Defects in the CD3/TCR complex and impairment of T cell function are necessary for tumor evasion, but the underlying mechanisms are incompletely understood. We found that culture supernatants from several types of solid tumor cell lines drove human monocytes to become tolerogenic semimature dendritic cells (TDCs). Upon encountering T cells, the TDCs triggered rapid down-regulation of CD3ε and TCR-α/β and subsequent apoptosis in autologous T cells. Consistent with these results, accumulation of immunosuppressive DCs coincided with CD3ε down-regulation and T cell deletion in cancer nests of human tumors. The impaired T cell function was mediated by factor(s) released by live TDCs after direct interaction with lymphocytes. Also, the TDC-induced effect on T cells was markedly reduced by blocking of NADPH oxidase but not by inhibition of arginase, inducible NO synthase (iNOS), IDO, or IFN-γ. Moreover, we found that hyaluronan fragments constituted a common factor produced by a variety of human tumor cell lines to induce formation of TDCs. These observations indicate that tumor microenvironments, including hyaluronan fragments derived from cancer cells, educate DCs to adopt a semimature phenotype, which in turn aids tumor immune escape by causing defects in the CD3/TCR complex and deletion of T cells.

Dendritic cells (DCs)3 are the most potent “professional” APCs, and they are responsible for integrating a variety of incoming signals and orchestrating the immune response (1). Bidirectional interactions between DCs and T cells initiate either an immunogenic or a tolerogenic pathway, both of which play crucial roles in autoimmune diseases and tumor immunity (2, 3). It is generally assumed that these two contrasting functions of DCs are associated with the maturation stages of the cells: fully mature DCs (mDCs) are efficient activators of naive T cells, whereas immature DCs (iDCs) have been implicated for anergy induction (4, 5). Furthermore, an intermediate stage of maturation was recently described in which the cells are referred to as semimature DCs (6). These DCs expressed high levels of MHC class II and costimulatory molecules, even though they exhibited an IL-12lowIL-10high phenotype. It was also observed that the semimature DCs can induce tolerance by generating regulatory T cells and/or T cell anergy (6).

Tumors can mimic some of the signaling pathways of the immune system to propagate conditions that favor immune tolerance and so escape tumor immunity (7). Clinical and experimental studies have demonstrated that tumor growth is closely associated with impaired differentiation and maturation of myeloid cells, particularly macrophages and DCs (8, 9, 10). Also, phenotypic and functional analyses of myeloid cells from cancer patients have revealed that tumor cells or tumor-derived factors do favor differentiation of monocytes into tolerogenic DCs (11, 12). Although the exact mechanisms underlying this effect are not yet clear, such abnormal development of DCs in the tumor microenvironment may contribute to the impact of these cells on the dysfunction or signaling of T cells.

TCRs are essential for the functions of lymphocytes, and thus defects in proximal TCR signaling have been recognized as important mechanisms to evade immune responses (7, 13). An impairment of TCR signaling that inactivates the effector phase of the antitumor response has been observed in both mice and patients with advanced tumors (14). Furthermore, evidence is emerging that expression of the CD3ζ chain is markedly decreased in both peripheral-blood and tumor-infiltrating lymphocytes in patients with different types of tumors (15, 16, 17). Compared with the ζ-chain, significantly less is known about the importance of the ε chain in tumor immunity. CD3ε is represented twice in the TCR complex, serving as a component of both the γ/ε and δ/ε dimers, both of which are essential for T cell survival (13, 18). CD3ε deficiency has been shown to represent a novel cause in patients with severe combined immunodeficiency (19). These mentioned findings, together with the recent reported down-regulation of CD3ε in patients with lung cancer (20), imply that CD3ε may play an important role in tumor immune escape.

In the present study, we observed that soluble factors derived from tumor cell lines drove human monocytes to develop into tolerogenic semimature DCs (TDCs). Upon encountering T cells, these TDCs impaired the expression of CD3ε and subsequently induced apoptosis in autologous T cells through generation of reactive oxygen species (ROS). This finding, along with the distinct expression patterns of CD3ε molecules in tumor samples, provides important insight into the collaborative action that occurs in tumor environments to counteract effective immune responses by blocking the TCR signaling for T cell survival. Our results also suggest that hyaluronan (HA) fragments represent a common factor that is released by tumor cells to induce the formation of TDCs.

The Abs and chemicals used and their sources were as follows: recombinant human GM-CSF, IL-4, an annexin V apoptosis detection kit, and blocking mAbs against human IFN-γ, TNF-α, and IL-10 were purchased from R&D Systems; anti-CD44 Ab and a control Ab were from Lab Vision; and cell isolation and tissue culture reagents were obtained from Invitrogen. The HA-specific blocking peptide Pep-1 (GAHWQFNALTVR) and a control peptide (WRHGFALTAVNQ) were synthesized by GL Biochem as described previously (21), and they were purified to >98% by reverse-phase HPLC. All other reagents were obtained from Sigma-Aldrich unless otherwise indicated in the text.

Human cervical (HeLa), hepatoma (SK-Hep-1 and HepG2), and leukemia (THP1) cell lines were obtained from American Type Culture Collection; cells of the lung carcinoma line 95D were from the Institute of Biochemistry and Cell Biology, Chinese Academy of Sciences; the stable HAS2-knockdown SK-Hep-1 clones (SiHAS2) and the mock transfectants were established in our previous study (22, 23). All cells were tested for mycoplasma contamination using a single-step PCR method (24), and they were maintained in complete medium composed of RPMI 1640 with 10% FBS. To prepare tumor cell supernatants (TSNs), 5 × 106 tumor cells were plated in 10 ml of complete medium in 100-mm dishes for 3–4 days, and the supernatants were subsequently centrifuged, filtered, and stored in aliquots at −80°C.

Intermediate-sized HA fragments (50–200 kDa) were prepared by partial digestion of high-molecular mass HA with limited amounts of testicular hyaluronidase as previously described (22). In short, high-molecular mass HA from rooster comb (H1751) was dissolved in digestion buffer (10 mM CH3COONa (pH 4.0), 500 U/ml hyaluronidase from bovine testes, and 5 mg of HA/ml) and incubated at 37°C for 30 min. The reaction was terminated by putting the digestion mixture into a boiling water bath. The sizes of HA fragments were determined by 0.5% agarose gel electrophoresis and visualized with cationic dye Stain-All.

