Telomerase-based GX301 cancer vaccine in patients with metastatic castration-resistant prostate cancer: a randomized phase II trial

This was a multicentre, randomized, parallel-group, open-label trial with blind assessment of the primary end-point. The study was carried out in compliance with the Helsinki Declaration. The protocol was approved by national competent authorities (AIFA and AEMPS, respectively) and the ethics committees of all participating hospitals and was registered with EudraCT (2014-000095-26) and ClinicalTrials.gov (NCT02293707). All patients were required to sign a written informed consent before enrolling into the study.

The primary end-points were safety assessment and evaluation of immunological response defined as the achievement of an immunological score ≥ 3 (see below). Analysis of clinical efficacy was a secondary end-point.

Main eligibility criteria were (a) previously histologically confirmed diagnosis of m CRPC; (b) documented achievement of response or disease stability after docetaxel chemotherapy.

GX301 vaccine is composed of four hTERT peptides (peptides 540–548, 611–626, 672–686, 766–780) and two adjuvants, Montanide ISA-51 VG and imiquimod. Each hTERT peptide was supplied as 625 µg lyophilised powder vials by Bachem AG, Bubendorf, Switzerland. Montanide was supplied as 3 mL vials by Seppic SA, La Garenne Colombes, France. Imiquimod is a medicinal product marketed as single-dose sachets containing 12.5 mg imiquimod as 5% cream (Meda Pharma SpA, Milan, Italy).

Each GX301 administration consisted of four intradermal injections (one for each peptide) given in the abdominal region and followed by topical application of imiquimod. Each intradermal injection consisted of a fixed hTERT peptide dose, 500 μg, reconstituted as a saline solution and mixed with Montanide (1:1) using a standardized disposable device.

The three GX301 regimens consisted of either eight administrations (Regimen 1) on days 1, 3, 5, 7, 14, 21, 35 and 63, four administrations (Regimen 2) on days 1, 14, 35 and 63, or two administrations (Regimen 3) on days 1 and 63 (Supplementary Table 1). Day 1 was the day of randomization.

Treatment-emergent adverse events (AEs) were recorded throughout on-study observation. AEs were graded for severity according to Common Terminology Criteria for Adverse Events (CTCAE), version 4.0. AEs that were fatal, life-threatening or requiring/prolonging hospitalization, resulting in significant disability, or otherwise judged as medically important events, were classified as serious.

Assessment of the immunological efficacy of GX301 regimens was based on the following tests performed on peripheral blood mononuclear cells (PBMC): (1) Peptide-specific ELISPOT assay for the evaluation of frequency of IFNγ–secreting T lymphocytes; (2) Peptide-specific intracellular staining and flow cytometry analysis for evaluation of the frequency of circulating IFNγ–secreting CD4 + and CD8 + T lymphocytes; (3) Peptide-specific cytotoxic assay.

Blood samples for immunological testing were taken at baseline (randomization), day 90 and day 180. Positive test responses found on days 90 and 180 were considered vaccine-related if they were either new (i.e. not detected at baseline) or greater than twice the baseline value. Individual immunological outcomes were scored as the sum of vaccine-related responses, achieving an immunological score ranging 0 to 6 (three tests by two time-points): the immunological success was defined as the achievement of an immunological score ≥ 3.

The frequencies of IFNγ-secreting CD8 + and CD8- T cells after incubation with the vaccination peptides were evaluated by intracytoplasmic immunofluorescence analyses, as follows. Peripheral blood mononuclear cells (PBMC) (1 × 10⁶ cells), obtained from heparinized peripheral blood by centrifugation on Ficoll gradient and re-suspended in RPMI conditioned by 10% AB serum, were stimulated overnight at 37 °C by a mix of the four GX301 vaccine peptides (5 µg/ml each) in presence of purified anti-human CD28 (clone CD28.2) and anti-CD49d (clone L25) mAbs both at 1 µg/ml concentration (BD). Brefeldine A (BFA, Sigma) (10 µg/ml) was added to samples for the last four hours of incubation. Samples cultured without peptides or stimulated with PMA and ionomycin (Sigma) were considered negative and positive controls, respectively.

