Eligible patients had histopathologically verified localized PC and were surgical candidates for RP based on a standard workup of PSA measurement, biopsy results and imaging as needed. Patients must have chosen RP as their definitive treatment and needed to have evaluable biopsy tissue available for analysis or be willing to undergo a targeted prevaccination prostate biopsy. Patients were ≥18 years of age, had an ECOG performance status of 0–1 and had adequate organ function as defined by liver, kidney and hematological laboratory tests. Patients could have no evidence of immunocompromise or significant autoimmune disease that was active or potentially life-threatening if activated.

This phase II, single-arm study evaluated the impact of PROSTVAC vaccination in the neoadjuvant PC setting. Patients received recombinant vaccinia (rV)-PSA(L155)-TRICOM (2×10⁸ IU subcutaneously) as a priming vaccination, followed by monthly boosts on weeks 5, 9 and 13 with recombinant fowlpox (rF)-PSA(L155)-TRICOM (1×10⁹ IU subcutaneously). After completing the vaccination series, patients underwent RP. The boosting schedule was subsequently amended to weeks 3, 5 and 9 to allow for earlier surgeries. A final follow-up safety evaluation was performed 12–15 months postoperatively. AEs were monitored using Common Terminology Criteria for Adverse Events V.4.0. The primary objective was to examine changes in CD4 and CD8 T-cell infiltrates in tumor tissue postvaccination. Secondary objectives included investigating changes in peripheral PSA-specific T-cell responses, intraprostatic Treg infiltrates, PSA and MRI parameters postvaccination.

We stained 5 µm-thick formalin-fixed paraffin-embedded sections from prostate biopsies and RP specimens using Opal multiplex 6-plex kits, according to the manufacturer’s protocol (PerkinElmer), for a panel of DAPI, CD4, CD8, FOXP3, Ki67, PanCK and PD-L1 (online supplementary methods).

H&E and multiplex immunofluorescence scans were captured by a PerkinElmer Vectra Polaris. Both biopsies and RP tissue slide scans were virtually segmented by a research pathologist into three distinct compartments: CT, tumor IM and NL. Five randomly selected regions of interest (ROI) (mm², original magnification 40×, 0.25 µm/pixel resolution) from each compartment were then selected from each slide for multispectral imaging (MSI). Vectra Polaris captures spectral information from an MSI image using a multispectral camera, and the intensity of each fluorescent target is extracted from the multispectral data using linear unmixing. Unmixed MSI images were analyzed using inForm OS 2.3.1 software (PerkinElmer). A common algorithm was built based on five representative images, with adjustment from one batch to another due to batch variation. The algorithm consisted of cell segmentation, phenotyping and thresholding. All immune-cell infiltrates were measured as cell counts/mm² of tissue (density).

Antigen-specific responses were assessed by intracellular cytokine staining following a period of in vitro stimulation (IVS) of PBMCs with overlapping 15-mer peptide pools encoding the TAA PSA. The PSA peptide pool contained a previously identified agonist epitope9; pools encoding for human leukocyte antigen (HLA) and CEFT (a mixture of cytomegalovirus, Epstein-Barr virus, influenza and tetanus toxin) served as negative and positive controls, respectively. PBMCs from patients before and 60 days after therapy were stimulated by IVS and stained with antibodies to identify the absolute number of CD4+ or CD8+ lymphocytes producing cytokine (IFN-γ, TNF-α or IL-2) or positive for CD107a, as previously described.10 The background signal (obtained with the HLA peptide pool) and values obtained prevaccination were subtracted from those obtained postvaccination. Values >250 were scored as positive for TAA-specific immune response postvaccination if they were also at least twofold greater than that obtained with HLA. To explore evidence of cross-priming and development of an immune response to TAAs not found in the vaccine (antigen cascade or antigen spreading), responses to the cascade antigens MUC-1 and brachyury were tested using the same method. Peptide pools of MUC-1 and brachyury contained previously identified agonist epitopes.11 12

When feasible, an MRI was performed at baseline and after the last vaccination to assess for changes in the prostate and PC tumors prevaccination and postvaccination, including prostate volume, number of lesions, largest lesion length, PI-RADS score and ADC mapping.13 ADC values have been found to be decreased in various malignancies due to hypercellularity, and lower pretreatment ADC values in cervical cancer have been associated with a higher risk of recurrence.14

