loss variants and subsequent tumor immune escape (31, 32).

Continued vaccination was utilized after adoptive transfer to enhance the persistence of T cells in vivo. Patients with a PR or SD were able to maintain high levels of HER2-specific T cells in the peripheral blood over time in contrast to those with PD. It is unknown whether continued vaccination after adoptive transfer is needed for HER2 T-cell persistence. In our previous phase I study of this approach, HER2-specific T cells persisted in the majority of patients for more than 70 days from first infusion without postinfusion immunizations (14). Our data suggest that evidence of activated T cells and elevated CD4 central memory T cells at the time of enrollment, identifies those patients more likely to have a response to treatment. CD69⁺, CD4, and CD8 T cells at higher levels at baseline could reflect a more immunogenic tumor and T cells that are not functionally exhausted. Higher levels of central memory T cells prior to vaccination could provide a T-cell pool that can rapidly proliferate when activated and become functional effector cells (33). As pharmacodynamic markers, an increase in both CD4 and CD8 EM T cells and a decrease in PD-1 expression on CD8 T cells during therapy was associated with beneficial outcomes. Recent investigations have suggested that clonal expansion of CD8 T-cell effector memory populations are associated with durable clinical responses after treatment with immune checkpoint inhibitors in melanoma (34). Several studies of various forms of immune therapy have reported that higher levels of PD-1 expression on CD8 T cells is associated with suboptimal response. A clinical trial of ipilimumab in advanced melanoma associated higher levels of CD8⁺ PD-1⁺ T cells with decreased OS (35). Similarly, in a study of imiquimod and chemotherapy in patients with breast cancer with refractory chest wall disease, PD-1 expression on either CD4 or CD8 T cells was associated with lack of response to treatment (36). Future trials will be needed to further validate these biomarkers.

There are limitations to the study. First, as the patients were heavily pretreated, we were only able to obtain limited samples that impacted the ability to fully phenotype peripheral blood immune cells and address intermolecular epitope spreading. In addition, our T-cell cultures for infusion were associated with limited polyfunctionality. Our median PSI score ∼125 is 4-fold lower than scores that have been associated with beneficial clinical outcomes after adoptive T-cell transfer (18). The PSI score may vary depending on antigen specificities of T-cell products, stimulation methods to promote cytokine secretions, and disease indications. Modifying our ex vivo T-cell culture conditions to include cytokines such as IL7 or those associated with the generation of polyfunctional T cells could improve the antitumor activity of the infused product (37, 38). Of note, polyfunctionality represents the cells that cosecreted at least two cytokines per cell in the analyzed single cells that included both cytokine-producing and nonproducing individual cells. It has been reported that these defined polyfunctional cells secreted not only the most numbers of different cytokines, but also the highest copy numbers of any individual cytokine (20). Although these polyfunctional T-cells comprised only 10%–20% of the total population, they functionally dominated the population of infused T cells. Finally, although disease stabilization lasted a median of 6 months, the median OS in the responders was 45 months. We were not able to assess clinical responses to subsequent treatment after patients ended the study. High levels of type I T cells could potentially sensitize tumors to become more responsive to subsequent chemotherapy treatment via SOCS1 inhibition of oncogenic signaling pathways as a potential mechanism for the prolonged survival of responders (39).

In conclusion, adoptive transfer of HER2 vaccine–primed T cells was feasible and moderately effective in patients with treatment-refractory HER2-positive breast cancer. Future studies will evaluate the combination of T-cell transfer with an anti–PD-1 mAb that may enhance efficacy of the approach by increasing T-cell polyfunctionality and allowing further expansion of HER2-specific T cells in vivo (40).

M.L. Disis reports nonfinancial support from NIH and other support from Helen B Slonaker Endowed Professor for Cancer Research, Epithany, and University of Washington during the conduct of the study as well as grants from NCI, Gateway for Cancer Research, Komen Foundation, American Cancer Society (ACS), Aston Sci, Precigen, Bavarian Nordisk, and Veanna outside the submitted work; in addition, M.L. Disis has a patent for University of Washington licensed. A.L. Coveler reports grants from Seagen, AbGenomics, Novocure, AstraZeneca, Actuate, Amgen, and Mirati and grants and nonfinancial support from Nucana outside the submitted work. J.S. Childs reports grants from NCI during the conduct of the study. J. Zhou reports other support from IsoPlexis Corporation outside the submitted work. S. Mackay reports personal fees from IsoPlexis during the conduct of the study as well as personal fees from IsoPlexis outside the submitted work. L.G. Salazar reports grants from National Cancer Institute during the conduct of the study. No disclosures were reported by the other authors.

M.L. Disis: Conceptualization, resources, data curation, formal analysis, supervision, funding acquisition, investigation, methodology, writing–original draft, project administration, writing–review, and editing. Y. Dang: Data curation, formal analysis, investigation, methodology, writing–original draft, writing–review, and editing. A.L. Coveler: Data curation, investigation, and methodology. J.S. Childs: Data curation, investigation, and methodology. D.M. Higgins: Data curation, investigation, and methodology. Y. Liu: Data curation, investigation, and methodology. J. Zhou: Data curation, formal analysis, investigation, methodology, writing–original draft, writing–review, and editing. S. Mackay: Data curation, formal analysis, investigation, methodology, writing–original draft, writing–review, and editing. L.G. Salazar: Conceptualization, data curation, investigation, methodology, writing–review, and editing.

This work was supported by NCI R01CA129517, R01CA85374, and the Gateway for Cancer Research. Patients were seen in the clinical research center supported by UL1 TR002319. L.G. Salazar was supported by K23CA10069. Data analysis by Y. Dang was supported by the Komen Foundation (SAC130058). M.L. Disis was supported by the Helen B. Slonaker Endowed Professor for Cancer Research and an ACS Clinical Professorship (CRP-15–106–01-LIB).

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