Then, washed samples were incubated with vitality dye LIVEDEAD (Molecular Probes, Thermo Fisher) before proceeding with surface staining. The following fluorochrome-conjugated mAbs were used: PE-conjugated anti-human CD8 clone SK1, APC-conjugated anti-human CD3 clone UCHT1 (BD). After surface staining, Cytofix/Cytoperm kit (BD) was used to fix and permeabilize the lymphocytes following the manufacturer’s instructions. The cells were washed in Perm-Wash buffer (BD) and incubated with a FITC-conjugated anti-human IFNγ mAb (BD). Thereafter, the samples were washed in Perm-Wash buffer, re-suspended in FACS Lysing solution (BD) and analysed by a LSR Fortessa X20 flow cytometer (BD) using the FACS DIVA software (BD) v8.1.0. The results were expressed as frequency of IFN-γ producing cells in CD3 + CD8 + or in CD3 + CD8- alive lymphocytes after subtracting the frequency of unstimulated T cells spontaneously producing IFNγ cytokine.

Positive responses were considered those either absent at baseline or greater than twice the baseline showing ≥ 0.1% background positive cells, as suggested for low frequency reactivity [33].

In order to detect IFNγ-producing T cells reactive against the GX301 peptides, ELISPOT analyses were performed on freshly isolated PBMC using the Human IFNγ ELISPOT Kit according to the manufacturer’s instructions (BD) and following the indications of international proficiency panels [34]. Briefly, PBMC (2 × 10⁵ cells in X-VIVO medium, Euroclone) were incubated overnight with a mix of the four GX301 vaccine peptides (5 µg/ml each) in the presence of anti-human CD28 and anti-human CD49d mAbs (BD) (both at 1 µg/ml), or with phytohaemagglutinin (PHA-P, MPBIO) at 1 mg/ml, as positive control, or medium alone as negative controls.

Positive responses were considered those either absent at baseline or greater than twice the baseline showing ≥ 10 spots and ≥ 2 × background spot number.

Vaccine-specific cytotoxic activity of circulating T lymphocytes was analysed by flow cytometry, as follows [35]. PBMC (10 × 10⁶) were re-suspended in 1 ml of PBS containing CFDA-SE 5 µM (Molecular Probes, Thermo Fisher) for 5 min at room temperature and then washed twice in PBS-1% AB serum at 4 °C. Monocytes were positively sorted from labelled-PBMC by CD14 Micro-Beads human Kit according to the manufacturer’s instructions (Miltenyi) and pulsed or not (1 × 10⁵/well) overnight with a mix of the four GX301 vaccine peptides (5 µg/ml each). The day after, PBMC (2 × 10⁶/ml) were incubated for 6 h at 37 °C with 1 × 10⁵ CFDA-SE-labelled, pulsed or un-pulsed, autologous monocytes as target cells. Thereafter, cells were washed with PBS and re-suspended in 300 µl of PBS added with 5 µl of 7-AAD (BD) before flow cytometer analysis. The samples were analysed by a FACSCanto II flow cytometer (BD) using FACS DIVA software (BD) v 6.1.3.

The percentage of specific lysis was calculated as

Achieved values were normalized for the percentage of CD3 + CD8 + T cells detected among PBMC. Positive responses were considered those either absent at baseline or greater than twice the baseline showing ≥ 15% of specific lysis.

