Vaccinations with tumor cells engineered to produce IL-4 prolonged survival and cured 30% of mice bearing pulmonary metastases, an effect abrogated by in vivo depletion of T cells. Vaccination induced type 2 T cell polarization in both CD4 and CD8 T lymphocyte subsets. We focused on the antitumor activity exerted by type 2 CD8+ T cells (Tc2) activated by IL-4 tumor cell vaccination. Tc2 lymphocytes lacked in vitro tumor cytotoxicity, but released IL-4 upon stimulation with tumor cells, as shown by limiting dilution analysis of the frequencies of tumor-specific pCTL and of CD8 cells producing the cytokine. In vivo fresh purified CD8+ T lymphocytes from IL-4-vaccinated mice eliminated 80–100% of lung metastases when transferred into tumor-bearing mice. CD8+ lymphocytes from IL-4-vaccinated IFN-γ knockout (KO), but not from IL-4 KO, mice cured lung metastases, thus indicating that IL-4 produced by Tc2 cells was instrumental for tumor rejection. The antitumor effect of adoptively transferred Tc2 lymphocytes needed host CD8 T cells and AsGM1 leukocyte populations, and partially granulocytes. These data indicate that Tc2 CD8+ T cells exert immunoregulatory functions and induce tumor rejection through the cooperation of bystander lymphoid effector cells. Tumor eradication is thus not restricted to a type 1 response, but can also be mediated by a type 2 biased T cell response.

In contrast to the current view indicating that a type 1 immune response is required for tumor regression, type 2 cytokines have been successfully used in murine models for augmenting the immunogenicity of tumor cells such to determine in vivo tumor rejection (1).

IL-4, initially known as B cell stimulatory factor (2), has a pivotal role in conditioning type 2 T cell differentiation (3) leading to IgG1 and IgE isotype switching. In several studies, IL-4 showed in vivo tumor growth inhibition through a strong inflammatory effect associated with local recruitment of granulocytes, macrophages, T lymphocytes, and dendritic cells (4, 5, 6, 7). Tumor rejection required polymorphonuclear cells, either eosinophils (8) or neutrophils (9), whereas the role of CD8 T cells remains controversial, since they have been found necessary in one study (6) but not in others (5, 10). IL-4-releasing tumor cell vaccines showed efficacy when used in preclinical therapeutic setting (5, 11), and pilot studies in cancer patients have been proposed (12, 13) and conducted (51).

The aim of this study was to test whether type 2 antitumor CD8 T cells are activated by IL-4-producing tumor cell vaccines and thus studying the mechanism by which they provoke tumor eradication. The colon carcinoma C26/IL-4/FRα cells, engineered to produce IL-4 and to express the human folate receptor α (FRα)3 as a model Ag, were irradiated and used to treat mice bearing C26/FRα pulmonary metastases. We have recently shown that vaccination with C26/IL-4/FRα elicits a strong anti-FRα IgG1 response that is not correlated with the therapeutic outcome, thus suggesting that mechanisms other than the Ab response should be implicated (14). We show here that treatment with IL-4 vaccine induces noncytotoxic CD8+ T cells producing type 2 cytokines able to induce tumor rejection through interaction with host CD8+, ASGm1+, and GR-1+ cells.

Eight- to ten-week-old female BALB/c mice were purchased from Charles River (Calco, Italy); BALB/c IL-4 knockout (KO) and BALB/c IFN-γ KO mice were from The Jackson Laboratory (Bar Harbor, ME). Mice were maintained and treated in compliance with institutional guidelines.

C26/FRα, C26/IL-4/FRα, and C26/IL-12/FRα cells producing murine IL-4 or IL-12 and expressing human FRα were obtained by retroviral transduction, as described (14). For immunotherapy experiments, lung metastases were induced by the i.v. injection of 104 C26/FRα cells, and mice were vaccinated on days 3, 6, 9, and 13, by injecting s.c. 3 × 106 15,000 rad irradiated C26/IL-4/FRα cells, producing 20–30 ng/ml IL-4 (106 cells/ml/48 h). CD4 and CD8 cell depletion was conducted by treating mice, on day 1 and every 2 wk thereafter, with mAbs (0.5 mg, i.p.) obtained from the hybridomas GK 1.5 and 2.43, respectively. For survival, follow-up mice were euthanized when showing dyspnea; survivors were shown to be tumor-free. Lymphocytes were collected from vaccinated mice 5–7 days after last vaccination.