PBMCs were isolated from buffy coats of blood from healthy donors by Ficoll density gradient centrifugation, as previously described (25). Monocytes were selected from PBMCs by anti-CD14 magnetic beads in a MACS column purification system (Miltenyi Biotec). To generate DCs, the purified monocytes were cultured for 6–7 days in complete RPMI medium supplemented with 40 ng/ml GM-CSF and IL-4 in the presence of HA, or 20% TSNs. Half of the culture medium was replaced on day 3 and day 5. DC maturation was induced by stimulating the cells with 200 ng/ml LPS for 24 h. In some experiments, the monocytes were pretreated with the CD44-blocking Ab, control IgG, Pep-1, or a control peptide at the indicated concentrations before exposure to TSNs.

DCs were collected, washed, and then resuspended in PBS supplemented with 1% heat-inactivated FBS. Thereafter, the cells were stained with fluorochrome-conjugated mAbs against CD14, CD1a, CD86, and HLA-DR, or a relevant control Ab (BD Pharmingen) and then analyzed by flow cytometry using CellQuest software version 7.5.3 (FACSVantage-SE, BD Immunocytometry Systems).

Concentrations of TNF-α, IL-12p70, IL-10, and IFN-γ were determined using ELISA kits (eBioscience).

The autostimulatory function of DCs was examined by analyzing a MLR using CD14+-depleted PBMCs as responder cells. The responder cells were resuspended (4 × 106/ml) in RPMI 1640 medium supplemented with 10% heat-inactivated FBS and 20 IU/ml IL-2 (PeproTech), and then cultured in 96-well round-bottom plates (4 × 105 cells/well). Autostimulatory DCs resuspended (2 × 105/ml) in the same medium were incubated with PBMCs at a ratio of 1:20. In some experiments, before coculturing the DCs were pretreated with mitomycin C, diphenyleneiodonium (DPI), catalase, superoxide dismutase (SOD), 1-methyl-tryptophan (1-MT), N-monomethyl-l -arginine, or N-hydroxy-nor-l -arginine (Calbiochem), or with specific blocking mAbs against TNF-α, IFN-γ, and IL-10 at the indicated concentrations.

Lymphocytes were washed and resuspended in PBS supplemented with 1% heat-inactivated FBS. Thereafter, the cells were stained with fluorochrome-conjugated mAbs against CD3ε, CD3ζ, CD4, CD8, CD69, TCR-α/β, and control Ab (BD Pharmingen) according to the manufacturer’s instructions, and they were subsequently analyzed by flow cytometry. Apoptosis was quantified with an annexin V apoptosis detection kit (26), using CD3ε to set the gate for T cells. Binding of annexin V and CD3ε to the cells was measured by flow cytometry using CellQuest software.

A total of 44 samples of hepatocellular, cervical, colorectal, and lung carcinomas were obtained from the Cancer Center of Sun Yat-Sen University and anonymized in accordance with local ethical guidelines, as stipulated by the Declaration of Helsinki and a protocol approved by the Review Board of our Cancer Center. Paraffin-embedded and formalin-fixed samples were cut into 5-μm sections, which were then processed for immunohistochemistry as previously described (22). Following incubation with the Ab against human CD3ε or CD8α (DakoCytomation), the adjacent sections were stained using the EnVision System with diaminobenzidine or amino-ethylcarbazide (DakoCytomation). Evaluation of immunohistochemical variables was performed by two independent observers. At low-power field (×100), the tissue sections were screened and the five most representative areas were manually selected using a Leica DM IRB inverted microscope. For evaluating the density of CD8 cells in different regions, the respective areas of cancer nests and adjacent peritumoral stroma were measured at high-power field (×400, ∼0.15 mm2/field). The number of CD8 cells was then counted manually and expressed as cells per field.

Frozen sections (15 for hepatocellular carcinomas and 5 for lung cancers) were stained with polyclonal rabbit anti-human DC-SIGN and mouse anti-human IL-10 (R&D Systems), rabbit anti-human CD3ε and mouse anti-human CD8α (Lab Vision), or rabbit anti-human DC-SIGN and mouse anti-human CD8α followed by Alexa Fluor 488-, 568-, or 633-conjugated goat anti-mouse IgG (H+L) and Alexa Fluor 488- or 568-conjugated goat anti-rabbit IgG (H+L). Positive cells were quantified by ImagePro Plus software and are expressed as the mean of the percentage of positive cells (±SD) in 10 high-powered fields at × 800 magnification using confocal microscopy.

Apoptotic cells in paraffin-embedded human tumor samples were examined with the In Situ Cell Death Detection kit, POD (Roche Diagnostics, Pleasanton, CA) according to the instructions provided by the manufacturer. In some experiments, after TUNEL staining, the frozen tissue sections were reincubated with rabbit anti-human CD3ε followed by Alexa Fluor 568-conjugated goat anti-rabbit IgG (H+L).

The data on cytokine concentrations and surface marker expression are given as means ± SEM and SD, respectively. Statistical significance was determined by Student’s t test. A value of p < 0.05 was considered statistically significant.

To study the mechanisms involved in the induction of tolerogenic DCs by the tumor environment, we first set out to establish conditions under which this process can be reliably reproduced in vitro. Monocytes were cultured with GM-CSF and IL-4 in the presence or absence of TSNs for 6 days. By day 6, the control cells had differentiated into iDCs with reduced CD14 and increased CD1a on their surface. Exposure of monocytes to 20% TSNs from SK-Hep-1, HepG2, HeLa, and 95D cells, respectively, resulted in impaired differentiation of DCs with retained CD14 and reduced CD1a (Fig. 1,A). These DCs exhibited a semimature phenotype with a 2- to 5-fold increase in expression of CD83, CD86, and HLA-DR, and a distinctive IL-12lowIL-10high cytokine production profile (Fig. 1, A and B).

FIGURE 1.