Immune phenotyping of peripheral blood lymphocytes was performed as follows. One hundred µl of washed whole blood, collected in Vacutainers containing tetrasodium EDTA, were incubated with pre-mixed, pre-optimized, multicolour ‘cocktails’ of antibodies within 12 × 75 mm flow cytometry tubes (Lyotube, Becton Dickinson, BD) for 30 min at 4 °C. The cocktails were optimized in two panels to evaluate the frequency of T, B, NK cell subpopulations, CD8 + and CD4 + T regulatory (Treg) cells, CD8 + and CD4 + T cell maturation and activation. The first panel included the following fluorochrome-conjugated monoclonal antibodies (mAbs): BD Horizon V450 (V450)-conjugated anti-human HLA-DR clone L243(G46-6), BD Horizon V450 (V500)-conjugated anti-human CD45 clone 2D1, fluorescein(FITC)-conjugated anti-human CD3 clone UCHT1, allophycocyanin(APC)-conjugated anti-human CD8 clone SK1, APC-H7-conjugated anti-human CD4 clone SK3, phycoerythrin(PE)-conjugated anti-human CD16 + CD56 + clones B73.1 and MY31, PE-Cyanin7(PE-Cy7)-conjugated anti-human CD19 clone SJ25C1. The second panel included the following fluorochrome-conjugated mAbs: V450-conjugated anti-human CD45RA clone HI100, V500-conjugated anti-human CD3 clone UCHT1, Brilliant violet(BV)711-conjugated anti-human CD8 clone SK1, FITC-conjugated anti-human CD127 clone HIL-7R-M21, peridinin-chlorophyll proteins(PerCP)-Cy5.5-conjugated anti-human CCR7 clone 150503, APC-conjugated anti-human CD39 clone TU66, APC-H7-conjugated anti-human CD4 clone SK3, PE-conjugated anti-human CD28 clone CD28.2, PE-Cy7-conjugated anti-human CD25 clone 2A3. Cells were then re-suspended in 100 µl of PBS and 10 µl of 7-AminoactinomycinD (7-AAD, BD) were added as viability staining solution to exclude dead cells. Samples were analysed by a LSR Fortessa X20 flow cytometer (BD) using the FACS DIVA software (BD) v8.1.0.

Progression-free survival (PFS) and overall survival (OS) were secondary end-points. Assessment of clinical efficacy was based on the evaluation of serum PSA time-course and on the evaluation of disease evolution through clinical examination and imaging analyses repeated at fixed time intervals or at any time if deemed necessary by the local investigators.

PFS and OS were estimated with the Kaplan–Meier method. Data of patients who were lost to on-study observation or follow-up were censored at the time of the last available information. GX301 regimens were compared for PFS and OS using the log-rank test. Association of selected putative prognostic factors (i.e. time to CRPC diagnosis, class of cumulative docetaxel dose and outcome of docetaxel chemotherapy) with PFS or OS was investigated with Cox regression model. All patients who received at least one GX301 administration were included in the analyses.

Ninety-nine patients were enrolled in the study. One of them withdrew spontaneously: the remaining 98 were randomized into the three regimen groups for receiving either eight (regimen 1, n = 32), four (regimen 2, n = 33) or two (regimen 3, n = 33) vaccine administrations, respectively. All the 98 randomized patients received the vaccine accordingly to the assigned schedule: however, among them, only 63 were assessable for immunological efficacy based on protocol criteria, due to withdrawn from observation before the 180-day time-point of 25 patients.

Baseline patient characteristics are summarized in Table 1.

Safety was analysed in all 98 patients who received at least one vaccine administration. Among AEs, panniculitis-like local inflammation at the site of vaccine administration was common to all patients. The frequency of inflammatory reaction at injection sites increased, as expected, with the number of vaccine administrations. Overall, SAEs were rare and mostly unrelated to GX301 vaccination. In particular, only one fatal event was registered due to the onset of a second neoplasia (glioblastoma multiforme). Table 2 summarizes serious non-fatal adverse events: a total of eight SAEs was observed in two, two and one patients in regimen 1, regimen 2 and regimen 3, respectively.

Importantly, no events relative to induction of severe lymphopenia after vaccination were registered in our series (not shown).

Moreover, no onset of autoimmunity-related clinical signs or of autoantibodies was observed.

Sixty-three patients were assessable for the immunological outcome (n = 20, n = 24 and n = 19 for regimen 1, regimen 2 and regimen 3, respectively). Representative analyses for each type of immunological tests are shown in Fig. 1.

Sixty out of 63 (95%) immunized patients who completed the vaccination protocol showed at least one positive response at one of the tests performed on days 90 and 180 after the first immunization. The only three patients who did not show any vaccine-specific immune response belonged to the regimen 3 (Table 3).