Recipient mice (5–10 mice/group) were i.v. injected with 7.5 × 104 C26/FRα cells to induce lung metastases and treated on the following day by i.v. injection of fresh spleen cells (3.5 × 107) obtained from donors vaccinated with C26/IL-4/FRα tumor cells, as detailed above, and treated with anti-CD4 mAb 3–4 days previously. CD8+ lymphocytes were enriched by elution on nylon wool columns (Wako Chemicals, Dusseldorf, Germany), and 5 × 106 lymphocytes were transferred. The transferred population was shown to be 85–95% CD8+ by flow cytometric analysis. Recipient mice were sacrificed 2 wk after lymphocyte transfer, and the number of superficial lung metastases was counted after lung insufflation, as previously described (14). Recipient mice were depleted of lymphoid subpopulations by treatment with specific mAbs. T cell depletion was obtained by surgical thymectomy at 1 mo of age, followed by treatment with anti-CD4 and/or anti-CD8 1 wk before the beginning of the experiment. Treatment with anti-AsialoGM1 rabbit antiserum (Wako Chemicals) and with anti-granulocyte mAb (RB6-8C5 hybridoma) was given 3 days before and on the day of lymphocyte transfer, at the dose of 10 μl and 0.5 mg, respectively. Ablating treatments were checked by FACS analysis and functional assays to verify target leukocyte depletion. Hybridomas were obtained from (American Type Culture Collection, Manassas, VA); mAbs were purified by using a commercial kit (E-Z-SEP; Amersham Pharmacia Biotech., Uppsala, Sweden).

Lymphocytes obtained from mice vaccinated with C26/IL-4/FRα or with C26/IL-12/FRα cells were restimulated in mixed lymphocyte tumor cell cultures (MLTC) with C26/FRα irradiated cells for 6 days, as described (15). CD4+ and CD8+ cells were then positively selected by paramagnetic beads conjugated with anti-CD8a mAb (Miltenyi Biotec, Bergish Gladbach, Germany) and tested for cytokine production by culturing lymphocytes (105/well) in 96-well flat-bottom plates precoated or not with 0.5 μg/well of anti-CD3 mAb (145-2C11 hybridoma) at 37°C for 20 h. Supernatants were assayed for cytokine content by specific ELISAs (PharMingen, San Diego, CA).

CTL response was assessed in a standard 4-h 51Cr-release assay after 6 days MLTC, as described (15). For LDA, lymphocytes were serially 1:2 diluted (from 8 × 104/well to 500/well) and cultured with 15,000 rad irradiated C26/FRα cells (5 × 103/well), 2000 rad irradiated syngeneic splenocytes as feeder cells (5 × 105/well), and 50 U/ml rhIL-2 (Chiron-Italia, Milan, Italy), in 32 identical replicates, in 96 U-well plates for 10 days. 51Cr-release assays were performed with 103 target cells/well. Specificity for C26/FRα was evaluated by testing replica plates with unrelated target cells. Values of lysis exceeding three times the SD of the mean spontaneous release were used as a threshold for scoring positive cytolysis. Calculation of the frequencies from limiting dilution data was done as previously described (16). The same method was used for calculation of frequencies of IL-4 and IFN-γ-producing precursors; in this case, after 18 h of incubation with unlabeled target cells, supernatants were tested for cytokine content by specific ELISA. The experiments were conducted by directly comparing lymphocytes from mice treated with IL-4 vaccine or with IL-12 vaccine.

