Purpose: Small cell lung cancer (SCLC) possesses high tendency to disseminate. However, SCLC patients with paraneoplastic syndrome mediated by immunity against onconeural antigens remain in limited-stage disease (LD) without distant metastases. Cumulative evidence regulates that a balance between immune and regulatory T (Treg) cells determines the magnitude of immune responses to not only self-antigens but also tumor-associated antigens. The purpose of this study was to elucidate the immunologic balance induced in SCLC patients.

Experimental Design: We analyzed T cells in the peripheral blood of 35 consecutive SCLC patients, 8 long-term survivors, and 19 healthy volunteers.

Results: Purified CD4+ T cells with down-regulated expression of CD62L (CD62Llow) produced IFN-γ, interleukin (IL)-4, and IL-17, thus considered to be immune effector T cells (Teff). Significantly more Teff cell numbers were detected in LD-SCLC patients than that of extended-stage SCLC (ED-SCLC). By contrast, induction of CD62LhighCD25+ CD4+ Treg cells was significantly higher in ED-SCLC patients. Long-term survivors of SCLC maintained a high Teff to Treg cell ratio, whereas patients with recurrent disease exhibited a low Teff to Treg cell ratio. Teff cells in LD-SCLC patients included more IL-17–producing CD4+ T cells (Th17). Moreover, dendritic cells derived from CD14+ cells of LD-SCLC patients secreted more IL-23.

Conclusion: These results show that CD4+ T-cell balance may be a biomarker that distinguishes ED-SCLC from LD-SCLC and predicts recurrence. This study also suggests the importance of inducing Teff cells, particularly Th17 cells, while eliminating Treg cells to control systemic dissemination of SCLC.

Translational Relevance

This study indicated that effector T cell to regulatory T cell ratio could be a useful biomarker to distinguish limited-stage disease from extended-stage small cell lung cancer and that therapy to increase Th17 and deplete regulatory T cells is a promising strategy.

Small cell lung cancer (SCLC) is an aggressive disease with a strong tendency to disseminate. Approximately 15% to 20% of SCLC patients whose tumors are confined to the hemithorax and mediastinum and lack detectable distant metastases are considered to be limited-stage disease (LD). These LD-SCLC patients are often cured by treatment management. However, although no distant metastases are detected in LD-SCLC patients, regional treatments such as surgical resection or thoracic radiation therapy alone seldom result in a cure suggesting systemic micrometastases. Thus, repeated invasion by SCLC cells into the peripheral blood, some of which remain in the blood as circulating tumor cells, is considered to occur not only in extended-stage disease (ED) patients possessing prominent distant metastases but also in LD patients (1, 2). The reason why circulating tumor cells are unable to establish visible distant metastases in LD patients is unclear because no biological differences have been detected between tumor cells of LD-SCLC and ED-SCLC.

SCLC is considered to be a relatively immunogenic tumor because it occasionally causes paraneoplastic syndromes such as the Lambert-Eaton myasthenic syndrome (LEMS) mediated by an immunologic mechanism that recognizes shared onconeural antigens. Interestingly, SCLC patients suffering from LEMS tend to remain in long-term LD state and have favorable prognoses (3). Thus, it is postulated that the immune response, which attacks the neuromuscular system in LEMS patients, also fights the SCLC cells to constrain tumor progression.

In the immune system, regulatory CD4+ T (Treg) cells with constitutive expression of the interleukin (IL)-2 receptor α chain (CD25) and the transcription factor forkhead box P3 (FOXP3) play a pivotal role in peripheral tolerance to self and non-self antigens, including tumor-associated antigens (4). It has been shown that Treg cell numbers increase in cancer-bearing patients and that the number of Treg cells correlates with the prognosis (510). We have reported that in addition to tumor-specific Treg cells, antitumor effector T (Teff) cells were generated in the same tumor-draining lymph nodes during tumor progression (11). T cells with down-regulated CD62L expression (CD62Llow) that were isolated from the tumor-draining lymph nodes mediated antitumor reactivity when infused i.v., resulting in the regression of established tumors (12, 13). When coinfused, Treg cells purified as CD62LhighCD25+ CD4+ from the same tumor-draining lymph nodes could inhibit the antitumor therapeutic efficacy of Teff cells. Importantly, the suppression of antitumor reactivity depended on the ratio of Teff to Treg cells (11). Taken together, it seems that the balance between induced CD4+ Teff and Treg cells determines the immune responses against tumors. However, the CD4+ T-cell balance in human malignancies has not been elucidated.

Here, we examined the peripheral blood mononuclear cells (PBMC) in 35 consecutive SCLC patients, 8 long-term survivors who had been disease-free for >3 years after treatment, and 19 healthy volunteers after obtaining a written informed consent. CD62Llow CD4+ T cells isolated from the peripheral blood exhibited primed effector T-cell function to secrete types 1 (Th1), 2 (Th2), and 17 (Th17) helper T-cell cytokines. On the other hand, the CD62LhighCD25+ CD4+ T-cell subpopulation showed cytosolic expression of FOXP3 and regulatory function to suppress cytokine production and inhibit the proliferation of Teff cells. A reciprocal balance was detected between the CD62LhighCD25+ CD4+ Treg cells and the effector CD62Llow CD4+ T cells. The former increased in ED-SCLC patients, and in contrast, the latter significantly increased without induction of Treg cells in LD-SCLC patients. Moreover, cytokine analyses revealed that the CD62Llow CD4+ T cells purified from LD-SCLC patients included more Th17 cells that produce preferentially IL-17 and that dendritic cells (DC) derived from CD14+ cells of LD-SCLC patients produced more IL-23.

Patients. The present study comprised 35 consecutive SCLC patients, 8 long-term survivors, and 19 healthy volunteers from a single institution (Niigata University Medical and Dental Hospital, Niigata, Japan; Table 1). LD-SCLC patients who had been disease-free for >3 y after treatment were considered to be long-term survivors. Specimens were collected after obtaining written informed consent approved by the Niigata University Ethical Committee.

Table 1.

