In the injured liver, hepatic stellate cells (HSCs) secrete many different cytokines, recruit lymphocytes, and thus participate actively in the pathogenesis of liver disease. Little is known of the role of HSCs in immune responses. In this study, HSCs isolated from C57BL/10 (H2b) mice were found to express scant key surface molecules in the quiescent stage. Activated HSCs express major histocompatibility complex class I, costimulatory molecules, and produce a variety of cytokines. Stimulation by interferon γ (IFN-γ) or activated T cells enhanced expression of these molecules. Interestingly, addition of the activated (but not quiescent) HSCs suppressed thymidine uptake by T cells that were stimulated by alloantigens or by anti-CD3–mediated T-cell receptor ligation in a dose-dependent manner. High cytokine production by the T cells suggests that the inhibition was probably not a result of suppression of their activation. T-cell division was also found to be normal in a CFSE dilution assay. The HSC-induced T-cell hyporesponsiveness was associated with enhanced T-cell apoptosis. Activation of HSCs was associated with markedly enhanced expression of B7-H1. Blockade of B7-H1/PD-1 ligation significantly reduced HSC immunomodulatory activity, suggesting an important role of B7-H1. In conclusion, the bidirectional interactions between HSCs and immune cells may contribute to hepatic immune tolerance. (HEPATOLOGY 2004;40:1312–1321.)
If you can't find a tool you're looking for, please click the link at the top of the page to "Go to old article view". Alternatively, view our Knowledge Base articles for additional help. Your feedback is important to us, so please let us know if you have comments or ideas for improvement.
The liver provides a microenvironment of diminished immunogenicity, and in some circumstances of tolerogenicity. Thus, liver grafts transplanted in many species survive without the requirement for immunosuppressive therapy.1–3 In humans, immunosuppressive therapy is required to prevent liver allograft rejection, but a liver immunomodulatory effect is also apparent. Transplantation across ABO barriers leads to accelerated rejection in kidneys and hearts, but is rarely witnessed in livers.4 Accelerated rejection caused by the presence of positive antidonor antibodies, observed commonly for kidneys, is rare in human liver allografts.5–6 Liver allograft loss from chronic rejection is infrequent compared with that of the kidney and heart.7, 8 There are well-documented cases of liver transplant patients who survive following withdrawal of immunosuppression.9–13 In addition, induction of systemic tolerance through the oral administration of antigens has also been attributed to the liver, because similar tolerance can be induced through direct delivery of antigen via the portal vein, but not via the intravenous route.14–16 Hepatic tolerance may also contribute to the liver's vulnerability to chronic infections, such as hepatitis B virus, hepatitis C virus, and hepatic parasitic infections.17
It is unlikely that hepatic tolerance is a result of deletion of specific T-cell clones, because spleen cells isolated from recipients with accepted liver allograft exhibit profound proliferative and specific cytotoxic responses to donor alloantigens in vitro.18 A T-cell receptor (TCR) Vβ usage study in which lymphocytes were isolated from such recipients showed no clonal deletion.18 The liver has unique immunological properties, suggesting unusual interactions within the hepatic milieu and immune cells. It has been shown in a mouse liver transplant model that significant CD4+ and CD8+ T-cell infiltration occurs in liver allografts within a few hours and peaks within a week, but it decreases gradually thereafter. This result is associated with high levels of T-lymphocyte apoptosis in the liver allografts,19, 20 which occurs as early as 18 hours following their infiltration.21, 22
Hepatic stellate cells (HSCs) play an important role in the regulation of sinusoidal blood flow (contractility), maintainance of hepatic architecture (extracellular matrix synthesis), and production of various growth factors. During liver injury, HSCs transform into myofibroblast-like cells and deposit extracellular matrix, causing liver fibrosis. Activated HSCs also recruit lymphocytes by producing chemokines,23 and they produce numerous cytokines with proinflammatory as well as anti-inflammatory activity.24–26 In this study, we demonstrated an inhibitory effect of HSCs on immune responses through interactions between HSCs and T cells. Activated murine HSCs strongly suppress T-cell responses. This was associated with apoptosis of activated T cells. Upregulation of B7-H1 (also called programmed death ligand-1), an inhibitory molecule of the B7 family, on activated HSCs may contribute to this immunomodulatory activity.