Lymphocytes from MLTC were cultured for 18 h with immobilized anti-CD3 mAb and 10 μg/ml brefeldin A added for the last 2 h (Sigma, Milan, Italy). Cells were suspended in PBS containing 1% FCS, fixed with paraformaldehyde 4% for 20 min at room temperature, and permeabilized with saponin buffer containing saponin 0.5% and 0.1% NaN3 (Sigma) for 10 min. For triple staining, FITC-labeled anti-IFN-γ, PE anti-IL-4, and CyChrome anti-CD4 mAbs were used; for double staining, purified CD8+ cells were labeled with FITC anti-CD8, PE anti-IL-4, and PE anti-IFN-γ. Isotype-matched conjugated mAbs were used for background determination. All mAbs were purchased from PharMingen. Cytofluorometric analysis was performed with a FACScan (Becton Dickinson, Mountain View, CA). mAbs dilutions were established by preliminary dose/response experiments, and 50,000 events were acquired.

Active immunotherapy with C26/FRα cells engineered to produce IL-4 increased survival and cured 33% of mice bearing C26/FRα lung metastases (Fig. 1, and Ref. 14). The therapeutic effect was dependent on host T cells, since it was abrogated in mice depleted of CD4 or CD8 T cells by in vivo treatment with specific mAbs (Fig. 1).

FIGURE 1.

Survival of mice bearing C26/FRα lung metastases treated or not by vaccination with C26/IL-4/FRα cells, and of vaccinated mice depleted of CD8 or CD4 T cells. Mice injected i.v. with C26/FRα cells were s.c. vaccinated with 3 × 106 irradiated tumor cells on days 3, 6, 9, and 13. Treatment with anti-CD4 and anti-CD8 mAbs were given on day 1 and repeated every 2 wk. Control mice were left untreated. p < 0.0001 by Log Rank test when comparing survival curves of vaccinated mice (n = 18, median survival time (mst) 34) with that of controls (n = 20, mst 27), of vaccinated mice depleted of CD8 cells (n = 19, mst 19), and of vaccinated mice depleted of CD4 cells (n = 16, mst 27).

FIGURE 1.

Survival of mice bearing C26/FRα lung metastases treated or not by vaccination with C26/IL-4/FRα cells, and of vaccinated mice depleted of CD8 or CD4 T cells. Mice injected i.v. with C26/FRα cells were s.c. vaccinated with 3 × 106 irradiated tumor cells on days 3, 6, 9, and 13. Treatment with anti-CD4 and anti-CD8 mAbs were given on day 1 and repeated every 2 wk. Control mice were left untreated. p < 0.0001 by Log Rank test when comparing survival curves of vaccinated mice (n = 18, median survival time (mst) 34) with that of controls (n = 20, mst 27), of vaccinated mice depleted of CD8 cells (n = 19, mst 19), and of vaccinated mice depleted of CD4 cells (n = 16, mst 27).

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We analyzed the type of T cell response activated by vaccinations with C26/IL-4/FRα cells by assessing cytokine production by ELISA and by flow cytometry. Splenic lymphocytes obtained 5–7 days after the course of vaccinations were restimulated in MLTC with C26/FRα cells for 6 days; CD4+ and CD8+ T cells were then purified and cytokine production examined after stimulation with anti-CD3 mAb for 18 h. The pattern of cytokines produced by either CD4+ (Fig. 2, upper panel) or CD8+ lymphocyte subsets (Fig. 2, lower panel) indicated that treatment with an IL-4-releasing cell vaccine induced polarization toward the type 2 phenotype in both CD4+ and CD8+ T cells. In fact, IL-4, IL-5, and IL-10 were measured at higher levels and IFN-γ at lower quantity, compared with T cells from mice vaccinated with C26/FRα cells engineered to produce IL-12, included in this analysis as a control of type 1 T cell activation (15, 16, 17). The analysis of cytokines produced by T cells from lymph nodes draining the vaccination site showed similar results (data not shown).

FIGURE 2.

ELISA measurement of cytokine production by CD4+ or CD8+ T cells upon vaccination with C26/IL-4/FRα (filled bars) or C26/IL-12/FRα cells (open bars). Lymphocytes were purified from MLTC and stimulated with immobilized anti-CD3 mAb.

FIGURE 2.