Patient characteristics

StageAgeGender
Healthy volunteer (n = 19)   
 30 
 29 
 29 
 32 
 32 
 36 
 35 
 31 
 30 
 64 
 75 
 67 
 65 
 56 
 55 
 57 
 62 
 59 
 57 
Long-term survivors (n = 8)   
    cT1N2M0 51 
    cT4N2M0 62 
    cT2N1M0 59 
    cT4N2M0 64 
    cT4N3M0 70 
    pT1N0M0 80 
    cT2N2M0 68 
    pT1N0M0 68 
ED-SCLC (n = 15)   
    cT1N3M1 66 
    cT2N3M1 58 
    cT1N3M1 61 
    cT4N2M1 65 
    cT4N3M1 62 
    cT1N2M1 57 
    cT1N3M1 58 
    cT1N3M1 48 
    cT4N2M1 62 
    cT3N3M1 77 
    cT1N3M1 63 
    cT2N3M1 58 
    cT1N2M1 59 
    cT4N3M1 71 
    cT1N3M1 58 
LD-SCLC (n = 20)   
    cT2N2M0 72 
    cT4N2M0 57 
    cT4N3M0 55 
    cT1N1M0 71 
    cT1N3M0 62 
    cT4N0M0 56 
    cT3N2M0 63 
    cT2N3M0 67 
    cT2N0M0 64 
    cT4N3M0 63 
    cT1N2M0 66 
    pT2N0M0 54 
    cT1N2M0 69 
    cT4N3M0 65 
    cT1N0M0 57 
    cT1N1M0 63 
    cT4N3M0 71 
    cT1N2M0 75 
    cT1N1M0 67 
    cT4N3M0 63 
StageAgeGender
Healthy volunteer (n = 19)   
 30 
 29 
 29 
 32 
 32 
 36 
 35 
 31 
 30 
 64 
 75 
 67 
 65 
 56 
 55 
 57 
 62 
 59 
 57 
Long-term survivors (n = 8)   
    cT1N2M0 51 
    cT4N2M0 62 
    cT2N1M0 59 
    cT4N2M0 64 
    cT4N3M0 70 
    pT1N0M0 80 
    cT2N2M0 68 
    pT1N0M0 68 
ED-SCLC (n = 15)   
    cT1N3M1 66 
    cT2N3M1 58 
    cT1N3M1 61 
    cT4N2M1 65 
    cT4N3M1 62 
    cT1N2M1 57 
    cT1N3M1 58 
    cT1N3M1 48 
    cT4N2M1 62 
    cT3N3M1 77 
    cT1N3M1 63 
    cT2N3M1 58 
    cT1N2M1 59 
    cT4N3M1 71 
    cT1N3M1 58 
LD-SCLC (n = 20)   
    cT2N2M0 72 
    cT4N2M0 57 
    cT4N3M0 55 
    cT1N1M0 71 
    cT1N3M0 62 
    cT4N0M0 56 
    cT3N2M0 63 
    cT2N3M0 67 
    cT2N0M0 64 
    cT4N3M0 63 
    cT1N2M0 66 
    pT2N0M0 54 
    cT1N2M0 69 
    cT4N3M0 65 
    cT1N0M0 57 
    cT1N1M0 63 
    cT4N3M0 71 
    cT1N2M0 75 
    cT1N1M0 67 
    cT4N3M0 63 

Cell purification. PBMCs were obtained by centrifugation over Ficoll-Hypaque gradients. CD4+ and CD8+ T cells were purified by positive selection using CD4+ or CD8+ T-cell isolation kits (Dynal Biotech). CD4+ T cells were further separated into CD62Lhigh and CD62Llow cells. Cells were incubated with specific CD62L monoclonal antibody (mAb; 1H3) followed by a panning technique in T-25 flasks that were precoated with goat anti-mouse immunoglobulin antibodies (Jackson ImmunoResearch Laboratories). CD62Lhigh T cells adhered to the plastic surface and could be easily obtained by scraping with a rubber policeman. To obtain highly purified CD62Llow cells, the nonadherent cells were further depleted of CD62L-positive cells using anti-CD62L mAb and sheep anti-mouse immunoglobulin antibody-coated DynaBeads M-450 (Dynal Biotech) following the manufacturer's suggested procedure. CD25+ cells were isolated by positive selection using anti-CD25 mAb-coated microbeads and autoMACS (Miltenyi Biotec). Cell purities were all >90%.

Monocyte-derived DCs. DCs were generated from CD14+ monocytes derived from peripheral blood. Briefly, CD14+ cells were positively isolated using anti-CD14 mAb-coated microbeads (Miltenyi Biotec). The isolated CD14+ cells were then cultured in complete medium (CM) supplemented with 80 ng/mL of recombinant human granulocyte macrophage colony-stimulating factor (a gift from Kirin) and 10 ng/mL IL-4 (eBioscience). On day 6, the nonadherent DCs were harvested by gentle pipetting. CM consists of RPMI 1640 supplemented with 10% heat-inactivated lipopolysaccharide-qualified FCS, 0.1 mmol/L nonessential amino acids, 1 μmol/L sodium pyruvate, 100 units/mL penicillin, and 100 μg/mL streptomycin sulfate (all from Life Technologies, Inc.).

Antibodies and phenotype analyses. The following mAbs were used: FITC-conjugated anti-CD3 (HIT3a) and CD4 (RPA-T4), phycoerythrin (PE)-conjugated anti-CD8 (RPA-T8) and CD25 (M-A251), PE-Cy7–conjugated anti-CD25 (M-A251), PE-Cy5–conjugated anti-CD62L (Dreg 56; all from BD PharMingen), and FITC-conjugated anti-CD62L (Dreg 56; eBioscience). Cell surface phenotypes were analyzed by direct immunofluorescence staining of 1 × 106 cells with conjugated mAbs. In each sample, 10,000 cells were analyzed using a FACScan flow microfluorometer (Becton Dickinson) and CellQuest software. Staining of cytosolic FOXP3 was done using PE-anti-human FOXP3 mAb (PCH101) and Staining set (eBioscience), according to the manufacturer's instructions.

Cytokine analysis. Responder T cells were stimulated with immobilized anti-CD3 mAb or DCs pulsed with irradiated tumor cells for 48 h. The supernatants were harvested and assayed for cytokine concentrations by using a BD-Cytometric Bead Array (CBA; Becton Dickinson) or a quantitative “sandwich” enzyme immunoassay with an IFN-γ and an IL-17A ELISA kit (eBioscience), according to the manufacturer's instructions.

Responder 2 × 105 monocyte-derived DCs were cultured in the presence of 10 ng/mL lipopolysaccharide (Sigma-Aldrich) in 200 μL CM for 24 h. The amounts of IL-23 were determined by using ELISA kit (eBioscience), according to the manufacturer's instructions.