The primary objective was to determine whether there were changes in CD4 or CD8 cells overall, as well as within subsets based on compartments or peaks within ROIs. Ratios were formed to compare results post/pre. They were tested for whether the ratio=1 by the Wilcoxon signed rank test. Changes in averaged values over five randomly selected ROIs for the CD4 and CD8 T-cell infiltrates were considered the primary end point (p<0.025 would be significant following Bonferroni correction for these two tests, per protocol). Other measures were considered exploratory and were interpreted in the context of the number of tests performed, but without formal adjustment for multiple comparisons. All p values are reported as obtained, without adjustment for multiple comparisons. With 24 evaluable patients with paired data, the trial had 80% power to detect a change from baseline equal to 0.67 SD of the change using a 0.025 significance level paired t-test for each of the two primary measures. In practice, a Wilcoxon signed rank test was to be used if the ratios were not normally distributed. To allow for a small number of non-evaluable patients, the accrual ceiling for the trial was 27 patients.

Between October 2014 and May 2016, 27 patients were enrolled in a phase II study at the Center for Cancer Research, National Cancer Institute (table 1). The first four patients received monthly boosting vaccinations; all subsequent patients received boosts on weeks 3, 5 and 9. All patients completed the full series of vaccinations, 26/27 subsequently underwent radical prostatectomy (RP) and 1 patient elected to come off-study prior to surgery.

Matched prevaccination (biopsy) and postvaccination (RP) prostate samples from 26 patients were available for evaluation, all of which provided adequate tumor for analysis by two approaches: non-compartmentalized analysis (NCA) and compartmentalized analysis (CA) (see ‘Methods’ section). Two RP cases and two biopsies were missing benign lymph glands (NL), and one biopsy was missing infiltrative margins (IM) due to complete involvement by tumor. Tumor CD4 T-cell infiltrates were significantly increased in postvaccination RP specimens compared with baseline biopsies in NCA (median 176/mm² vs 152/mm²; IQR 136–317/mm² vs 69–284/mm²; p=0.0249; median ratio 1.21; IQR 0.64–2.25). Using the same analysis, CD8 T-cell infiltrates did not significantly increase postvaccination compared with baseline (median 147/mm² vs 107/mm²; IQR 111–226/mm² vs 94–203/mm²; p=0.23 (figure 1); median ratio 1.03; IQR 0.66–1.71 (figure 1, online supplementary figure 3)). The CA showed that both CD4 and CD8 T-cell infiltrates were seen more at the IM compared with the tumor core (CT) (figure 2). In this analysis, increased CD4 T-cell infiltrates at IM (median 198/mm² vs 151/mm²; IQR 123–500/mm² vs 85–256/mm²; p=0.042; median ratio 1.44; IQR 0.59–4.17) and CD8 T-cell infiltrates at CT (median 140/mm² vs 105/mm²; IQR 91–175/mm² vs 83–163/mm²; p=0.036; median ratio 1.25; IQR 0.88–2.09) were noted in postvaccination RP specimens compared with baseline biopsies (figure 2). Similarly, an increase in CD4 T-cell infiltrates was seen postvaccination in RP specimens at NL (median 263/mm² vs 86/mm²; IQR 93–531/mm² vs 41–225/mm²; p=0.0002; median ratio 2.33; IQR 1.05–5.22) compared with baseline biopsies (figure 2, online supplementary figure 3). In fact, 21/25 patients (84%) had a >1.5-fold increase in CD4 T cells and 18/25 patients (72%) had a >1.5-fold increase in CD8 T cells in at least one of three compartments (online supplementary table 3). T-cell infiltrates were increased in the tumor core (CT) in postvaccination RP specimens compared with untreated control RP specimens (n=5) for both CD4 (median 65.17/mm²; IQR 11.79–110.4/mm²; p=0.0132) and CD8 (median 18.3/mm²; IQR 14.29–53.61/mm²; p=0.0007) (online supplementary figure 4).