ELISA measurement of cytokine production by CD4+ or CD8+ T cells upon vaccination with C26/IL-4/FRα (filled bars) or C26/IL-12/FRα cells (open bars). Lymphocytes were purified from MLTC and stimulated with immobilized anti-CD3 mAb.

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Intracellular staining of purified CD4+ T cells showed a higher number of cells producing IL-4 rather than IFN-γ in IL-4-vaccinated mice (Fig. 3,A), whereas an opposite pattern of expression was found in mice treated with IL-12 vaccine (Fig. 3,B). In addition, these data indicated that IL-4 and IFN-γ were expressed by distinct cells in IL-4-vaccinated mice. Similarly, in CD8+ T lymphocytes, the frequency of IL-4-producing cells was higher than that of IFN-γ-producing cells (Fig. 3, C and D).

FIGURE 3.

Left panel, Expression of IL-4 and IFN-γ by CD4+ T cells from mice treated with C26/IL-4/FRα (A) and with C26/IL-12/FRα cell vaccines (B), as detected by flow cytometric analysis. Lymphocytes from MLTC were stimulated with immobilized anti-CD3 mAb in the presence of brefeldin A, fixed and stained with anti-CD4 Cy, and with anti-IL-4 PE and anti-IFN-γ FITC after permeabilization. By gating on CD4-positive cells, in A, lymphocytes stained with anti-IL-4 mAb were 6.2% and with anti-IFN-γ 2.3%, while in B, 1.3% and 28.7%, respectively. Right panel, Expression of IL-4 (C) and IFN-γ (D) by CD8+ T cells from mice treated with C26/IL-4/FRα vaccines. Cells were stained with anti-CD8 FITC and with anti-IL-4 PE or anti-IFN-γ PE. A total of 4.3% CD8+ cells was stained by anti-IL-4 mAb (C) and 0.8% by anti-IFN-γ mAb (D).

FIGURE 3.

Left panel, Expression of IL-4 and IFN-γ by CD4+ T cells from mice treated with C26/IL-4/FRα (A) and with C26/IL-12/FRα cell vaccines (B), as detected by flow cytometric analysis. Lymphocytes from MLTC were stimulated with immobilized anti-CD3 mAb in the presence of brefeldin A, fixed and stained with anti-CD4 Cy, and with anti-IL-4 PE and anti-IFN-γ FITC after permeabilization. By gating on CD4-positive cells, in A, lymphocytes stained with anti-IL-4 mAb were 6.2% and with anti-IFN-γ 2.3%, while in B, 1.3% and 28.7%, respectively. Right panel, Expression of IL-4 (C) and IFN-γ (D) by CD8+ T cells from mice treated with C26/IL-4/FRα vaccines. Cells were stained with anti-CD8 FITC and with anti-IL-4 PE or anti-IFN-γ PE. A total of 4.3% CD8+ cells was stained by anti-IL-4 mAb (C) and 0.8% by anti-IFN-γ mAb (D).

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Type 2 CD8+ T cells activated by C26/IL-4/FRα vaccine were tested for CTL activity against the tumor. Absence or low levels of C26/FRα tumor cell lysis were detected in short-term cytotoxicity assays (Fig. 4), a result confirmed by testing the frequencies of anti-C26/FRα pCTL in LDA assays. In fact, the frequencies of pCTL against C26/FRα in IL-4-vaccinated mice were shown to be 5- to 10-fold lower than in lymphocytes from IL-12-vaccinated mice in three different assays (Table I).

FIGURE 4.

Cytotoxicity of lymphocytes from mice vaccinated with C26/FRα/IL-4 cells. Splenic lymphocytes were cultured in MLTC for 6 days and then used as effector cells against C26/FRα and P815 target cells in 51Cr-release assay. Lymphocytes from mice treated with C26/FRα/IL-12 cell vaccine were assayed as lytic effectors, untreated tumor bearing control mice as negative control effectors.

FIGURE 4.