Proliferation assay. The isolated T cells (2 × 105) were stimulated in 200 μL CM for 24 h in 96-well plates precoated with anti-human CD3 (BD Biosciences). The ratio of CD62Llow to CD62Lhigh CD4+CD25+ T cells was 2:1. CD62Llow T cells were labeled with 5 μmol/L 5-(6)-carboxyfluorescein diacetate succinimidyl diester (CFSE; Molecular Probes, Inc.) in HBSS at 37°C for 15 min and washed twice before CD3 stimulation. After stimulation for 24 h, the cells were counted and washed twice in HBSS. The T cells were then cultured in CM supplemented with 10 units/mL recombinant human IL-2 (a gift from Shionogi) at a concentration of 1 × 105/mL. Three wells were analyzed under each condition.

Statistical analyses.P values were calculated by using two-sided Student's t test; P < 0.05 was considered to be statistically significant. For comparison of percentages of cells from the same patients before and after chemotherapy, a paired t test was used instead.

Shifted CD4+ regulatory and effector balance in patients with LD-SCLC and ED-SCLC. Before purification, PBMCs were analyzed for CD3, CD4, CD8, CD62L, and CD25 expression. Table 1 shows characteristics of the healthy volunteers and patients included in this study. As shown in Fig. 1A, the percentages of CD4+ T cells did not differ among the healthy volunteers and LD-SCLC and ED-SCLC patients. Figure 1B and C shows the percentages of CD4+ cells that were CD62Llow and CD62LhighCD25+ cells. Approximately 5% to 10% of CD4+ T cells belonged to the CD62LhighCD25+ subpopulation in the healthy volunteers (Fig. 1C). The healthy volunteers included individuals both younger than (<40 years) and as old as the SCLC patients. No differences were observed between them with regard to the percentages of CD62Llow and CD62LhighCD25+ cells. Consistent with previous studies that showed that the Treg cell subpopulation increases in cancer-bearing patients (1420), the CD62LhighCD25+ subpopulation significantly increased in ED-SCLC patients compared with the healthy volunteers (P = 0.0009). In contrast, the percentages of the CD62LhighCD25+ subpopulation in LD-SCLC patients showed no differences from those of the healthy volunteers. Further, a significantly larger CD62Llow T-cell subpopulation was induced in the LD-SCLC patients compared with that in the healthy volunteers or ED-SCLC patients (P = 0.000009 and 0.00003, respectively; Fig. 1C). No significant difference was observed in the percentages of CD62Llow CD4+ T cells between the healthy volunteers and ED-SCLC patients. To illustrate the difference in the CD4+ T-cell balance between the CD62Llow CD4+ T cells and CD62LhighCD25+ CD4+ cells, we calculated their ratio. As shown in Fig. 1D, a distinct CD62Llow T-cell–dominant CD4+ T-cell balance was induced in LD-SCLC patients, whereas a CD62LhighCD25+ CD4+ T-cell–dominant balance was observed in ED-SCLC patients. This study included four LEMS patients. All of them were LD-SCLC patients and exhibited effector-dominant CD4+ T-cell balance. In contrast to the percentages of CD4+ cells, the percentages of CD62Llow primed effector CD8+ T cells show no differences among LD-SCLC and ED-SCLC patients and healthy volunteers (Fig. 1E and F).

Fig. 1.

PBMCs were stained with FITC-conjugated anti-CD62L, PE-Cy5–conjugated anti-CD25, PE-conjugated anti-CD4, or PE-conjugated anti-CD8 mAbs. A and E, percentages of CD4+ and CD8+ cells in relation to the total number of PBMCs in the lymphocyte region in forward and side scatter. B and C, percentages of CD62Llow and CD62LhighCD25+ cells in relation to CD4+ cells in SCLC patients and healthy volunteers. D, ratio of CD62Llow CD4+ T cells to CD62LhighCD25+ CD4+ T cells. The ratios were calculated as follows: ratio = percentage of CD62Llow cells/percentage of CD62Lhigh CD25+ cells. F, percentages of CD62Llow in relation to CD8+ cells.

Fig. 1.

PBMCs were stained with FITC-conjugated anti-CD62L, PE-Cy5–conjugated anti-CD25, PE-conjugated anti-CD4, or PE-conjugated anti-CD8 mAbs. A and E, percentages of CD4+ and CD8+ cells in relation to the total number of PBMCs in the lymphocyte region in forward and side scatter. B and C, percentages of CD62Llow and CD62LhighCD25+ cells in relation to CD4+ cells in SCLC patients and healthy volunteers. D, ratio of CD62Llow CD4+ T cells to CD62LhighCD25+ CD4+ T cells. The ratios were calculated as follows: ratio = percentage of CD62Llow cells/percentage of CD62Lhigh CD25+ cells. F, percentages of CD62Llow in relation to CD8+ cells.

Close modal

CD62LhighCD25+ CD4+ T cells in SCLC patients expressed FOXP3 and possessed Treg cell functions.Figure 2A shows a representative expression pattern of CD62L and CD25 on CD4+ cells. The CD4+ cells were isolated from one of LD-SCLC patients. We defined zones A, B, or C as CD62Llow, CD62LhighCD25, and CD62LhighCD25+, respectively. Expression of FOXP3, which is considered as the master switch for Treg cells, was analyzed in CD4+ T cells. As shown in Fig. 2B, only the CD62LhighCD25+ CD4+ T cells expressed FOXP3. To determine whether these cells possessed regulatory properties, CD62Llow T cells were cocultured with CD62LhighCD25+ CD4+ T cells in the presence of CD3 stimulation. CD62Llow CD4+ T cells, CD62LhighCD25 CD4+ T cells, and CD62LhighCD25+ CD4+ T cells were purified from PBMCs with magnetic beads as described. Experiments were repeated with T cells isolated from LD-SCLC and ED-SCLC patients and healthy volunteers. Figure 2C and D shows the representative data obtained with T cells derived from an ED-SCLC patient. CD62LhighCD25+ CD4+ T cells inhibited cytokine production and the proliferation of CD62Llow CD4+ T cells. Thus, CD62LhighCD25+ CD4+ T cells represented Treg cells.

Fig. 2.

