For the present study, we used PBMCs (peripheral blood mononuclear cells) from 15 patients (8 males and 7 females) with pancreatic ductal adenocarcinoma (according to the TNM classification for pancreatic tumors) (35), who were already enrolled in a previous study approved by the local Ethics Committee (24). The age of PDAC patients ranged from 36 to 92 years (mean age, 63 years). All patients underwent surgical resection of the primary lesion but did not receive chemotherapy. Patients with evidence of serious illness, immunosuppression, autoimmune or infectious diseases were excluded. Characteristics of PDAC patients and ENO1 expression of PDAC tissue are summarized in Table I. For comparison with healthy controls, 15 blood donors (8 males and 7 females, mean age 60) were studied.

T cell cultures were carried out in RPMI-1640 culture medium (SERO-Med GmbH, Vienna, Austria) supplemented with 10% FCS HyClone Laboratories (Gibco Laboratories, Grand Island, NY, USA) and recombinant human interleukin-2 (IL-2) (Eurocetus, Milan, Italy). Unlabeled or fluorochrome-conjugated anti-CD3, CD4, CD8, CD25, and isotype-matched control mAbs were purchased from BD Biosciences (San Jose, CA, USA). Fluorochrome-conjugated anti-IL-10, anti-TGF-β and anti-FoxP3 mAbs were purchased from eBioscience (San Diego, CA, USA).

Recombinant α-enolase was overexpressed in the E. Coli strain BL21 (DE3)/pLysS (kindly provided by A. Giallongo, Institute of Biomedicine and Molecular Immunology, National Research Council, Palermo, Italy) and was produced as previously reported (23). Anti-ENO1 IgG levels were measured by an in-house ELISA kit (23).

The fasting venous blood (2 ml) at the early morning of the study objects was drawn and the serum was centrifuged and stored at −20°C until assayed. The electrochemiluminescence immunoassay method was used for detecting the levels of CA19-9. The procedures were conducted strictly according to the instructions of kits. The critical value of CA19-9 was 37 U/ml.

Immunohistochemical analysis on 15 PDAC tissues was performed with an anti-α-enolase mAb (1:100; clone 19/12) followed by a horseradish peroxidase (HRP)-conjugated goat anti-mouse IgG (Santa Cruz Biotechnology, Celbio s.p.a., Milan, Italy) and DAB detection kit (Ventana Medical Systems, Inc., Tucson, AZ, USA). Samples were fixed with formalin and paraffin-embedded. Slides were incubated with the primary and secondary Ab and developed in a Ventana ES automated stainer. Stained sections were counterstained with hematoxylin.

All images were taken with a x60 objective. Images were acquired on a Leica DM LA upright microscope equipped with a DC300F camera and were analyzed with IM50 software (Leica Microsystems, Heidelberg, Germany).

To assess the presence of ENO1-specific T cells in the peripheral blood of PDAC patients, PBMCs obtained by density gradient centrifugation on Ficoll-Hypaque were re-suspended in medium supplemented with 3% human serum. PBMCs (3×10⁵) were cultured for 96 h in the presence of medium alone or ENO1 (10 μg/ml). At 16 h before harvesting, 0.5 μCi/well [³H]-thymidine (Amersham Pharmacia Biotech, Uppsala, Sweden) was added, and the radionuclide uptake was measured by a β-counter. The mitogenic index (MI) was calculated as the ratio between mean values of cpm (counts per minute) obtained in ENO1-stimulated cultures and those obtained in the presence of medium alone. An MI ≥3 was considered as a positive result.

ENO1-specific T cell lines were generated as previously described (25). Briefly, 10⁶ PBMCs in 2 ml RPMI-1640 medium supplemented with 2 mM L-glutamine, 2×10⁻⁵ M 2-ME, and 5% human serum (complete medium) were stimulated with ENO1 (10 mg/ml) in 24-well flat-bottomed plates for 5 days. Then, human IL-2 (30 U/ml) was added, and cultures were maintained for 7 days. Viable T-cell blasts were re-suspended in complete medium and cloned under limiting dilution, as previously described (36). Briefly, single T cell blasts were seeded in microwell plates (0.3 cells/well) in the presence of 2×10⁵ irradiated (9000 rad) PBMCs, phytohemagglutinin (PHA) (0.5% vol/vol) and IL-2 (50 U/ml). At weekly intervals, 2×10⁵ irradiated PBMC and IL-2 were added to each microculture to maintain the expansion of growing clones.

Tcc were screened for responsiveness to ENO1 by measuring [³H]-thymidine uptake after a 60-h co-culture with irradiated autologous mononuclear cells in the presence of medium or ENO1 (10 mg/ml). An MI ≥3 was considered as a positive result.