Cytotoxicity of lymphocytes from mice vaccinated with C26/FRα/IL-4 cells. Splenic lymphocytes were cultured in MLTC for 6 days and then used as effector cells against C26/FRα and P815 target cells in 51Cr-release assay. Lymphocytes from mice treated with C26/FRα/IL-12 cell vaccine were assayed as lytic effectors, untreated tumor bearing control mice as negative control effectors.

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Table I.

Frequencies of anti-C26/FRα pCTL and IL-4- and IFN-γ-producing CD8+ lymphocytes activated by treatment with IL-4 vaccine or with IL-12 vaccine, as measured by LDA

Lymphocytes from mice vaccinated with:Frequencies of Anti-C26/FRα Precursorsa
CTLIL-4-producingIFN-γ-producing
C26/IL-4/FRα 18 (22-12) 588 (714-398) 138 (171-105) 
C26/IL-12/FRα 80 (117-89) 5 (7-2) 1061 (1236-885) 
    
Lymphocytes from mice vaccinated with:Frequencies of Anti-C26/FRα Precursorsa
CTLIL-4-producingIFN-γ-producing
C26/IL-4/FRα 18 (22-12) 588 (714-398) 138 (171-105) 
C26/IL-12/FRα 80 (117-89) 5 (7-2) 1061 (1236-885) 
    
a

Results are expressed as CD8+ precursors/106 lymphocytes; numbers in parentheses represent the upper and lower 95% confidence limits. For pCTL determination, cytotoxicity was assayed by 6 h of incubation with 51Cr-labeled target cells; cytokine release was tested after overnight incubation with nonlabeled targets. Specificity for C26/FRα was evaluated by testing replica plates with unrelated target cells.

The same LDA was used to determine the frequencies of CD8+ lymphocytes producing cytokines upon recognition of tumor target cells (16). In this assay, to avoid any bias due to CD4-secreted cytokines, we used lymphocytes depleted of CD4+ T cells. In lymphocytes from IL-4-vaccinated mice, the frequency of CD8+ T cell precursors producing IL-4 and specific for C26/FRα was 1/1700, whereas the frequency of those producing IFN-γ was 1/20,345; in the same experiment, lymphocytes from IL-12-vaccinated mice showed a frequency of 1/207,597 and 1/6006 cells producing IL-4 and IFN-γ, respectively (Table I).

To test whether type 2 CD8+ T cells (Tc2) may induce tumor rejection, splenic lymphocytes from C26/IL-4/FRα-vaccinated mice were depleted of CD4 cells, and CD8+ lymphocytes were purified or not before being adoptively transferred into syngeneic recipients bearing C26/FRα lung metastases. Two weeks after lymphocyte transfer, recipient mice were sacrificed to count superficial lung metastatic nodules. Splenocytes depleted of the CD4 T cell population from IL-4-vaccinated mice completely eliminated lung metastases (mean number of metastases, 1.6 ± 0.7), whereas lymphocytes from C26/FRα tumor-bearing or from naive donors did not affect the number of metastatic nodules compared with untreated controls (169 ± 38, 139 ± 39, and 147 ± 46). The transfer of CD8+ cells, purified from the CD4-depleted splenocytes, determined >85% reduction of metastases (24 ± 7), while lymphocytes from vaccinated donors depleted of both CD4 and CD8 cells induced 60% reduction of lung metastases (52 ± 13) (Fig. 5 A). These data indicated that adoptively transferred Tc2 cells can eliminate lung metastases, and that their therapeutic activity is enhanced by coadministration of uncharacterized splenocyte populations not belonging to CD4+ cells.

FIGURE 5.