CD4+ T cells purified from PBMCs were stained with FITC-conjugated anti-CD62L, PE-Cy5–conjugated anti-CD25, and PE-conjugated anti-FOXP3 mAbs. A, CD62L and CD25 expression on the isolated CD4+ T cells. Zones A, B, and C were defined as CD62Llow, CD62LhighCD25, and CD62LhighCD25+, respectively. The percentages of CD62LhighCD25+ CD4+ T cells and CD62Llow CD4+ T cells in relation to the numbers of CD4+ cells were analyzed. B, cytosolic FOXP3 expression of gated CD62Llow, CD62LhighCD25, and CD62LhighCD25+ CD4+ cells. C, IFN-γ production by 1 × 105 CD62Llow CD4+ T cells in the presence or absence of 5 × 104 CD62LhighCD25+ CD4+ T cells in 200 μL CM in a 96-well plate for 48 h. D, changes in the CD62Llow CD4+ T-cell counts; 1 × 105 CD62Llow T cells stained with CFSE were stimulated overnight with immobilized anti-CD3 mAbs in 96-well plates in the presence or absence of 5 × 104 CD62LhighCD25+ CD4+ T cells. They were then cultured in CM supplemented with 10 units/mL of recombinant human IL-2 at a concentration of 1 × 105/mL for 48 h. The cells were counted and analyzed using a microfluorometer before and after culture. Fold expansion was calculated as follows: number of CSFE+ cells after the 48-h culture period/number of CFSE+ cells before the 48-h culture period.

Fig. 2.

CD4+ T cells purified from PBMCs were stained with FITC-conjugated anti-CD62L, PE-Cy5–conjugated anti-CD25, and PE-conjugated anti-FOXP3 mAbs. A, CD62L and CD25 expression on the isolated CD4+ T cells. Zones A, B, and C were defined as CD62Llow, CD62LhighCD25, and CD62LhighCD25+, respectively. The percentages of CD62LhighCD25+ CD4+ T cells and CD62Llow CD4+ T cells in relation to the numbers of CD4+ cells were analyzed. B, cytosolic FOXP3 expression of gated CD62Llow, CD62LhighCD25, and CD62LhighCD25+ CD4+ cells. C, IFN-γ production by 1 × 105 CD62Llow CD4+ T cells in the presence or absence of 5 × 104 CD62LhighCD25+ CD4+ T cells in 200 μL CM in a 96-well plate for 48 h. D, changes in the CD62Llow CD4+ T-cell counts; 1 × 105 CD62Llow T cells stained with CFSE were stimulated overnight with immobilized anti-CD3 mAbs in 96-well plates in the presence or absence of 5 × 104 CD62LhighCD25+ CD4+ T cells. They were then cultured in CM supplemented with 10 units/mL of recombinant human IL-2 at a concentration of 1 × 105/mL for 48 h. The cells were counted and analyzed using a microfluorometer before and after culture. Fold expansion was calculated as follows: number of CSFE+ cells after the 48-h culture period/number of CFSE+ cells before the 48-h culture period.

Close modal

Th1, Th2, and Th17 cells were in the CD62Llow CD4+ T-cell subpopulation. Next, we examined the cytokine production profile of each T-cell subpopulation. Sufficient number of purified T cells for cytokine analyses were obtained from 6 healthy volunteers, 10 LD-SCLC patients, and 7 ED-SCLC patients. Purified T cells were stimulated with immobilized anti-CD3 mAbs, and the supernatant was analyzed for IFN-γ, tumor necrosis factor-α, IL-2, IL-4, IL-5, and IL-17 by BD-CBA or ELISA according to the manufacturer's instructions. Figure 3A shows a representative cytokine production profile obtained from one of the LD-SCLC patients. CD62Llow CD4+ T cells secreted large amounts of Th1, Th2, and Th17 cytokines, such as IFN-γ, tumor necrosis factor-α, IL-4, IL-5, and IL-17, except for IL-2 that was mainly secreted by CD62LhighCD25 CD4+ T cells. Thus, the CD62Llow CD4+ T cells included Th1, Th2, and Th17 cells, and most CD62LhighCD25 CD4+ T cells were naive T cells.

Fig. 3.

In a 96-well plate, 2 × 105 CD62Llow CD4+ T cells, CD62LhighCD25 CD4+ T cells, or CD62LhighCD25+ CD4+ T cells isolated from 10 LD-SCLC, 7 ED-SCLC, and 6 healthy volunteers were stimulated for 48 h with immobilized anti-CD3 mAbs in 200 μL CM. The supernatant was examined for IFN-γ, tumor necrosis factor-α (TNFα), IL-2, IL-4, IL-5, and IL-17 secreted by fractionated CD4+ T cells using BD-CBA and ELISA. A, representative data obtained from one LD-SCLC patient. B and C, amounts of IL-17 and IFN-γ produced by CD62Llow CD4+ T cells obtained from LD-SCLC and ED-SCLC patients and healthy volunteers. D, amounts of IL-23 secreted in 200 μL CM by 2 × 105 monocyte-derived DCs on stimulation of 10 ng/mL lipopolysaccharide for 24 h. E, antigen-stimulated secretion of IFN-γ by CD4+ T cells or CD8+ T cells isolated from PBMCs of the SCLC patient p-T2N0M0 who had undergone surgical resection. In a 96-well plate, 1 × 105 CD62LhighCD25 CD4+ T cells, CD62Llow CD4+ T cells, or CD62Llow CD8+ T cells were stimulated for 48 h with 5 × 104 CD14+ cell-derived DCs in the presence or absence of 5 × 104 CD62LhighCD25+ CD4+ T cells in 200 μL CM. DCs were pulsed overnight with an equal number of 5,000 cGy–irradiated SCLC tumor cells, and they were purified with CD11c microbeads before coculture.

Fig. 3.

In a 96-well plate, 2 × 105 CD62Llow CD4+ T cells, CD62LhighCD25 CD4+ T cells, or CD62LhighCD25+ CD4+ T cells isolated from 10 LD-SCLC, 7 ED-SCLC, and 6 healthy volunteers were stimulated for 48 h with immobilized anti-CD3 mAbs in 200 μL CM. The supernatant was examined for IFN-γ, tumor necrosis factor-α (TNFα), IL-2, IL-4, IL-5, and IL-17 secreted by fractionated CD4+ T cells using BD-CBA and ELISA. A, representative data obtained from one LD-SCLC patient. B and C, amounts of IL-17 and IFN-γ produced by CD62Llow CD4+ T cells obtained from LD-SCLC and ED-SCLC patients and healthy volunteers. D, amounts of IL-23 secreted in 200 μL CM by 2 × 105 monocyte-derived DCs on stimulation of 10 ng/mL lipopolysaccharide for 24 h. E, antigen-stimulated secretion of IFN-γ by CD4+ T cells or CD8+ T cells isolated from PBMCs of the SCLC patient p-T2N0M0 who had undergone surgical resection. In a 96-well plate, 1 × 105 CD62LhighCD25 CD4+ T cells, CD62Llow CD4+ T cells, or CD62Llow CD8+ T cells were stimulated for 48 h with 5 × 104 CD14+ cell-derived DCs in the presence or absence of 5 × 104 CD62LhighCD25+ CD4+ T cells in 200 μL CM. DCs were pulsed overnight with an equal number of 5,000 cGy–irradiated SCLC tumor cells, and they were purified with CD11c microbeads before coculture.