Exposure to TSNs redirected monocytes to develop into tolerogenic semimature DCs. Purified monocytes were cultured for 6 days with GM-CSF and IL-4 in the presence (dashed lines) or absence (solid lines) of 20% TSNs from various tumor cell lines. A and B, Expression of surface markers and production of cytokines were determined by flow cytometry and ELISA, respectively. C and D, Thereafter, the DCs were stimulated with LPS for 24 h and were subsequently analyzed. The histograms in A and C are representative of 10 separate experiments. The data on cytokine production represent the means ± SE of 10 experiments; ∗∗, p < 0.01 compared with DCs that were cultured in medium alone.

FIGURE 1.

Exposure to TSNs redirected monocytes to develop into tolerogenic semimature DCs. Purified monocytes were cultured for 6 days with GM-CSF and IL-4 in the presence (dashed lines) or absence (solid lines) of 20% TSNs from various tumor cell lines. A and B, Expression of surface markers and production of cytokines were determined by flow cytometry and ELISA, respectively. C and D, Thereafter, the DCs were stimulated with LPS for 24 h and were subsequently analyzed. The histograms in A and C are representative of 10 separate experiments. The data on cytokine production represent the means ± SE of 10 experiments; ∗∗, p < 0.01 compared with DCs that were cultured in medium alone.

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To further elucidate the functional states of TSN-exposed DCs, we next examined how they responded to LPS. In normal iDCs, this stimulus induced full maturation involving events such as up-regulation of CD83, CD86, and HLA-DR, and production of IL-12p70 and TNF-α (27). However, with the exception of augmented generation of IL-10, that pattern was significantly attenuated in DCs that had been exposed to TSNs from solid tumor cell lines (Fig. 1, C and D). In contrast, TSNs from leukemia (THP1) and normal liver (LO2) cells had only marginal effects on the cytokine production profile and the surface marker expression in both immature and mature DCs (Fig. 1, B and D; data not shown), even when used at a high concentration (40% TSNs). These results clearly indicate that soluble mediators released from solid tumor cell lines compel monocytes to develop into tolerogenic semimature DCs. The TSNs used in this study did not contain any measurable levels of TNF-α, IL-12p70, or IL-10, and thus the cytokines we detected must have been produced by DCs.

The results regarding tumor-induced DC dysfunction were further confirmed in human tumor samples stained for DC-SIGN (marker for DC) (28) and IL-10. In hepatocellular carcinomas, DC-SIGN-positive cells accumulated in the cancer nest, whereas they were hardly detected in adjacent normal tissue (Fig. 2,A). Furthermore, 65 ± 12% of the DC-SIGN-positive cells (23 ± 5 cells/field, n = 4) in cancer nests were also positive for IL-10 (Fig. 2 A), which indicates that they may have an immunosuppressive phenotype in situ. Similar results were obtained in lung cancer tissues (data not shown).

FIGURE 2.

Localization and phenotype of tumor-infiltrating leukocytes in adjacent sections. A, Frozen sections of human hepatocellular carcinoma tissue and peritumor normal tissue were stained for DC-SIGN (green) and IL-10 (blue). B, Frozen sections of human hepatocellular carcinoma tissue were stained for CD3ε (green) and CD8α (red). Note that CD8+ cells in the cancer nests do express CD3ε, but at much lower levels. C, Frozen sections of hepatocellular carcinoma tissue were stained for DC-SIGN (red) and CD8α (green). Contact of CD8+ cells and DC-SIGN+ cells in tumor in situ could be identified (right panels). One of 15 representative patient samples is shown in A and B, and 1 of 8 representative samples is shown in C.

FIGURE 2.

Localization and phenotype of tumor-infiltrating leukocytes in adjacent sections. A, Frozen sections of human hepatocellular carcinoma tissue and peritumor normal tissue were stained for DC-SIGN (green) and IL-10 (blue). B, Frozen sections of human hepatocellular carcinoma tissue were stained for CD3ε (green) and CD8α (red). Note that CD8+ cells in the cancer nests do express CD3ε, but at much lower levels. C, Frozen sections of hepatocellular carcinoma tissue were stained for DC-SIGN (red) and CD8α (green). Contact of CD8+ cells and DC-SIGN+ cells in tumor in situ could be identified (right panels). One of 15 representative patient samples is shown in A and B, and 1 of 8 representative samples is shown in C.

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Previous studies have shown that, depending on their phenotype, DCs can induce either tolerance or activation in T cells (4, 5). Therefore, we investigated the effect of TSN-exposed semimature DCs (TDCs) on T cell responses in a MLR assay using CD14+-depleted autologous PBMCs as responders. The results showed that 24-h coculturing with TDCs, but not with iDCs or mDCs, significantly decreased the extracellular expression of the TCR/CD3 complex, including CD3ε and TCR-α/β chains, but had no effect on the intracellular expression of CD3ζ in T cells (Fig. 3,A and Table I). Similar results were obtained when we used purified CD4+ or CD8+ cells as responders in the MLR assay (data not shown).

FIGURE 3.

TSN-exposed DCs induced down-regulation of CD3ε and subsequent deletion of autologous T cells. CD14+-depleted PBMCs were cocultured (20:1) with normal iDCs (solid lines), normal mDCs (bold lines), or TSN-exposed DCs (dashed lines) for the indicated times. A, Expression of CD3ε, CD3ζ, and TCR-α/β in T cells was determined after 24 h by flow cytometry. B, The kinetic rate of apoptosis in the T cells was quantified by flow cytometry performed using an annexin V kit. C, Expression of CD3ε and binding of annexin V in T cells were analyzed by flow cytometry after 48 h. D, Apoptosis in CD4 or CD8 T cells was analyzed by multicolor flow cytometry. The results shown are representative of at least five independent experiments. The mean fluorescence intensity (MFI) of these molecules (mean ± SD) are indicated in Table I.

FIGURE 3.

TSN-exposed DCs induced down-regulation of CD3ε and subsequent deletion of autologous T cells. CD14+-depleted PBMCs were cocultured (20:1) with normal iDCs (solid lines), normal mDCs (bold lines), or TSN-exposed DCs (dashed lines) for the indicated times. A, Expression of CD3ε, CD3ζ, and TCR-α/β in T cells was determined after 24 h by flow cytometry. B, The kinetic rate of apoptosis in the T cells was quantified by flow cytometry performed using an annexin V kit. C, Expression of CD3ε and binding of annexin V in T cells were analyzed by flow cytometry after 48 h. D, Apoptosis in CD4 or CD8 T cells was analyzed by multicolor flow cytometry. The results shown are representative of at least five independent experiments. The mean fluorescence intensity (MFI) of these molecules (mean ± SD) are indicated in Table I.