Intratumoral heterogeneity of various immune-cell infiltrate frequencies was more evident in RP specimens than in biopsies. Non-standard peak densities and a newly proposed heterogeneity score (peak to average immune score (PARIS), defined as peak density divided by average density, figure 3) were exploratory and used to assess the heterogeneity of immune infiltrates. The analysis of peak cell densities showed overall increases in CD4 and CD8 T-cell infiltrates in most compartments (p<0.05). This is particularly true for IM CD4 (median 533/mm² vs 174/mm²; IQR 287–1221/mm² vs 100–433/mm²; p=0.0001; median ratio 2.82; IQR 1.08–9.06), and NL CD4 (median 554/mm² vs 106/mm²; IQR 194–1431/mm² vs 67–276/mm²; p<0.0001; median ratio 4.65; IQR 1.99–9.85), as well as CT CD8 (median 281/mm² vs 159/mm²; IQR 186–429/mm² vs 98–243/mm²; p=0.001 (figure 3); median ratio 1.85; IQR 0.69–2.91 (figure 3)). PARIS score means for CD4 T cells postvaccination (2.26, 2.64, 2.43) were compared with prevaccination score means (2.21, 1.3, 1.25) in CT, IM and NL, respectively. PARIS score means for CD8 T cells postvaccination (2.11, 2.37, 2.10) were also compared with prevaccination score means (1.58, 1.33, 1.13) in CT, IM and NL, respectively. These scores were increased in all three compartments for both CD4 and CD8 T cells (p<0.05), except for CD4 T cells in CT (p=0.07) (figure 3).

In the NCA, an increase in effector (CD4+FOXP3–) T helper cell infiltrates (mean 307/mm² vs 194/mm²; 95% CI 184–430/mm² vs 117–271/mm²; p=0.03) was seen in postvaccination RP specimens compared with baseline biopsies (figure 4). Tumor-infiltrating T-regulatory cells (Tregs) (CD4+FOXP3+) tended to substantially decrease in postvaccination RP specimens compared with baseline biopsies (mean 5/mm² vs 17/mm²; 95% CI 4–7/mm² vs 5–29/mm²; p=0.06) (figure 4). Similarly, proliferative (Ki67+) effector cytotoxic CD8 T-cell infiltrates tended to have lower density in postvaccination RP specimens compared with baseline biopsies (mean 14/mm² vs 9/mm²; 95% CI 6–23/mm² vs 4–14/mm²; p=0.44). There was no change in CD8+Ki67– T-cell infiltrates (mean 165/mm² vs 156/mm²; 95% CI 121–209/mm² vs 90–222/mm²; p=0.56) in postvaccination versus baseline specimens.

Programmed death-ligand 1 (PD-L1) expression was focal (1%–10% on tumor cells) and only seen in 3/26 RP specimens (11.5 %) compared with 0/26 (0%) baseline specimens (online supplementary figure 5). Immune-cell PD-L1 expression was also focal and seemed to be present on macrophage-like cells. These cells were not stained with a dedicated marker or quantitated. On average, although variable by case and ROI, cases with PD-L1+ status had a higher post/pre ratio of immune-cell infiltrates (online supplementary figure 5).

Of 25 patients evaluable for PSA-specific CD4+ and CD8+ T-cell responses, 7 (28%) developed measurable PSA-specific increases in T-cell cytokine production and/or CD107a positivity postvaccination (online supplementary table 4). Further evaluation showed that seven (28%) and six (24%) patients, respectively, developed measurable responses to the cascade antigens MUC-1 and brachyury. A total of 13/25 patients (52%) developed responses to any of the three TAAs tested, and 5 of the 13 had responses to more than one evaluated antigen (figure 5). None of the four patients who experienced early biochemical recurrence (BCR) developed antigen-specific T-cell responses to any of the TAAs tested, compared with the 12/20 responses in the non-recurrence group (p=0.093 by Fisher’s exact test). Polyfunctional TAA-specific responses, defined as CD4+ or CD8+ T cells that express >2 of the markers interferon (IFN)-γ, tumor necrosis factor (TNF)-α, interleukin (IL)-2 or CD107a, were also measured before and after vaccination. Using the criterion of a >threefold increase postvaccination versus prevaccination, or the presence of >100 polyfunctional cells postvaccination per 1×10⁶ peripheral blood mononuclear cells (PBMCs) (if negative at prevaccination), polyfunctional T cells specific for at least one of the TAAs tested were generated in 28% of patients (online supplementary figure 6A). Representative flow plots from a patient (#22) developing multifunctional MUC1-specific CD4+ T cells after vaccination is shown in online supplementary figure 6B.