Close modal

Greater IL-17 secretion by Teff cells from LD-SCLC and ED-SCLC patients. It has recently been described that a novel subset of CD4+ T cells producing IL-17A, IL-17F, and IL-22 (i.e., Th17 cells) is essential in several autoimmune diseases, such as multiple sclerosis (2128). Moreover, recently, it was shown that Th17 cells mediated potent antitumor reactivity to eradicate large established tumor (29). Th1 cells also have been considered to play an important role in antitumor immunity. To determine the size of the CD4+ T-cell subpopulation secreting IL-17, or IFN-γ, CD4+ T cells isolated from SCLC patients and healthy volunteers were stimulated with immobilized anti-CD3 mAb, and the supernatants were quantified for IL-17A and IFN-γ using ELISA. As shown in Fig. 3B, CD62Llow CD4+ T cells isolated from LD-SCLC patients produced significantly more IL-17A than those isolated from ED-SCLC and healthy volunteers (P = 0.012 and 0.004, respectively). In contrast, levels of IFN-γ produced by the same number of CD62Llow CD4+ T cells showed no difference among the LD-SCLC and ED-SCLC patients and healthy volunteers (Fig. 3C). Thus, it is likely that Teff cells induced in LD-SCLC patients deviated to Th17 cells.

IL-23 produced by DCs is believed to play a critical role in expanding a Th17 subpopulation (30). It was reported that DCs derived from monocytes of the multiple sclerosis patients, in whom Th17 cells are highly pathogenic and essential, produced more IL-23 (24). Then, we tested if DCs derived from CD14+ cells of the SCLC patients secrete IL-23. As shown in Fig. 3D, DCs derived from the LD-SCLC patients produced significantly more IL-23 on lipopolysaccharide stimulation.

Autologous tumor antigen–specific IFN-γ production by CD62Llow T cells derived from LD-SCLC patients. Next, to assess whether the CD62Llow subpopulation comprised T cells that recognize antigens on autologous SCLC tumor cells, we evaluated cytokine production by T cells isolated from a stage IB LD-SCLC patient who had undergone surgical resection and adjuvant chemotherapy. The resected tumor was digested with a mixture of 0.1% collagenase, 0.01% DNase, and 2.5 units/mL hyaluronidase (Sigma) for 3 hours at room temperature. Tumor cells were cryopreserved by supplementation with 5% DMSO and 6% hydroxyethyl starch cryoprotectant mixture (CP-1; Kyokuto) at −80°C according to the manufacturer's instructions until they were used for coculture. CD62Llow CD4+ T cells, CD62LhighCD25low CD4+ T cells, or CD62Llow CD8+ T cells isolated from the peripheral blood of the patient were stimulated with monocyte-derived DCs pulsed with 50 Gy–irradiated autologous tumor cells. One million monocyte-derived DCs were cocultured with 1 × 106 tumor cells in 2 mL of CM for 24 hours. CD11c+ cells were further purified with CD11c microbeads and autoMACS after coculture for T-cell stimulation. In a 96-well plate, 1 × 105 CD62LhighCD25 CD4+ T cells, CD62Llow CD4+ T cells, or CD62Llow CD8+ T cells were stimulated for 48 hours with 5 × 104 CD14+ cell-derived DCs in the presence or absence of 5 × 104 CD62LhighCD25+ CD4+ T cells in 200 μL CM and the supernatants were measured for IFN-γ by using ELISA. As shown in Fig. 3E, CD62Llow CD4+ T cells secreted IFN-γ specific to autologous tumor antigens. In contrast, the naive CD62LhighCD25 CD4+ T-cell subpopulation did not show tumor antigen–specific IFN-γ release. Moreover, the addition of CD62LhighCD25+ CD4+ T cells inhibited tumor-specific IFN-γ secretion. CD62Llow CD8+ T cells also showed antigen-specific IFN-γrelease.

Effect of chemotherapy on Teff to Treg cell ratio. It has been perceived that chemotherapy affects the general immune responsiveness in treated patients. However, recent studies indicated that certain cytotoxic agents may augment immune reactivities by reducing the number of Treg cells (3133). To address the effects of chemotherapy on CD4+ T-cell balance, T cells obtained from PBMCs were examined after two courses of platinum-based chemotherapy and analyzed using the paired Student's t test. Seven LD-SCLC and five ED-SCLC patients were included for the analyses. Eight patients were treated with cisplatin (CDDP) and etoposide; three patients, with carboplatin (CBDCA) and etoposide; and one patient, with CBDCA, ifosfamide, and etoposide. The peripheral blood was obtained from patients whose WBC count recovered to >3,000/μL 18 to 20 days after the last chemotherapy session. As shown in Fig. 4A, the percentages of CD62Llow cells from the total number of CD4+ T cells increased significantly (P = 0.003). In contrast, the percentage of Treg cells decreased (P = 0.003; Fig. 4B). Thus, platinum-based chemotherapy increased the Teff to Treg cell ratio (P = 0.004; Fig. 4C). It is probable that Treg cells is more sensitive to chemotherapeutic agents and that Teff cells possess a potent ability to proliferate. Thus, even in the ED-SCLC patients, the level of Treg cells significantly decreased and Teff cells readily proliferated after chemotherapy, resulting in an increase in the Teff to Treg cell ratio. During certain periods after chemotherapy, the LD-SCLC and the ED-SCLC patients exhibited high Teff to Treg cell ratio without significant differences.

Fig. 4.

Effect of chemotherapy on the CD62Llow CD4+ T cells and CD62LhighCD25+ CD4+ T cells was evaluated. The PBMCs were stained with FITC-conjugated anti-CD4, PE-conjugated anti-CD25, and PE-Cy5–conjugated anti-CD62L mAbs. A and B, percentages of CD62Llow and CD62LhighCD25+ cells in relation to the numbers of CD4+ cells before and after chemotherapy. All these cells were subjected to two courses of platinum-based chemotherapy. C, ratio of the percentage of CD62Llow CD4+ T cells to that of CD62LhighCD25+ CD4+ T cells. ○, LD-SCLC patients; •, ED-SCLC patients.