To verify the clonality of ENO1-specific Tcc, the repertoire of the T cell receptor (TCR) Vβ chain of single Tcc was analyzed with the TCR Vβ Repertoire kit (Beckman Coulter, Fullerton, CA, USA); isotype-matched non-specific IgG were used as negative controls and the T cell blasts were gated by anti-CD3 monoclonal antibody.

We analyzed surface marker (CD3, CD4 and CD8) expression in blasts of single Tcc by flow cytometry as previously described (37), along with the production of the intracellular marker (FoxP3) and cytokines (IL-10 and TGF-β) of ENO1-specific Tcc (unable to secrete IFN-γ, IL-4 and IL-17). For intracellular analysis, blasts were stimulated with PMA (phorbol myristate acetate) (10 ng/ml) plus ionomycin (200 ng/ml), and stained with anti-IL-10, anti-TGF-β and anti-FoxP3 mAbs. Tcc that were negative for FoxP3, IL-10 and TGF-β were defined as T null, and Tcc that were positive for FoxP3, IL-10 and TGF-β were defined as Tregs.

To evaluate the amount of secreted cytokines, T cell blasts of single Tcc were resuspended at a concentration of 10⁶ cells/ml of medium and cultured for 36 h in the presence of PMA (10 ng/ml) plus ionomycin (200 ng/ml). Cell-free supernatants were collected and assayed in duplicate for IFN-γ, IL-4 and IL-17 content by commercial ELISA kits (BioSource International, Camarillo, CA, USA). Supernatants presenting cytokine levels that were 5 SD above the mean levels in control supernatants derived from irradiated APCs alone were considered positive. Based on the cytokine profile and the CD4/CD8 expression evaluation, we divided the clones into the following groups: Th1/Tc1 (only IFN-γ), Th2/Tc2 (only IL-4), Th0/Tc0 (IFN-γ and IL-4) and Th17/Tc17 (IL-17).

B cells were prepared using the B cell isolation kit II (Miltenyi Biotec GmbH, Bergisch Gladbach, Germany). To obtain EBV-transformed lymphoblastoid B cell lines (EBV-B cells), autologous B cells from each patient were incubated for 48 h with EBV-producing marmoset B95.8 cell line supernatants, and subsequently expanded in complete medium supplemented with 15% FCS.

Perforin-mediated cytolytic activity of Tcc was assessed as previously reported (24). T cell blasts of ENO1-specific T clones were incubated at an effector-to-target ratio of 10-, 5- and 2.5 to-1 with ⁵¹Cr-labeled autologous (EBV)-B cells pre-incubated with ENO1 (1 μg/ml). After centrifugation, microplates were incubated for 8 h at 37°C, and 0.1 ml supernatant was removed from each microculture for measuring ⁵¹Cr release. To evaluate whether Tcc-specific cytotoxicity against target cells was ENO1/HLA-dependent, cytotoxicity tests were also carried out under two other different conditions: i) with ENO1-pulsed ⁵¹Cr-labeled autologous EBV-B cells in the presence of anti-HLA-DR blocking mAbs (10 mg/ml) to evaluate the ENO1-specific CD4⁺ Tcc; ii) in the presence of anti-HLA-A-B-C to assess the ENO1-specific CD8⁺ Tcc. Anti-HLA-DR and anti-HLA-A-B-C mAbs were purchased from Immunotech (Beckman Coulter, Marseille, France).

The ability of ENO1-specific Tcc to induce Fas-FasL-mediated apoptosis was assessed using Fas⁺ Panc1 cells as targets. T cell blasts from each clone were co-cultured with ⁵¹Cr-labeled Panc1 cells at an effector-to-target ratio of 10-, 5- and 2.5- to-1 for 18 h in the presence of PMA (10 ng/ml) and ionomycin (200 ng/ml), as previously reported (24). To block the Fas-FasL interaction, the anti-Fas antagonistic monoclonal antibody M3 (Amgen, Thousand Oaks, CA, USA) was used, at a final concentration of 5 mg/ml, for a 30-min pre-treatment of ⁵¹Cr-labeled Panc1 cells, as reported (24).

To assess the ability of antigen-specific Treg clones to suppress the antigen-induced autologous cell proliferation of ENO1-specific effector Tcc, 2×10⁴ of these blasts were cultured with 4×10³ irradiated (9000 rad) autologous ENO1-loaded or TT (Tetanus toxoid)-loaded dendritic cells (DCs) in the presence of 2×10⁴ cells of ENO1-specific Treg. DCs were obtained using the Blood Dendritic Cell Isolation kit II (Miltenyi Biotec). At day 4, after 8 h of pulsing with 0.5 mCi 3H-TdR/well (Amersham, Little Chalfont, UK), cultures were harvested and the radionuclide uptake was measured by β-counter analysis. Tetanus toxoid was used for specificity control of T cell clones.