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Table I.

TSN-exposed DCs impaired the expression of TCR-α/β and CD3ε in T cellsa

TreatmentTCR-α/βCD3εCD3ζ
iDCs 117 ± 21 411 ± 29 308 ± 26 
mDCs 108 ± 15 401 ± 37 300 ± 36 
SK-Hep-1 65 ± 13* 179 ± 21* 317 ± 28 
HepG2 58 ± 17* 156 ± 25* 299 ± 26 
HeLa 34 ± 11* 101 ± 18* 333 ± 21 
95D 29 ± 10* 89 ± 17* 301 ± 28 
TreatmentTCR-α/βCD3εCD3ζ
iDCs 117 ± 21 411 ± 29 308 ± 26 
mDCs 108 ± 15 401 ± 37 300 ± 36 
SK-Hep-1 65 ± 13* 179 ± 21* 317 ± 28 
HepG2 58 ± 17* 156 ± 25* 299 ± 26 
HeLa 34 ± 11* 101 ± 18* 333 ± 21 
95D 29 ± 10* 89 ± 17* 301 ± 28 
a

CD14+-depleted PBMCs were cocultured (20:1) with normal iDCs, mDCs, or TSN-exposed DCs for 24 h. Thereafter, the autologous T cells were collected and subsequently stained with anti-TCR-α/β, anti-CD3ε, anti-CD3ζ Ab, or relevant control Abs, and analyzed by multicolor flow cytometry. The data shown are the mean fluorescence intensities, and each value represents the mean ± SD of results from at least four separate experiments; ∗, p < 0.01, indicates significantly different from normal iDCs.

The signals transduced by extracellular interaction with the TCR/CD3 complex are essential for T cell survival and proliferation (18, 29). Accordingly, we next determined whether down-regulation of CD3ε and TCR-α/β in T cells would lead to their apoptosis as measured as annexin V binding by the gated proportion of lymphocytes. As illustrated in Fig. 3,B, T cells cocultured with TDCs, but not with control iDCs or mDCs, underwent apoptosis in a time-dependent manner, with annexin V positivity ranging from ∼60% to >90% after 72 h. Kinetic experiments using double staining revealed that the TDC-exposed T cells exhibited down-regulation of CD3ε much sooner than they showed when binding to annexin V, and that all of the apoptotic lymphocytes displayed attenuated expression of CD3ε (Fig. 3,C and data not shown). These results suggest that down-regulation of CD3ε was initiated before apoptosis in TDC-exposed T cells. Moreover, we found that both CD4 and CD8 cells were equally susceptible to apoptosis after coculture with TDCs (Fig. 3 D).

The results described above suggested that tumor microenvironments induce the formation of tolerogenic DCs, which in turn leads to down-regulation of CD3ε and subsequent deletion of T cells in cancer nests. To test this hypothesis, we examined the expression of CD3ε in serial sections of 24 human cancer specimens, including hepatocellular, cervical, colorectal, and lung carcinomas. In all samples analyzed, T cells were present throughout the tissue, but often predominantly in the peritumoral stroma surrounding the cancer nests. The T cells in peritumoral stroma showed marked expression of CD3ε, whereas those in the cancer nests exhibited significantly reduced levels of that protein (Figs. 2,B and 4,A). As illustrated in Fig. 4,B, a weak signal for CD3ε expression in the cancer nest was detected after a prolonged developing process. Similar results were obtained in complementary experiments performed to examine the expression of CD8α in adjacent sections of tumor samples, and those findings included significantly reduced numbers of CD8+ cells (cancer nests, 19 ± 10 cells/field vs peritumoral stroma, 116 ± 45 cells/field; n = 10; p < 0.001) with attenuated CD3ε expression in the cancer nests (Figs. 2, B and C, and 4,A). However, the expression of CD8α protein on each CD8+ cell seems unaffected, indicating that the down-regulation of CD3ε is a selective event for T cells in cancer nests (Fig. 4 A).

FIGURE 4.

Expression patterns of CD3ε and apoptotic status of T cells in human tumor samples. A, Adjacent sections of paraffin-embedded hepatocellular, cervical, or colorectal carcinomas were stained with an anti-CD3ε or anti-CD8α Ab. The micrographs at higher magnification show the stained cancer nest (1 ) and peritumoral stroma region (2 ). B, Expression of CD3ε in hepatocellular carcinomas after a prolonged developing process. C and D, Apoptosis of tumor-infiltrating lymphocytes in hepatoma sample. Adjacent sections of paraffin-embedded hepatoma samples were stained with an anti-CD3ε, anti-CD8α Ab, or TUNEL assay (C). D, Frozen tissue sections were stained for TUNEL assay (green) and with an anti-CD3ε Ab (red) and examined by confocal microscope. The micrograph shows the presence of TUNEL+ cells in the cancer nests, with all of them exhibiting an attenuated CD3ε expression (inset). Three of 24 representative patient samples are shown in A, and 1 of 10 representative samples is shown in C and D.

FIGURE 4.

Expression patterns of CD3ε and apoptotic status of T cells in human tumor samples. A, Adjacent sections of paraffin-embedded hepatocellular, cervical, or colorectal carcinomas were stained with an anti-CD3ε or anti-CD8α Ab. The micrographs at higher magnification show the stained cancer nest (1 ) and peritumoral stroma region (2 ). B, Expression of CD3ε in hepatocellular carcinomas after a prolonged developing process. C and D, Apoptosis of tumor-infiltrating lymphocytes in hepatoma sample. Adjacent sections of paraffin-embedded hepatoma samples were stained with an anti-CD3ε, anti-CD8α Ab, or TUNEL assay (C). D, Frozen tissue sections were stained for TUNEL assay (green) and with an anti-CD3ε Ab (red) and examined by confocal microscope. The micrograph shows the presence of TUNEL+ cells in the cancer nests, with all of them exhibiting an attenuated CD3ε expression (inset). Three of 24 representative patient samples are shown in A, and 1 of 10 representative samples is shown in C and D.