Antitumor effect of Tc2 lymphocytes against C26/FRα lung metastases. Data are expressed as mean ± SD. ∗, p values ranging from p < 0.05 to p < 0.0001 by comparison with controls, as determined by Student’s t test. A, Adoptive transfer with splenic lymphocytes from CD4 T cell-depleted syngeneic naive donors (SPC naive), C26/FRα tumor-bearing controls (SPC control), or C26/IL-4/FRα vaccinated (SPC IL-4-vacc), and of purified CD8+ T cells (CD8) or of spleen lymphocytes after further CD8 T cell depletion (SPC CD8 depl) from vaccine-treated mice. Controls were left untreated. CD8′ group vs SPC CD4/CD8 depl group, p < 0.02; CD8′ group vs SPC CD4 depl, p < 0.05. B, Adoptive transfer of CD8+ lymphocytes obtained from IL-4 KO (CD8 IL-4KO) and wild-type mice (CD8 wt) vaccinated with C26/IL-4/FRα cells. Controls were left untreated. Mean number of metastases: controls, 98 ± 2; CD8 IL-4 KO, 91 ± 10; and CD8 wt, 8 ± 6. C, Adoptive transfer of CD8+ lymphocytes from IFN-γ KO (CD8 IFNKO) and wild-type (CD8 wt) mice treated with C26/FRα/IL-4 vaccine. Controls, 250 ± 10; CD8 IFNKO, 69 ± 9; and CD8 wt, 55 ± 11.

FIGURE 5.

Antitumor effect of Tc2 lymphocytes against C26/FRα lung metastases. Data are expressed as mean ± SD. ∗, p values ranging from p < 0.05 to p < 0.0001 by comparison with controls, as determined by Student’s t test. A, Adoptive transfer with splenic lymphocytes from CD4 T cell-depleted syngeneic naive donors (SPC naive), C26/FRα tumor-bearing controls (SPC control), or C26/IL-4/FRα vaccinated (SPC IL-4-vacc), and of purified CD8+ T cells (CD8) or of spleen lymphocytes after further CD8 T cell depletion (SPC CD8 depl) from vaccine-treated mice. Controls were left untreated. CD8′ group vs SPC CD4/CD8 depl group, p < 0.02; CD8′ group vs SPC CD4 depl, p < 0.05. B, Adoptive transfer of CD8+ lymphocytes obtained from IL-4 KO (CD8 IL-4KO) and wild-type mice (CD8 wt) vaccinated with C26/IL-4/FRα cells. Controls were left untreated. Mean number of metastases: controls, 98 ± 2; CD8 IL-4 KO, 91 ± 10; and CD8 wt, 8 ± 6. C, Adoptive transfer of CD8+ lymphocytes from IFN-γ KO (CD8 IFNKO) and wild-type (CD8 wt) mice treated with C26/FRα/IL-4 vaccine. Controls, 250 ± 10; CD8 IFNKO, 69 ± 9; and CD8 wt, 55 ± 11.

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To assess the role of IL-4 production in the antitumor effect of the transferred lymphocytes, syngeneic IL-4 KO mice were used as lymphocyte donors. C26/IL-4/FRα vaccination of IL-4 KO mice induced CD8+ T cells producing the same level of IFN-γ than the wild-type vaccinated counterparts, but no IL-4 in response to stimulation with C26/FRα tumor cells (data not shown). The transfer of IL-4 KO-purified CD8+ T cells was ineffective (Fig. 5,B), indicating that IL-4 production by the infused lymphocytes is needed for mediating the antitumor effect. In contrast, the transfer of lymphocytes from IFN-γ KO-vaccinated donors significantly reduced the number of metastases, thus ruling out an effect of IFN-γ-producing CD8+ cells within the Tc2 population in the rejection of metastases (Fig. 5 C).

To investigate the host lymphoid effectors enrolled in the mechanism of tumor rejection, recipient mice were depleted of different leukocyte populations by treatment with specific mAbs before lymphocyte transfer. When recipients were depleted of CD8 T cells or of AsGM1-positive cells, the eradication of metastases was abolished, supporting a fundamental role of host CD8 and NK cells that are likely activated by transferred Tc2 cells, to eliminate lung metastases. Also, granulocyte depletion reduced the antimetastatic effect, whereas depletion of CD4 cells did not change the therapeutic effect of adoptively transferred Tc2 lymphocytes (Fig. 6). Thus, Tc2 lymphocytes eradicated lung metastases through the cooperation of bystander host CD8 T cells, NK cells, and granulocytes, but not CD4 lymphocytes.

FIGURE 6.