Genomic DNA was extracted from ENO1-specific T cell clones obtained from three tumor samples and three peripheral blood samples of the same patient using the Maxwell 16 Tissue DNA Purification kit and the Maxwell 16 Instrument (Promega, Madison, WI, USA).

TCR gene rearrangement analysis was performed using a gene clonality assay based on the BIOMED-2/Euroclonality multiplex-PCR protocol (38), known as the IdentiClone TCRβ TCRγ T-Cell Clonality Assa-ABI Fluorescence Detection (Invivoscribe, San Diego, CA, USA). The assay consists of five tubes each containing primers that target the TCRβ (tubes A-B-C) and the TCRγ (tubes D–E) chain locus. Briefly, 100 ng of genomic DNA per tube was subjected to PCR amplification according to the manufacturer's instructions. PCR products were run on an ABI 3130 Genetic Analyzer and Genescan profiles were obtained using GeneMapper (version 3.2) software (Applied Biosystems, Foster City, CA, USA).

Statistical analysis of the mitogenic index was evaluated by the unpaired two-tailed Student's t-test. A P-value <0.05 was considered statistically significant. A two-tailed Pearson r coefficient was used to correlate the titer of anti-ENO1 IgG and the percentage of ENO1-specific T clones obtained from PDAC patients. The correlation between the number of ENO1-specific Tcc and survival of patients and hazard ratio (HR) were evaluated by Kaplan-Meier analysis (log-rank Mentel-Cox test).

In order to assess the presence of ENO1-specific T cells in the peripheral blood of PDAC patients, PBMCs from 15 patients, which were previously characterized for the presence of tumor infiltrating T cells, (16) were cultured for 5 days in the presence of medium alone or recombinant ENO1. Notably, PBMCs from 10 of these PDAC patients (67%) (Table I and Fig. 1A) proliferated in the presence of ENO1. Conversely, in the healthy controls, only 3 cases out of 15 (20%) showed proliferation in response to ENO1. In addition, the proliferation of PBMC of PDAC patients showed a significantly greater mitogenic index (mean MI ± SEM, 14.4±2.5) with respect to that of the three healthy controls (mean MI ± SEM, 3.3±0.8; P=0.0002).

As five PDAC patients did not have detectable ENO1-specific T cells in their peripheral blood, but four out of these five patients did, however, present ENO1-specific tumor-infiltrating T cells (16) we expanded and cloned peripheral blood lymphocytes, by IL-2, from all PDAC resectable patients; we obtained 181 Tcc from all PDAC patients except patients 3, 6 and 10 (Table I).

There were 88% (159/181) of CD4⁺ Tcc and 12% of CD8⁺ Tcc (22/181). Interestingly, out of 22 CD8⁺ Tcc, 7 were isolated from the blood of patient 5 Pcp, who was reported as having the highest number of CD8⁺ Tcc.

As ENO1 is a TAA that induces an integrated humoral and cellular response in PDAC patients (15). Tcc obtained were assayed for proliferation in response to ENO1, and 30% of the Tcc was (54/181, Table I) proliferated upon stimulation with ENO1. With the exception of three cases (Pcp 3,6,10), we obtained ENO1-specific Tcc from all patients; in particular patients 9 and 15 were recorded as having the greatest number of Tcc (8 and 9, respectively).

Clonality was confirmed by cytofluorimetric analysis of TCR-Vβ expression. Each Tcc was only revealed by one of the TCR-Vβ chain-specific monoclonal antibodies, showing a single peak of fluorescence intensity (Fig. 1B).

We have already shown that all patients (except for patient PCp10) had detectable serum anti-ENO1 IgG (Table I). Interestingly, we observed that patients with higher titers of anti-ENO1 antibodies also showed the highest percentage of ENO1-specific T clones (r=0.8745 and P-value <0.0001) (Fig. 1C); particularly patients 9 and 15.

Of the ENO1-specific Tcc, 51 were CD4⁺, 21 (41%) produced IL-17 (Th17) and among these 12 (24%) secreted both IL-17 and IFN-γ (Fig. 1D). A total of 15 ENO1-specific CD4⁺ Tcc secreted IL-4, either alone (6 Th2, 12%) or with IFN-γ (9 Th0, 18%). Finally, 14 (27%) ENO1-specific CD4⁺ Tcc produced only IFN-γ, showing a Th1 profile. Notably, taken together, of the 14 Th1 clones and the 12 Th17 clones producing IFN-γ and the 9 Th0 Tcc, we found that the majority of ENO1-specific CD4⁺ Tcc (%) were able to produce IFN-γ. The last ENO1-specific CD4⁺ Tcc was FoxP3⁺ and produced IL-10 and TGF-β, showing a Treg profile. Of the remaining three ENO1-specific CD8⁺ Tcc, one had a regulatory profile (defined as Tcreg) and two had a Th1 profile (defined as Tc1) (Fig. 1D).