Responders to vaccination, as per protocol criteria, ranged from 42 to 65% of patients with a proportional relationship between rates of immunological responders and number of immunizations administered by each regimen (Table 3).

Since one of the four immunogenic peptides (hTERT_540–548) included in the GX301 vaccine is restricted by the HLA-A₂ allele, we compared the responder rates between the HLA-A₂ positive and negative patients and no differences were observed (not shown).

Interestingly, taking into consideration the total number of positive and negative responses to the six immunological tests performed at days 90 and 180 in either the total patient population receiving the vaccine (n. 98 patients) or the immunological assessable patient population (63 patients), the comparison of the rates of positive responses at any of the six immunological tests among the different regimens showed a significant difference between regimen 2 and regimen 3, while no differences were observed between either regimen 1 and regimen 3 or regimen 1 and regimen 2 (Fig. 2).

The circulating frequencies and absolute numbers of different T cell subsets were assessed at baseline and after 90 and 180 days from baseline. T cell subsets to be analysed were selected for (a) maturation stage, in terms of CCR7 + CD45RA + naïve, CCR7 + CD45RA- central memory (CM), CCR7 + CD45RA- effector memory (EM), and CCR7 + CD45RA + terminal effector memory cells (TEM), and b) regulatory commitment, in terms of both CD4 + CD127-CD25hi and CD8 + CD28-CD127loCD39 + Treg, respectively.

The comparison of circulating T cell subset frequencies or absolute numbers between immunologically responder and non-responder patients showed that at baseline the only difference concerned the frequency of naïve CD8 + T cells, that was lower in responders than in non-responder patients (Fig. 3a).

In order to have a picture on the dynamics of T cell subset frequencies and absolute numbers upon GX301 vaccination, the differences (Δ) were calculated between values at day 90 or at day 180 and values at baseline; then, such differences were compared between responders and non-responders. This analysis showed that responders had a significantly higher increase in absolute number of circulating CD4 + T cells at day 180 than non-responders (Fig. 3b).

Fifty per cent of patients underwent disease progression within day 163 and 75% within day 183 (Fig. 4a), with no statistically significant or trend differences among GX301 regimens (Fig. 4b).

Post-study follow-up was completed by 95% of patients. OS was 62% at 18 months and 48% at 24 months (Fig. 4c) with no statistically significant differences among GX301 regimens (Fig. 4d).

In order to have a rough estimate of clinical efficacy of the vaccine, the analysis of survival following disease progression was calculated for patients with documented disease progression (n = 82 out of 98 enrolled patients). Median survival of patients progressing after vaccine administration was 17.3 months (Fig. 5a). This value increases to 19.9 months limiting the analysis to the 59 patients who were treated at progression with either abiraterone acetate, enzalutamide or cabazitaxel (used alone or in sequential combinations) (Fig. 5b).

In order to investigate on the possible association between immunological response to GX301 vaccine and clinical outcome, PFS and OS were compared between responders and non-responders, irrespective of the assigned regimen. Figure 6a, b shows that no significant differences were observed between the two groups.

Then, we wondered whether the level or dynamics of some T cell subsets could be predictive of clinical outcome. Hence, we found that baseline absolute number of CD4 + Treg impacted on PFS since patients with baseline number < 30.3 CD4 + Treg/μl had a lower risk of progression than patients with baseline number > 30.3 CD4 + Treg/μl (Fig. 7a). Concerning OS, we observed that patients who had a day 180 vs baseline Δ < 37.2 of CD3 + T cell number/μl and < − 0.4 of CD8 + T cell percentage had a more prolonged survival than patients with Δ values > 37.2 CD3 + T cell number/μl and > − 0.4 CD8 + T cell percentage, respectively (Fig. 7b, c).

Interestingly, a decreased number at day 180 of CD8 + Treg (identified as shown in Supplementary Fig. 1) was associated with a better prognosis (Fig. 7d), a phenomenon non-dependent on the trend of the non-Treg CD8 + T cell subpopulation (Fig. 7e).