Fig. 4.

Effect of chemotherapy on the CD62Llow CD4+ T cells and CD62LhighCD25+ CD4+ T cells was evaluated. The PBMCs were stained with FITC-conjugated anti-CD4, PE-conjugated anti-CD25, and PE-Cy5–conjugated anti-CD62L mAbs. A and B, percentages of CD62Llow and CD62LhighCD25+ cells in relation to the numbers of CD4+ cells before and after chemotherapy. All these cells were subjected to two courses of platinum-based chemotherapy. C, ratio of the percentage of CD62Llow CD4+ T cells to that of CD62LhighCD25+ CD4+ T cells. ○, LD-SCLC patients; •, ED-SCLC patients.

Close modal

Effector CD4+ T cells dominant in long-term survivors. We analyzed whether the effector-dominant CD4+ T-cell balance was maintained in SCLC patients who had been cured by treatment. As shown in Fig. 5, long-term survivors still retained significantly greater levels of CD62Llow CD4+ Teff cells compared with healthy volunteers (P < 0.00001). Although long-term survivors exhibited significantly but slightly greater levels of Treg cells than those in healthy volunteers (P < 0.01), their effector-dominant CD4+ T-cell balance was the same as that in LD-SCLC patients before treatment (Figs. 1D and E and 5). In contrast, the percentages of CD4+ Teff cells in LD-SCLC patients with recurrent disease were as low as those in ED-SCLC patients, although they had been diagnosed with LD-SCLC before treatment. In Fig. 5, the number indicates that recurrent disease was confined to the LD state. These patients without distant, hematogenous disease still maintained relatively effector-dominant balance.

Fig. 5.

A and B, percentages of CD62Llow CD4+ and CD62LhighCD25+ CD4+ T cells in relation to the number of CD4+ cells in SCLC patients. C, ratio of CD62Llow to CD62LhighCD25+ T cells. LS, long-term survivors who had been disease-free for >3 y after treatment; RD, LD-SCLC patients who had recurrent disease after treatment. #, patients who had local recurrent diseases without distant, hematogenous metastases. *, P < 0.01; **, P < 0.05, compared with healthy volunteers.

Fig. 5.

A and B, percentages of CD62Llow CD4+ and CD62LhighCD25+ CD4+ T cells in relation to the number of CD4+ cells in SCLC patients. C, ratio of CD62Llow to CD62LhighCD25+ T cells. LS, long-term survivors who had been disease-free for >3 y after treatment; RD, LD-SCLC patients who had recurrent disease after treatment. #, patients who had local recurrent diseases without distant, hematogenous metastases. *, P < 0.01; **, P < 0.05, compared with healthy volunteers.

Close modal

In this study, we showed the presence of a distinct reciprocal balance between Treg and Teff cells in the peripheral blood of LD-SCLC and ED-SCLC patients. LD-SCLC patients exhibited a unique effector-dominant CD4+ T-cell balance. Moreover, Teff cells induced in LD-SCLC patients included a larger Th17 cell subpopulation. In contrast, ED-SCLC patients exhibited Treg cell–dominant balance. The manner in which the distinct CD4+ T-cell balance is generated in cancer-bearing hosts remains unclear. In several human malignancies, it was reported that Treg cells were induced in tumor microenvironments and tumor-draining lymph nodes (10, 14, 16, 17, 34, 35). Tumor cells themselves can convert immature DCs into transforming growth factor-β–secreting DCs that selectively induce Treg cells (36). Immature myeloid suppressor cells induced by the tumor are also thought to be responsible for Treg cell induction (37). We found that immature myeloid suppressor cells increased in ED-SCLC patients compared with healthy volunteers (data not shown). Thus, tumor cells tend to activate the peripheral tolerance machinery to induce Treg cells. Indeed, the number of Treg cells increased in the peripheral blood of ED-SCLC patients, who comprise 80% to 85% of SCLC patients.

The question arises why Teff cells were induced in the LD-SCLC patients. Mutated gene products can induce immunity by triggering the recognition of normally silent epitopes, including self and tumor-associated antigens (38). Subsequently, tumor cells themselves are likely to have the potential to induce immunity because they all possess gene mutations. It was shown that a certain type of mutation, such as truncation and amino acid substitutions that promote T helper cell responses, further enhanced immunity. Thus, one possible explanation is that LD-SCLC cells but not ED-SCLC cells have immunogenic gene mutations. However, this theory still cannot explain why Teff cells induced in LD-SCLC deviated to Th17 cells and how Treg cell generation was inhibited in LD-SCLC.

Th17 cells develop via a lineage distinct from the Th1 and Th2 lineages and play an essential role in a variety of autoimmune diseases, such as multiple sclerosis (2126, 28). It was shown that monocyte-derived DCs from multiple sclerosis patients produced more IL-23 and that expansion of the pathogenic Th17 subpopulation was driven by IL-23 (30, 39). We detected that monocyte-derived DCs from LD-SCLC produced more IL-23 than that from ED-SCLC patients. Thus, it is likely that DCs in LD-SCLC patients have tend to expand a Th17 subpopulation. Recently, it was shown that Th17 and Treg cells are reciprocally induced depending on the cytokine balance (40). Transforming growth factor-β alone induces Treg cells; however, the addition of IL-6 to transforming growth factor-β results in Th17 cell differentiation and inhibits Treg cell induction. It is known that Th17 cells secrete not only IL-17 but also IL-6. Thus, it is possible that deviation to Th17 inhibits Treg cell development and facilitates priming of Th17 as a positive feedback mechanism in LD-SCLC patients.

It remains uncertain if the effector-dominant CD4+ T-cell balance did in fact prevent distant metastases in LD-SCLC patients. In murine models, the adoptive transfer of tumor antigen–reactive effector CD62Llow CD4+ T cells tilted the Treg cell–dominant CD4+ T-cell balance toward a dominant Teff cell environment that resulted in the regression of the growing tumor (13). Moreover, adoptive transfer of Treg cells isolated from tumor-draining lymph nodes promoted tumor growth (11). Recently, it was reported that Th17-polarized cells mediated eradication of large established tumor and that IL-23 worked as a cancer vaccine adjuvant to increase antitumor T cells and enhance effector T-cell function (29, 41). Thus, it is likely that the effector-dominant CD4+ T-cell balance and Th17-polarized cells induced in LD-SCLC patients is not only a result but also plays a role in the defense mechanism against tumor cells. Consistent with this theory, the long-term survivors maintained their effector-dominant CD4+ T-cell balance, whereas LD-SCLC patients with recurrent disease due to distant metastases acquired Treg cell–dominant balance.