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We next used TUNEL assay to examined the apoptotic status of tumor-infiltrating T cells. As shown in Fig. 4,C, the cancer nests contain more TUNEL+ lymphocytes. The presence of more TUNEL+ T cells in the cancer nests in situ was further confirmed by confocal microscopic analysis on frozen tumor tissues. The numbers of CD3 cells in the intratumor and peritumoral stroma areas are 23 ± 8 and 97 ± 23 cells/field (×800 magnification, n = 4, p < 0.001), respectively. A significantly higher apoptotic rate of CD3 cells is found in the intratumoral area (27.9 ± 9.6% vs 2.1 ± 0.9%; n = 4; p < 0.001). Consistent with the results from in vitro study, the TUNEL+ T cells exhibited a significantly attenuated CD3ε expression as compared with TUNEL T cells (Fig. 4,D). Moreover, we also observed that 32.1 ± 7.3% of CD8+ cells (n = 4) had physical interaction with DC in the cancer nests (Fig. 2 C).

It is known that DCs regulate T cell responses via membrane-bound molecules and secretion of soluble mediators (1). Accordingly, we performed a series of experiments to investigate the mechanism by which TDCs induce down-regulation of CD3ε and apoptosis in T cells. Initially, TSN-induced semimature DCs (prepared as described above) were pretreated with mitomycin C or fixed with polyformaldehyde and subsequently cocultured with autologous T cells. Such treatment completely blocked the ability of TDCs to induce T cell dysfunction, as shown for mitomycin C-treated TDCs in Fig. 5,A. We obtained similar results when we cocultured TDCs and lymphocytes in different chambers of the transwell plates (Fig. 5 B), which suggests that the membrane-bound molecules or soluble mediators released solely by TDCs are not sufficient to trigger the above-mentioned T cell dysfunction.

FIGURE 5.

TSN-exposed DCs caused T cell dysfunction via a bidirectional cell-contact mechanism. A, CD14+-depleted PBMCs were cultured alone (Lym) or with (20:1) normal iDCs, HeLa-TDCs (TDCs), or mitomycin C-treated TDCs (MMC-TDCs) for 24 h, after which levels of expression of CD3ε and apoptosis in T cells were determined by flow cytometry. B, CD14+-depleted PBMCs were cultured alone (Lym) or with mitomycin C-treated TDCs (Lym + MMC-TDC) in the lower assay chamber of 24-well transwell plates (0.22 μm pore size; Costar), and with live TDCs or live TDCs plus lymphocytes in the upper chamber, as indicated. After 24 h, the cells in the lower chamber were collected for analysis. C and D, The CD14+-depleted PBMCs were left untreated or were cultured together with normal iDCs, TDCs, or MMC-TDCs for the indicated times. Thereafter, the expression of surface markers (after 24 h) and production of cytokines were determined by flow cytometry and ELISA, respectively. The results shown in A, B, and C are representative of four separate experiments. The data on cytokine production represent the means ± SE of four experiments; ∗, p < 0.05 and ∗∗, p < 0.01 indicate significantly different from TDCs.

FIGURE 5.

TSN-exposed DCs caused T cell dysfunction via a bidirectional cell-contact mechanism. A, CD14+-depleted PBMCs were cultured alone (Lym) or with (20:1) normal iDCs, HeLa-TDCs (TDCs), or mitomycin C-treated TDCs (MMC-TDCs) for 24 h, after which levels of expression of CD3ε and apoptosis in T cells were determined by flow cytometry. B, CD14+-depleted PBMCs were cultured alone (Lym) or with mitomycin C-treated TDCs (Lym + MMC-TDC) in the lower assay chamber of 24-well transwell plates (0.22 μm pore size; Costar), and with live TDCs or live TDCs plus lymphocytes in the upper chamber, as indicated. After 24 h, the cells in the lower chamber were collected for analysis. C and D, The CD14+-depleted PBMCs were left untreated or were cultured together with normal iDCs, TDCs, or MMC-TDCs for the indicated times. Thereafter, the expression of surface markers (after 24 h) and production of cytokines were determined by flow cytometry and ELISA, respectively. The results shown in A, B, and C are representative of four separate experiments. The data on cytokine production represent the means ± SE of four experiments; ∗, p < 0.05 and ∗∗, p < 0.01 indicate significantly different from TDCs.

Close modal

Next, we cocultured T cells alone or with mitomycin C- or polyformaldehyde-treated TDCs in the lower assay chamber and with live TDCs or live TDCs plus lymphocytes in the upper chamber. Analysis of the cells collected from the lower chamber revealed that down-regulation of CD3ε and apoptosis in T cells occurred only when live TDCs and lymphocytes were cocultured in the same chamber, even if there was no direct contact with TDCs (Fig. 5, A and B, and data not shown). These results indicate that the observed T cell dysfunction is mediated by one or more factors that are released from live TDCs, but only after those cells have interacted with autologous T cells. In other words, it seems that the dysfunction is not induced by a combination of membrane molecules and soluble mediators that are released from TDCs alone.

T cells are more susceptible to apoptosis when they are activated, and hence we examined the activation status and cytokine production profile of T cells exposed to TDCs. As shown in Fig. 5,C, the expression of CD69, which is a marker of T cell activation (30), was significantly up-regulated on T cells that had been exposed to live TDCs for 24 h. Measuring cytokines in the coculture system over time revealed a rapid accumulation of IFN-γ, TNF-α, and IL-10 in the culture supernatants, and the levels of IFN-γ and TNF-α reached a maximum or a plateau within 24 h (Fig. 5,D). Similar to CD3ε down-regulation and apoptosis, these responses were significantly attenuated when the TDCs were pretreated with mitomycin C (Fig. 5, C and D).

Previous studies using a mouse model have shown that tumor-associated macrophages or immature myeloid cells can induce T cell deletion via production of IFN-γ and TNF-α (31, 32). Inasmuch as we had detected high levels of these cytokines after coculture of lymphocytes and TDCs (Fig. 5,D), we performed new experiments using a neutralizing Ab against IFN-γ or TNF-α to ascertain whether these molecules give rise to T cell dysfunction. The results showed that use of an Ab at a concentration that effectively neutralized IFN-γ or TNF-α in the coculture system (Table II) did not affect the TDC-induced CD3ε down-regulation and subsequent apoptosis in autologous T cells (Fig. 6 A).