Host effector cells determine rejection of metastases upon adoptive transfer of Tc2 lymphocytes. Data are expressed as mean ± SD. ∗, p values ranging from p < 0.05 to p < 0.0001 by comparison with controls, as determined by Student’s t test. Untreated recipient mice (untreated, 2 ± 0.7), thymectomized recipients depleted of CD4 (CD4 depl, 11 ± 10) or of CD8 T cell subsets (CD8 depl, 143 ± 34), or both (CD4/CD8 depl, 210 ± 16), mice depleted of granulocytes (granulocyte depl, 64 ± 18) and in AsialoGM1-depleted mice (AsGM1 depl, 198 ± 32) were adoptively transferred with Tc2 lymphocytes. Control mice not injected with lymphocytes had 139 ± 29 metastases (data not shown).

FIGURE 6.

Host effector cells determine rejection of metastases upon adoptive transfer of Tc2 lymphocytes. Data are expressed as mean ± SD. ∗, p values ranging from p < 0.05 to p < 0.0001 by comparison with controls, as determined by Student’s t test. Untreated recipient mice (untreated, 2 ± 0.7), thymectomized recipients depleted of CD4 (CD4 depl, 11 ± 10) or of CD8 T cell subsets (CD8 depl, 143 ± 34), or both (CD4/CD8 depl, 210 ± 16), mice depleted of granulocytes (granulocyte depl, 64 ± 18) and in AsialoGM1-depleted mice (AsGM1 depl, 198 ± 32) were adoptively transferred with Tc2 lymphocytes. Control mice not injected with lymphocytes had 139 ± 29 metastases (data not shown).

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Initially shown for CD4 T cells, it is now clear that both CD4 and CD8 T cell subsets differentiate into Ag-specific type 1 and type 2 cells characterized by the production of IFN-γ or of IL-4 (18, 19). The IFN-γ or IL-4 conditioned cytokine milieu in which primary Ag stimulation takes place has a major role in directing the differentiation of CD8 T cells from naive T lymphocytes into either type 1 or type 2 effectors (20, 21, 22, 23). Tc2 have also been isolated from PBL and mucosal tissues (24, 25) in healthy individuals and during infectious diseases (26, 27). Tc2 are cytotoxic in some systems (20, 22) but not in others (28, 29), can provide B cell help (29), induce delayed-type hypersensitivity reactions (30), and their type 2 phenotype is maintained in memory responses (22, 23). Although one of the first reports showing IL-4 production by CD8 lymphocytes dealt with tumor immune mice (31), no data are available about the presence and function of tumor-specific Tc2 cells. For this reason, after Tc2 cells were detected in mice treated with IL-4 vaccine, the present study was focused on the assessment of their antitumor potential.

Tc2 cells activated in mice treated with IL-4-vaccine were poorly cytolytic. When purified from MLTC and stimulated by anti-CD3 mAb, CD8+ cells from mice treated with IL-4 vaccine were shown to produce IL-4, IL-5, and IL-10, while IFN-γ was measured at lower quantities as compared with lymphocytes obtained from mice treated with the IL-12-producing vaccine. In addition, TNF-α and GM-CSF were released at levels similar between the two groups (data not shown). Intracellular immunostaining directly demonstrated the presence of IL-4-producing CD8+ cells. However, the general poor staining of intracellular IL-4 (32), did not allow to obtain a trustworthy quantitative data about the frequency of IL-4-producing CD8+ cells in lymphocytes from mice treated with IL-4 vaccine. To solve this difficulty, we set up LDA experiments that showed frequencies of IL-4-producing, tumor-specific CD8 lymphocytes 10-fold higher than that of IFN-γ-producing cells and 20-fold higher than that of cytotoxic pCTL.

The adoptive transfer of enriched CD8+ cells from mice vaccinated with C26/IL-4/FRα cells in mice bearing C26/FRα lung metastases resulted in an 80–100% reduction in the number of tumor nodules in four different experiments. Noncytolytic CD8 lymphocytes obtained from tumor infiltrate (TIL), as well as CD4 Th cells, have been reported to eradicate tumors upon adoptive transfer through their release of cytokines in different murine models (33, 34, 35, 36) and in a recent pilot trial (37). Although the production of type 2 cytokines has not been evaluated in TILs (38, 39), the ability of a Th2-type CD4 T cell clone to initiate tumor eradication has been reported (40).