In contrast to CD4+ T cells, the size of the effector CD8+ T-cell population, which exhibited tumor antigen specificity, was not affected by disease stages or by distinct CD4+ T-cell balance in SCLC patients. Although it has been reported that Treg cells can suppress either CD4+ or CD8+ effector T-cell proliferation (4244), recent in vivo studies showed that antigen-specific CD8+ T cells undergo essentially normal clonal expansion and effector differentiation in the presence of antigen-specific Treg cells (4548). Our proliferation assays indicated that the Treg cells derived from SCLC patients delayed effector CD8+ T-cell proliferation but did not inhibit it (data not shown). Thus, Treg cells may suppress a terminal effector function but not the size of the effector CD8+ T-cell population (48).

The ratio of effector CD4+ cells in relation to Treg cells could be a useful biomarker for assessing immunologic responses and distinguishing LD-SCLC patients from ED-SCLC patients. Further, the data in this article shed light on the significance of IL-23/Th17 axis for antitumor immunity. Furthermore, these results indicated that immunotherapy that increases tumor-reactive Teff cell levels and depletes Treg cells, thereby maintaining a Th17 cell-dominant CD4+ T-cell balance, may be essential in establishing effective antitumor immunity.

No potential conflicts of interest were disclosed.

Grant support: Grant-in-Aid for Scientific Research from the Ministry of Education, Science and Culture of Japan; Niigata University Grant for Promotion of Project; and Niigata University Grant for Scientific Research.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Note: K. Koyama and H. Kagamu contributed equally to this work.

We thank Dr. Suyu Shu for reviewing the manuscript and helpful discussion.