Table II.

The blocking effects of IL-10, TNF-α, and IFN-γ in the coculture system by specific mAbs

TreatmentIL-10 (pg/mL)TNF-α (pg/mL)IFN-γ (pg/mL)
Untreated 1745 ± 396 4217 ± 636 3308 ± 711 
IL-10 Ab 321 ± 65** 3899 ± 553 3121 ± 578 
TNF-α Ab 1438 ± 299* 23 ± 5** 2148 ± 499* 
IFN-γ Ab 1621 ± 421 4007 ± 817 479 ± 35** 
TreatmentIL-10 (pg/mL)TNF-α (pg/mL)IFN-γ (pg/mL)
Untreated 1745 ± 396 4217 ± 636 3308 ± 711 
IL-10 Ab 321 ± 65** 3899 ± 553 3121 ± 578 
TNF-α Ab 1438 ± 299* 23 ± 5** 2148 ± 499* 
IFN-γ Ab 1621 ± 421 4007 ± 817 479 ± 35** 

CD14+-depleted PBMCs were cocultured (20:1) with HeLa-exposed TDCs in the presense of neutralizing Ab against IL-10 (1 μg/mL), TNF-α (1 μg/mL), or IFN-γ (5 μg/mL) for 24 h. Thereafter, the levels of IL-10, TNF-α, and IFN-γ in the coculture system were determined by ELISA. Each value represents the mean ± SD of results from at least four separate experiments; ∗, p < 0.05 and ∗∗, p < 0.01 indicate significantly different from untreated TDCs.

FIGURE 6.

TDC triggered T cell dysfunction via an oxygen-dependent pathway. The CD14+-depleted PBMCs were cocultured (20:1) for the indicated times with normal iDCs, HeLa-TDCs (TDCs), or TDCs that have been pretreated with 1 mM 1-MT, 5 μM DPI, 100 U/ml catalase, 200 U/ml SOD, or a blocking Ab to IL-10 (1 μg/ml), TNF-α (1 μg/ml), or IFN-γ (5 μg/ml) for 1 h. A, DPI abolished the TDC-induced down-regulation of CD3ε and apoptosis in T cells. B, DPI impaired the up-regulation of CD69 and the down-regulation of TCR-α/β in T cells after 24-h exposure to TDCs. C, SOD and catalase partially reversed the TDC-induced T cell deletion. Values represent the means ± SE of four separate experiments; ∗, p < 0.05 and ∗∗, p < 0.01 indicate significantly different from TDCs.

FIGURE 6.

TDC triggered T cell dysfunction via an oxygen-dependent pathway. The CD14+-depleted PBMCs were cocultured (20:1) for the indicated times with normal iDCs, HeLa-TDCs (TDCs), or TDCs that have been pretreated with 1 mM 1-MT, 5 μM DPI, 100 U/ml catalase, 200 U/ml SOD, or a blocking Ab to IL-10 (1 μg/ml), TNF-α (1 μg/ml), or IFN-γ (5 μg/ml) for 1 h. A, DPI abolished the TDC-induced down-regulation of CD3ε and apoptosis in T cells. B, DPI impaired the up-regulation of CD69 and the down-regulation of TCR-α/β in T cells after 24-h exposure to TDCs. C, SOD and catalase partially reversed the TDC-induced T cell deletion. Values represent the means ± SE of four separate experiments; ∗, p < 0.05 and ∗∗, p < 0.01 indicate significantly different from TDCs.

Close modal

In studies using other systems (31, 32, 33, 34, 35), it was found that arginase, IDO, and iNOS seemed to be involved in inhibition of T cells mediated by DCs, mouse macrophages, and myeloid suppressor cells. To determine whether any of these molecules are involved in the present observations of T cell dysfunction, we pretreated TDCs with selective inhibitors of these enzymes and then cocultured them with lymphocytes in the presence of the same inhibitor. As shown in Fig. 6 A, pretreatment with 1-MT, a selective IDO inhibitor, did not affect the ability of TDCs to induce CD3ε down-regulation and subsequent apoptosis in autologous T cells. We also investigated possible involvement of arginase and iNOS in this T cell dysfunction by pretreating TDCs with the selective arginase inhibitor N-hydroxy-nor-l -arginine (50 μM) or the selective iNOS inhibitor N-monomethyl-l -arginine (1 mM) (35) before coculture with the lymphocytes. Neither of these inhibitors had any effect on the TDC-induced T cell dysfunction (data not shown).

ROS have been implicated in phagocyte-mediated T cell suppression in both human and mouse tumors (10, 36). To determine whether such a mechanism is involved in the TDC-induced T cell dysfunction, we pretreated TDCs with DPI (5 μM), a flavoprotein inhibitor of NADPH oxidase (26), and subsequently exposed them to autologous T cells. We found that DPI had no effect on spontaneous apoptosis in the T cells (data not shown), whereas it almost completely abolished the TDC-induced CD3ε down-regulation and apoptosis (Fig. 6,A). Such treatment also significantly attenuated the TDC-induced up-regulation of CD69 and down-regulation of TCR-α/β in T cells (Fig. 6 B).

To further confirm the role of ROS in TDC-induced T cell dysfunction, we assessed the effects of the two ROS scavengers SOD and catalase (26) on this process. As shown in Fig. 6 C, addition of these scavengers markedly reversed the TDC-mediated CD3ε down-regulation and apoptosis in T cells. These results clearly show that generation of ROS is a prerequisite for the ability of TDCs to cause T cell dysfunction.

We have recently observed that HA fragments constitute a common factor that is produced by several types of human tumors to induce the formation of immunosuppressive macrophages (22). In the present study, we found that intermediate-sized HA fragments can mimic the effect of TSNs to induce the formation of semimature DCs showing reduced expression of CD1a and elevated production of HLA-DR. Upon stimulation with LPS, the HA-treated DCs were unable to achieve full maturation, and instead exhibited an IL-12lowIL-10high phenotype (Fig. 7, A and B). Additionally, these HA-conditioned DCs could efficiently trigger the CD3ε down-regulation and apoptosis in T cells via a cell-contact mechanism (data not shown).