To evaluate the anticancer properties of the ENO1-specific Tcc isolated from the peripheral blood of PDAC patients, we assessed their cytotoxicity using ENO1-pulsed ⁵¹Cr-labeled autologous EBV-B cells as targets. At an effector-to-target ratio of 10:1, all 14 Th1, and 7 out of 9 Th0 clones were shown to lyse their targets (Fig. 2A). Among the Th17 population, only the 12 Tcc IL-17/IFN-γ double producers were able to kill their target cells.

None of the six Th2 or the ENO1-specific Treg Tcc lysed the EBV-B cells, while only the two ENO1-specific Tc1 CD8⁺ clones killed the EBV-B cells.

As expected, in the presence of anti-HLA class II blocking mAbs, all the ENO1-specific CD4⁺ Tcc were unable to kill the EBV-B cells, and similar results were obtained for the two ENO1-specific Tc1 in the presence of anti-HLA class I blocking mAbs (Fig. 2A).

Effector T cells can kill their targets by Fas-FasL-mediated apoptosis, and therefore, the ability of ENO1-specific Tcc to induce ⁵¹Cr release by Fas⁺ Panc1 cells was evaluated. Upon mitogen activation, 57% of CD4⁺ (29/51) and 66% of CD8⁺ (2/3) ENO1-specific Tcc induced apoptosis in target cells (Fig. 2B). Among the CD4⁺ Tcc, 14 were Th1, 12 were Th17, and 3 were Th0, but none were Th2 or Treg (Fig. 2B). The role of the Fas-FasL interaction in ⁵¹Cr release was confirmed by its inhibition (range, 38.3–57.6%) using an anti-Fas antibody (data not shown).

Finally, we evaluated the ability of the unique ENO1-specific Treg Tcc (Tcc 6) to inhibit the proliferation of ENO1-effector Tcc (Th1 and Th17) obtained from the same patient (Pcp 8). As shown in Fig. 2C, the ENO1-specific Treg clone inhibited proliferation of both ENO1-specific effector Tcc: the Th1 (Tcc 18) and the Th17 (Tcc 24).

As ENO1-specific Tcc was detected in both the peripheral blood and tumor areas in 73% of the analyzed PDA patients (24), we tested the hypothesis that the same T cell clones were circulating from the periphery to the tumor area and vice versa. We reasoned that if at least one Tcc from the periphery and one Tcc from the tumor area of the same PDAC patient showed the same rearrangement of their TCR, this would provide a proof of concept that the same T lymphocyte was able to recirculate from the tumor to the peripheral blood and vice versa.

Three ENO1-specific Tcc (1Th1, 1 Th17 and 1 Tc1) from the blood and from the central tumor area of the same PDAC patient (Pcp9) were analyzed for TCRβ and γ chain rearrangements by using a classical gene clonality assay (36). Results confirmed the identity of the CD8⁺ Tcc with a Tc1 profile from blood and from tumors (Fig. 3).

Because the majority of ENO1-specific Tcc specifically killed pancreatic cancer cells and were able to re-circulate from the periphery to the tumor and vice versa, we evaluated if the number of peripheral ENO1-specific Tcc correlated with patient survival from diagnosis.

For this purpose, we divided the patients into two groups based on the percentage of peripheral ENO1-specific Tcc and on serum anti-ENO1 specific antibodies: i) <20, and ii) >20 (Table I). Kaplan-Meier analysis revealed that the patients of group ii showed a better survival compared to group i (P<0.0001, Hazard Ratio=31) suggesting a positive correlation between the percentage of circulating ENO1-specific cells and a longer survival (Fig. 4). In addition, the levels of serum CA19-9 (Table I) in patients of group I were significantly (P<0.0001) higher than those in PDAC patients of ii group (data not shown). On the contrary, the PDAC expression of ENO1 is similar in the different patients of the two groups. PDAC tissues diffusely expressed α-enolase in the cells lining the neoplastic ducts infiltrating the stroma. In other words, Enolase is overexpressed in most of PC tissues, and to discriminate between high and lower overexpression is difficult. Thus, it is not possible to achieve a correlation between the expression of Enolase and the presence of specific ENO1-specific Tcc.