Our data underline a direct relationship between IL-4 production and the rejection of metastases. Tc2-secreted IL-4 may act through the enhancement of cytotoxicity of host CD8 T cells (41) and by activating eosinophils and basophils, in concert with IL-5 (42). In addition, the released TNF-α may synergize with IL-4 in activating endothelial cells, thus increasing vascular permeability and facilitating leukocyte infiltration, and by induction of NK cell activation (43). Moreover, IL-10 has a chemoattracting effect on CD8 T cells (44).

Depletion experiments clearly showed the participation of host CD8 T cells, asialoGM1-positive lymphocytes, and granulocytes in the process of metastases rejection. Although asialoGM1 is not expressed only on NK cells but also on a subset of CD8 T cells, these data indicate that all the mentioned leukocyte populations participate in the reduction of metastases.

The antitumor activity of Tc2 cells in adoptive immunotherapy can be potentiated by other donor leukocyte populations. In fact, injection of 35 × 106 spleen cells depleted of CD4 lymphocytes reduced the number of metastases to a greater extent compared with injection of 5 × 106 purified CD8+ cells, the estimated CD8+ content of such splenocytes (Fig. 6 A). Whether antitumor Ab-producing B cells, activated macrophages, or granulocytes play the major role in this phenomenon remains to be determined. Whether Tc2 are endowed with antitumor activity similar to that of Tc1 cells was not evaluated in the present study. When tested in parallel, CD8+ lymphocytes obtained from mice treated with IL-4 vaccine or with IL-12 vaccine reduced metastases to the same extent upon adoptive transfer (data not shown). However, in the studied tumor model, IL-12-releasing vaccine showed a superior therapeutic effect compared with IL-4 vaccine treatment, probably because of the activation of a stronger Ab response (14). Mice cured after either vaccine treatment were immune to a subsequent tumor challenge injection, and either Tc2 or Tc1 CD8+ cells were shown when the memory response was assayed in vitro (data not shown).

The data presented here have a number of important implications for tumor immunotherapy. First, they indicate Tc2 cells as a new lymphocyte population endowed with antitumor potential. Second, they highlight the independence from a strict type 1 response for immune tumor rejection. The T1/T2 paradigm implicated that cytokines produced by T1 cells being primarily associated with cell-mediated immune response would augment T cell response against tumors, whereas type 2 cytokines, like IL-4 and IL-10, were considered immunosuppressive and therefore inhibiting antitumor responses (45). Recent reports suggest that the T1/T2 paradigm is not predictive of whether a particular pathway is protective or not (46, 47). In addition, the requirement for both IL-4 and IFN-γ-dependent responses for obtaining B16 melanoma rejection in immune mice has been demonstrated (48). The data shown here would argue for the inconsistency of T1/T2 paradigm in immune response against tumors.

Finally, our data suggest that activation of cytokine-producing tumor-specific T cells, not only IFN-γ but also IL-4, might be a useful indicator for the immunological follow-up of cancer patients treated with active immunotherapy. To date, the major effort in monitoring vaccination trials has focused on the detection of antitumor CTL, though several preclinical and clinical studies had shown that CTL activation is not correlated with the clinical response (16, 49, 50). The activation of noncytotoxic Tc2 cells should be evaluated in patients treated with tumor vaccines producing IL-4.

We thank Mariella Parenza for excellent technical assistance and Ms. Grazia Barp for editorial help.

1

This study was supported by Italian Association for Cancer Research, by Istituto Superiore di Sanita Special Program on Gene Therapy, and by Consiglio Nazionale delle Ricerche Target Project on Biotechnology.

3

Abbreviations used in this paper: FRα, folate receptor α; KO, knockout; MLTC, mixed lymphocytes tumor cell culture; LDA, limiting dilution analysis; Tc2, type 2 CD8+ T cells.

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