FIGURE 7.

Tumor-derived HA induced suppression of DCs. A, Purified monocytes were cultured with (solid lines) or without (bold lines) 25 μg/ml INT-HA in the presence of GM-CSF and IL-4 for 6 days. B, The cells were subsequently stimulated with LPS for 24 h, and the expression of surface markers and production of cytokines were determined by flow cytometry and ELISA, respectively. C, Monocytes were left untreated or were pretreated with 10 μg/ml CD44-blocking Ab or control Ab (IgG1), or with 200 μg/ml Pep-1 or control peptide (Cpep), and then cultured with GM-CSF and IL-4 in the presence or absence of TSNs from various tumor cell lines for 6 days, after which maturation was induced by exposure to LPS for 24 h. D, Monocytes were cultured with GM-CSF and IL-4 in the presence or absence of TSNs from parental, mock, or SiHAS2 cells for 6 days, and maturation was subsequently induced by treatment with LPS for 24 h. Levels of IL-12p70 in the culture supernatants were determined by ELISA. Values represent the means ± SE of four separate experiments; ∗, p < 0.05 and ∗∗, p < 0.01 indicate comparison with monocytes preincubated in medium alone (C) or significantly different from parental and mock cells (D).

FIGURE 7.

Tumor-derived HA induced suppression of DCs. A, Purified monocytes were cultured with (solid lines) or without (bold lines) 25 μg/ml INT-HA in the presence of GM-CSF and IL-4 for 6 days. B, The cells were subsequently stimulated with LPS for 24 h, and the expression of surface markers and production of cytokines were determined by flow cytometry and ELISA, respectively. C, Monocytes were left untreated or were pretreated with 10 μg/ml CD44-blocking Ab or control Ab (IgG1), or with 200 μg/ml Pep-1 or control peptide (Cpep), and then cultured with GM-CSF and IL-4 in the presence or absence of TSNs from various tumor cell lines for 6 days, after which maturation was induced by exposure to LPS for 24 h. D, Monocytes were cultured with GM-CSF and IL-4 in the presence or absence of TSNs from parental, mock, or SiHAS2 cells for 6 days, and maturation was subsequently induced by treatment with LPS for 24 h. Levels of IL-12p70 in the culture supernatants were determined by ELISA. Values represent the means ± SE of four separate experiments; ∗, p < 0.05 and ∗∗, p < 0.01 indicate comparison with monocytes preincubated in medium alone (C) or significantly different from parental and mock cells (D).

Close modal

To further confirm the role of HA in the TSN-induced dysfunction of DCs, we pretreated monocytes with agents that antagonize the interactions between HA and its receptors, and then we exposed the cells to TSNs in the presence of GM-CSF and IL-4 for 6 days. CD44 is the major cell surface receptor for HA, and thus we conducted an experiment using the mAb 5F12, which specifically blocks HA-CD44 binding (22, 37). As shown in Fig. 7,C, pretreatment with mAb 5F12 partially restored the ability of TSN-exposed DCs to produce IL-12p70 after stimulation with LPS. Similar results were obtained when the monocytes were preexposed to the HA-specific blocking peptide Pep-1 (21, 22), whereas the isotype-matched IgG1 or a random control peptide had no effect (Fig. 7 C).

Compared with the TSNs from leukemic THP-1 cells, the TSNs from solid tumor cell lines induced formation of semimature DCs that contained significantly higher levels of HA, as compared with amounts detected in our previous study (22). Also in agreement with our earlier study, we noted that silencing of HA synthase 2 (HAS2) partially reduced the levels of HA in the TSNs (22), and consequently our next objective was to examine the effects of TSNs from the stable HAS2-knockdown SK-Hep-1 clones on DC differentiation. We found that the capacity to release IL-12 upon stimulation with LPS was partially restored in iDCs that were exposed to HAS2-knockdown cells for 6 days (Fig. 7 D). These results clearly indicate that HA is a component of TSNs that induces the formation of tolerogenic semimature DCs.

The impaired T cell functions and altered DC phenotype considered herein have been recognized as important mechanisms of tumor immune escape in both mice and cancer patients (7, 12, 38). The present results show that soluble tumor-derived factors, including HA fragments, compel human monocytes to develop into tolerogenic semimature DCs (TDCs). We observed that, upon coming in contact with T cells, the TDCs triggered rapid down-regulation of CD3ε and TCR-α/β and subsequent apoptosis in autologous T cells. Our findings also indicate that the induction of T cell dysfunction does not require the participation of IFN-γ or TNF-α, or the activity of arginase, IDO, or iNOS from TDCs. Instead, we found that pretreatment of TDCs with an inhibitor of NADPH oxidase or ROS scavengers markedly blocked the TDC-mediated T cell dysfunction.

The ability of DCs to initiate immunity or tolerance is largely determined by the incoming signals in the local environment (11, 12). In the present study, we observed that TSNs from a variety of solid tumor cell lines effectively induced the formation of TDCs that exhibited a distinctive IL-12lowIL-10high phenotype and rapidly induced defects in the CD3/TCR complex and apoptosis in autologous T cells. In accordance with that, we found that accumulation of immunosuppressive DCs was associated with CD3ε down-regulation and reduction in the number of T cells in the cancer nests of several different types of human tumors. Interestingly, transwell assays revealed that neither the membrane-bound molecules nor the soluble mediators released by TDCs cultured alone are sufficient to trigger T cell dysfunction. The down-regulation of CD3ε and apoptosis in T cells were observed only after the cells had been directly exposed to live TDCs, which indicates that bidirectional interaction between TDCs and lymphocytes is necessary to generate factor(s) responsible for inducing T cell dysfunction. Indeed, we found that coculture of TDCs and T cells in the upper chamber of the transwell assay system also efficiently induced CD3ε down-regulation and apoptosis in T cells collected from the lower chamber. Such a collaborative influence may reflect a novel immune escape mechanism by which tumors counteract effective T cell responses by inducing the formation of TDCs. Consistent with our observations, other investigators have recently reported that interaction with regulatory T cells can significantly up-regulate the expression of IDO or B7-H4 on APCs, and those APCs in turn suppress T cell responses (3, 39).