1
Saito T, Kobayashi M, Harada R, Uemura Y, Taguchi H. Sensitive detection of small cell lung carcinoma cells by reverse transcriptase-polymerase chain reaction for prepro-gastrin-releasing peptide mRNA.
Cancer
2003
;
97
:
2504
–11.
2
Begueret H, Vergier B, Begueret J, et al. Detection of circulating cells expressing chromogranin A gene transcripts in patients with lung neuroendocrine carcinoma.
Eur J Cancer
2002
;
38
:
2325
–30.
3
Maddison P, Newsom-Davis J, Mills KR, Souhami RL. Favourable prognosis in Lambert-Eaton myasthenic syndrome and small-cell lung carcinoma.
Lancet
1999
;
353
:
117
–8.
4
Sakaguchi S, Sakaguchi N, Shimizu J, et al. Immunologic tolerance maintained by CD25+ CD4+ regulatory T cells: their common role in controlling autoimmunity, tumor immunity, and transplantation tolerance.
Immunol Rev
2001
;
182
:
18
–32.
5
Viguier M, Lemaitre F, Verola O, et al. Foxp3 expressing CD4+CD25(high) regulatory T cells are overrepresented in human metastatic melanoma lymph nodes and inhibit the function of infiltrating T cells.
J Immunol
2004
;
173
:
1444
–53.
6
Liyanage UK, Moore TT, Joo HG, et al. Prevalence of regulatory T cells is increased in peripheral blood and tumor microenvironment of patients with pancreas or breast adenocarcinoma.
J Immunol
2002
;
169
:
2756
–61.
7
Woo EY, Yeh H, Chu CS, et al. Cutting edge: regulatory T cells from lung cancer patients directly inhibit autologous T cell proliferation.
J Immunol
2002
;
168
:
4272
–6.
8
Sasada T, Kimura M, Yoshida Y, Kanai M, Takabayashi A. CD4+CD25+ regulatory T cells in patients with gastrointestinal malignancies: possible involvement of regulatory T cells in disease progression.
Cancer
2003
;
98
:
1089
–99.
9
Ichihara F, Kono K, Takahashi A, Kawaida H, Sugai H, Fujii H. Increased populations of regulatory T cells in peripheral blood and tumor-infiltrating lymphocytes in patients with gastric and esophageal cancers.
Clin Cancer Res
2003
;
9
:
4404
–8.
10
Curiel TJ, Coukos G, Zou L, et al. Specific recruitment of regulatory T cells in ovarian carcinoma fosters immune privilege and predicts reduced survival.
Nat Med
2004
;
10
:
942
–9.
11
Hiura T, Kagamu H, Miura S, et al. Both regulatory T cells and antitumor effector T cells are primed in the same draining lymph nodes during tumor progression.
J Immunol
2005
;
175
:
5058
–66.
12
Kagamu H, Touhalisky JE, Plautz GE, Krauss JC, Shu S. Isolation based on L-selectin expression of immune effector T cells derived from tumor-draining lymph nodes.
Cancer Res
1996
;
56
:
4338
–42.
13
Kagamu H, Shu S. Purification of L-selectin(low) cells promotes the generation of highly potent CD4 antitumor effector T lymphocytes.
J Immunol
1998
;
160
:
3444
–52.
14
Wolf AM, Wolf D, Steurer M, Gastl G, Gunsilius E, Grubeck-Loebenstein B. Increase of regulatory T cells in the peripheral blood of cancer patients.
Clin Cancer Res
2003
;
9
:
606
–12.
15
Barnett B, Kryczek I, Cheng P, Zou W, Curiel TJ. Regulatory T cells in ovarian cancer: biology and therapeutic potential.
Am J Reprod Immunol
2005
;
54
:
369
–77.
16
Miller AM, Lundberg K, Ozenci V, et al. CD4+CD25 high T cells are enriched in the tumor and peripheral blood of prostate cancer patients.
J Immunol
2006
;
177
:
7398
–405.
17
Petersen RP, Campa MJ, Sperlazza J, et al. Tumor infiltrating Foxp3+ regulatory T-cells are associated with recurrence in pathologic stage I NSCLC patients.
Cancer
2006
;
107
:
2866
–72.
18
Loddenkemper C, Schernus M, Noutsias M, Stein H, Thiel E, Nagorsen D. In situ analysis of FOXP3+ regulatory T cells in human colorectal cancer.
J Transl Med
2006
;
4
:
52
.
19
Liyanage UK, Goedegebuure PS, Moore TT, et al. Increased prevalence of regulatory T cells (Treg) is induced by pancreas adenocarcinoma.
J Immunother (1997)
2006
;
29
:
416
–24.
20
Nummer D, Suri-Payer E, Schmitz-Winnenthal H, et al. Role of tumor endothelium in CD4+ CD25+ regulatory T cell infiltration of human pancreatic carcinoma.
J Natl Cancer Inst
2007
;
99
:
1188
–99.
21
Amadi-Obi A, Yu CR, Liu X, et al. TH17 cells contribute to uveitis and scleritis and are expanded by IL-2 and inhibited by IL-27/STAT1.
Nat Med
2007
;
13
:
711
–8.
22
Bettelli E, Oukka M, Kuchroo VK. T(H)-17 cells in the circle of immunity and autoimmunity.
Nat Immunol
2007
;
8
:
345
–50.
23
Lohr J, Knoechel B, Wang JJ, Villarino AV, Abbas AK. Role of IL-17 and regulatory T lymphocytes in a systemic autoimmune disease.
J Exp Med
2006
;
203
:
2785
–91.
24
Vaknin-Dembinsky A, Balashov K, Weiner HL. IL-23 is increased in dendritic cells in multiple sclerosis and down-regulation of IL-23 by antisense oligos increases dendritic cell IL-10 production.
J Immunol
2006
;
176
:
7768
–74.
25
Toh ML, Miossec P. The role of T cells in rheumatoid arthritis: new subsets and new targets.
Curr Opin Rheumatol
2007
;
19
:
284
–8.
26
Harrington LE, Hatton RD, Mangan PR, et al. Interleukin 17-producing CD4+ effector T cells develop via a lineage distinct from the T helper type 1 and 2 lineages.
Nat Immunol
2005
;
6
:
1123
–32.
27
Chen Y, Langrish CL, McKenzie B, et al. Anti-IL-23 therapy inhibits multiple inflammatory pathways and ameliorates autoimmune encephalomyelitis.
J Clin Invest
2006
;
116
:
1317
–26.
28
Nakae S, Nambu A, Sudo K, Iwakura Y. Suppression of immune induction of collagen-induced arthritis in IL-17-deficient mice.
J Immunol
2003
;
171
:
6173
–7.
29
Muranski P, Boni A, Antony PA, et al. Tumor-specific Th17-polarized cells eradicate large established melanoma.
Blood
2008
;
112
:
362
–73.
30
Kikly K, Liu L, Na S, Sedgwick JD. The IL-23/Th(17) axis: therapeutic targets for autoimmune inflammation.
Curr Opin Immunol
2006
;
18
:
670
–5.
31
Beyer M, Kochanek M, Darabi K, et al. Reduced frequencies and suppressive function of CD4+CD25hi regulatory T cells in patients with chronic lymphocytic leukemia after therapy with fludarabine.
Blood
2005
;
106
:
2018
–25.
32
Taieb J, Chaput N, Schartz N, et al. Chemoimmunotherapy of tumors: cyclophosphamide synergizes with exosome based vaccines.
J Immunol
2006
;
176
:
2722
–9.
33
Su YC, Rolph MS, Cooley MA, Sewell WA. Cyclophosphamide augments inflammation by reducing immunosuppression in a mouse model of allergic airway disease.
J Allergy Clin Immunol
2006
;
117
:
635
–41.
34
Wolf D, Wolf AM, Rumpold H, et al. The expression of the regulatory T cell-specific forkhead box transcription factor FoxP3 is associated with poor prognosis in ovarian cancer.
Clin Cancer Res
2005
;
11
:
8326
–31.
35
Schaefer C, Kim GG, Albers A, Hoermann K, Myers EN, Whiteside TL. Characteristics of CD4+CD25+ regulatory T cells in the peripheral circulation of patients with head and neck cancer.
Br J Cancer
2005
;
92
:
913
–20.
36
Ghiringhelli F, Puig PE, Roux S, et al. Tumor cells convert immature myeloid dendritic cells into TGF-β-secreting cells inducing CD4+CD25+ regulatory T cell proliferation.
J Exp Med
2005
;
202
:
919
–29.
37
Huang B, Pan PY, Li Q, et al. Gr-1+CD115+ immature myeloid suppressor cells mediate the development of tumor-induced T regulatory cells and T-cell anergy in tumor-bearing host.
Cancer Res
2006
;
66
:
1123
–31.
38
Engelhorn ME, Guevara-Patino JA, Noffz G, et al. Autoimmunity and tumor immunity induced by immune responses to mutations in self.
Nat Med
2006
;
12
:
198
–206.
39
Langrish CL, Chen Y, Blumenschein WM, et al. IL-23 drives a pathogenic T cell population that induces autoimmune inflammation.
J Exp Med
2005
;
201
:
233
–40.
40
Bettelli E, Carrier Y, Gao W, et al. Reciprocal developmental pathways for the generation of pathogenic effector TH17 and regulatory T cells.
Nature
2006
;
441
:
235
–8.
41
Overwijk WW, de Visser KE, Tirion FH, et al. Immunological and antitumor effects of IL-23 as a cancer vaccine adjuvant.
J Immunol
2006
;
176
:
5213
–22.
42
Sakaguchi S, Sakaguchi N, Asano M, Itoh M, Toda M. Immunologic self-tolerance maintained by activated T cells expressing IL-2 receptor α-chains (CD25). Breakdown of a single mechanism of self-tolerance causes various autoimmune diseases.
J Immunol
1995
;
155
:
1151
–64.
43
Sakaguchi S. Naturally arising Foxp3-expressing CD25+CD4+ regulatory T cells in immunological tolerance to self and non-self.
Nat Immunol
2005
;
6
:
345
–52.
44
Shevach EM. CD4+ CD25+ suppressor T cells: more questions than answers.
Nat Rev Immunol
2002
;
2
:
389
–400.
45
Lin CY, Graca L, Cobbold SP, Waldmann H. Dominant transplantation tolerance impairs CD8+ T cell function but not expansion.
Nat Immunol
2002
;
3
:
1208
–13.
46
Klein L, Khazaie K, von Boehmer H. In vivo dynamics of antigen-specific regulatory T cells not predicted from behavior in vitro.
Proc Natl Acad Sci U S A
2003
;
100
:
8886
–91.
47
Chen ML, Pittet MJ, Gorelik L, et al. Regulatory T cells suppress tumor-specific CD8 T cell cytotoxicity through TGF-β signals in vivo.
Proc Natl Acad Sci U S A
2005
;
102
:
419
–24.
48
Mempel TR, Pittet MJ, Khazaie K, et al. Regulatory T cells reversibly suppress cytotoxic T cell function independent of effector differentiation.
Immunity
2006
;
25
:
129
–41.