Inadequate performance of the TCR/CD3 complex is one of the main mechanisms by which tumors evade T cell responses (7). In this context, both peripheral and tumor-infiltrating lymphocytes from cancer patients have been found to exhibit reduced expression of CD3ζ (15, 16, 17). By comparison, little is known about the role of CD3ε in tumor immunity, although CD3ε is required for generation and/or survival of mature T cells in mice (13, 18). We noted that down-regulation of CD3ε is a critical event in TDC-mediated T cell deletion. The results of kinetic analysis revealed that the reduction of CD3ε occurred before the onset of apoptosis, and all apoptotic T cells displayed reduced CD3ε expression. Agents that prevent T cell apoptosis were found to block CD3ε down-regulation. Moreover, the decreased expression of CD3ε coincided with a significant T cell reduction in the cancer nests in samples of several different types of human tumors. Therefore, such down-regulation of CD3ε might represent a novel mechanism of tumor immune escape. In contrast to previous reports (15, 16, 17, 33), we discovered that exposure to TDCs did not significantly affect the expression of CD3ζ in T cells. Other researchers have observed that CD3ζ down-regulation induced by tumor APCs can be reversed by IL-2 (17, 40), and we found that IL-2 in our coculture systems could not reverse the down-regulation of CD3ε (data not shown). Thus, our findings further support the notion that TDCs suppress CD3ε via a mechanism different from that exerted by tumor-derived macrophages to down-regulate CD3ζ.

In our coculture system, interactions between TDCs and lymphocytes resulted in a rapid accumulation of TNF-α and IFN-γ (Fig. 5 D), both of which have been implicated in the deletion of T cells triggered by immature myeloid cells in mice (31, 32). However, the role of TNF-α and IFN-γ in TDC-induced T cell dysfunction is challenged by our results showing that such T cell dysfunction was not affected by the neutralizing Abs that effectively blocked these cytokines in our culture supernatants. In addition to cytokines, the effects of arginase, IDO, iNOS, and NADPH oxidase have been shown to mediate T cell suppression (3, 32, 33, 34, 35, 36). However, arginase I is barely detectable, and iNOS is generally assumed to be inactive in human APCs (35, 41), which implies that the regulatory mechanisms differ between humans and mice. We tested selective inhibitors of these molecules and found that only an inhibitor of NADPH oxidase was able to effectively block the TDC-mediated T cell dysfunction, indicating that generation of ROS by TDCs is a prerequisite for induction of the dysfunction. This conclusion is supported by our finding that two ROS scavengers also markedly reversed the TDC-effected down-regulation of CD3ε and apoptosis in T cells. Although the mechanisms in this context are not yet completely understood, it has been shown that ROS directly or indirectly target mitochondria and release cytochrome c from those organelles into the cytosol, which leads to caspase activation. Moreover, increased levels of ROS can suppress expression of Bcl-2 and induce T cell death via the intrinsic apoptosis pathway, and that cascade is essential for activated T cell autonomous death (42). Interestingly, Thorén and colleagues have observed that the antioxidative properties of normal myeloid DCs can protect T cells and NK cells from inactivation and apoptosis induced by oxygen radicals (43). Therefore, it is plausible that such a mechanism contributes to the opposing effects of TDCs on T cell functions.

In cancer patients, HA concentrations are usually higher in malignant tumors than in corresponding benign or normal tissues (44, 45). Experimental studies have revealed that not only does HA promote tumor growth and metastasis, but it also modulates the functions of APCs (21, 46). In support of those findings, the results of three sets of experiments in our investigation provide evidence that HA is a common factor that is produced by several types of solid tumor cell lines to alter DC maturation. First, we found that purified HA fragments were able to mimic the ability of TSNs to induce the formation of TDCs. Second, pretreatment with an anti-CD44 mAb or Pep-1 to antagonize the interactions between HA and its receptors partially restored the capacity of TDCs to release IL-12p70. Third, silencing of HAS2 in tumor cells, which partially reduced the HA levels in TSNs, attenuated the ability of those cells to induce DC dysfunction. Therefore, HA fragments generated in tumor microenvironments may constitute a common mediator of the formation of TDCs.

In all our experiments, anti-CD44 mAb or Pep-1 could only partially restored the ability of IL-12 production by TSN-exposed DCs, but had few, if any, effects on their ability to induce T cell dysfunction (data not shown). These results suggest that the effect of the TSNs involved additional soluble factors from tumor cells. Tumor cells are known to release a variety of factors, including immune suppressive cytokines, α-fetoprotein, glycoprotein, prostanoids, MUC1, and gangliosides to alter the maturation and function of DCs (1, 7, 11, 12, 47, 48, 49, 50).

Our results give important new insights into the collaborative action of tumors that is exercised to counteract effective T cell responses. HA and other soluble factors derived from cancer cells can drive monocytes to develop into suppressive TDCs. After interacting directly with T cells, the TDCs act via oxygen-dependent pathways to rapidly induce defects in the TCR/CD3 complex and subsequently provoke apoptosis in T cells, and in this way they create conditions of unresponsiveness in the cancer nests. In support of this conclusion, we observed that the number of suppressive APCs in the cancer nests of primary hepatocellular carcinomas was inversely associated with the overall survival of the patients, which can serve as an independent predictor of relapse-free survival and overall survival (our unpublished results). Therefore, it is possible that studies of the mechanisms that can selectively modulate the phenotype of APCs will provide a novel strategy for anticancer therapy (38, 51).

We thank Patricia Ödman for linguistic revision of the manuscript and Guoping Zhang for help with confocal microscopy.

The authors have no financial conflicts of interest.

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.

1

This work was supported by the Outstanding Young Scientist Fund and project grants from the National Natural Science Foundation of China (30425025, 30672388 and 30730086), the “973” Program (2004CB518801 and 2007CB512404), and the Natural Science Foundation of Guangdong (05200303).

3

Abbreviations used in this paper: DC, dentritic cell; TSN, tumor culture supernatant; TDC, TSN-exposed tolerogenic semimature DC; iNOS, inducible nitric oxide synthase; iDC, immature DC; mDC, mature DC; ROS, reactive oxygen species; HA, hyaluronan; DPI, diphenyleneiodonium; SOD, superoxide dismutase; 1-MT, 1-methyl-tryptophan.

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