HSC, hepatic stellate cell; B10, C57BL/10; TCR, T-cell receptor; OVA, ovalbumin; mAb, monoclonal antibody; MHC, major histocompatibility complex; DC, dendritic cell; IL, interleukin; IgG, immunoglobulin G; CFSE, carboxyfluorescein diacetate succinimidyl ester; MLR, mixed leukocyte reaction; cpm, counts per minute; TUNEL, terminal deoxynucleotidyl transferase–mediated dUTP nick end labeling; IFN, interferon; mRNA, messenger RNA; TGF, transforming growth factor; CTL, cytotoxic T lymphocyte
Materials and Methods
Male C57BL/10 (B10; H-2b), C3H (H-2k), BALB/c (H-2d), and DO11.10 (H-2d) mice were purchased from the Jackson Laboratory (Bar Harbor, ME). The TCR on CD4+ T cells in the DO11.10 mice is encoded by transgenes that recognize a chicken ovalbumin (OVA)-derived peptide, OVA323–339, presented by I-Ad. This TCR can be identified by the anticlonotypic monoclonal antibody (mAb) KJ1.26. The Des TCR-transgenic mice (H-2k) express the clonotype TCR on CD8+ T cells that is specific for major histocompatibility complex (MHC) class I H-2Kb molecules as described previously.24 All animals were maintained in the specific pathogen-free facility of the University of Pittsburgh Medical Center, were provided with Purina rodent chow (Ralston Purina, St. Louis, MO) and tap water ad libitum, and were used at 8 to 12 weeks of age in accordance with the guidance of the National Institutes of Health (publication 86-23, revised 1985).
Isolation and Culture of HSCs.
HSCs were isolated from mouse livers as described previously,25, 26 with some modifications. Briefly, the liver was perfused via the portal vein with 30 mL Ca2+-Mg2+–free Hank's balanced salt solution (Mediatech Inc., Herndon, VA) in situ (5 mL/min), flowed by perfusion with 1 mL collagenase IV (1 mg/mL, Sigma, St. Louis, MO). The liver was removed, meshed, and agitated in collagenase (1 mg/mL) at 37°C for 30 minutes. Cells were filtered through a nylon mesh and purified via Percoll (Sigma) gradient centrifugation. The isolated HSCs were cultured (105/mL) in an uncoated plastic flask (Nunclon, Roskilde, Denmark) with RPMI-1640 (Mediatech Inc.) supplemented with 10% fetal calf serum and 10% horse serum in 5% CO2 in air at 37°C for 2 (quiescent), 4 to 5 (intermediate), or 7 to 10 (activated) days. Cell viability was greater than 90% as determined using trypan blue exclusion. The purity of HSCs ranged from 90% to 95% as determined by desmin immunostaining and the typical light microscopic appearance of the lipid droplets as described previously.27 HSCs cultured for 2 days demonstrated quiescent features, with a round or star shape, abundant lipid droplets, and a lack of α smooth muscle actin expression (quiescent). Cells from the days 7 to 10 culture were spread out, had few lipid droplets, and expressed α smooth muscle actin (activated). Cells from the days 4 to 5 culture had an intermediate appearance as described previously.27, 28 HSCs cultured on flask for 7 days are referred hereafter as “activated HSCs” unless specified otherwise.
Culture of Dendritic Cells.
Bone marrow cells isolated from mouse femurs and tibias were lysed of red blood cells using red blood cell lysis buffer (Sigma), cultured in RPMI-1640 medium containing 10% v/v heat-inactivated fetal calf serum, 20 mmol/L hydroxyethylpiperazine-N-2 ethanesulfonic acid, 2 mmol/L L-glutamine, 0.1 mmol/L nonessential amino acids, 1 mmol/L sodium pyruvate, 20 μmol/L 2-mercaptoethanol, and antibiotics (100 U/mL penicillin, 100 mg/mL streptomycin; referred to subsequently as complete medium) in the presence of mouse recombinant granulocyte-macrophage colony-stimulating factor (4 ng/mL) and interleukin (IL)-4 (1,000 U/mL) (both from Schering-Plough, Kenilworth, NJ). Nonadherent cells were released spontaneously from the proliferating cell clusters. These cells were then harvested, washed, and resuspended in complete medium as described previously.29
mAb and Flow Cytometry.
Expression of cell surface molecules was determined via flow cytometric analysis using an EPICS ELITE flow cytometer (Coulter Corp., Hialeah, FL). Cells were stained with primary hamster or rat mAbs against B220, CD3, CD14, CD40, CD45, CD54, CD80, CD86, CD11a, or CD11b followed by secondary goat anti–hamster immunoglobulin G (IgG) or anti–rat IgG2a, as described previously.24, 30 MHC class I and II antigens were detected with mAbs against H-2b and I-Ab, respectively (all from BD PharMingen, San Diego, CA). Anti–mouse DO11.10 TCR (KJ1.26) was purchased from Caltag Laboratories (Burlingame, CA). The Des TCR determinant was stained with anti–Des mAb (mouse IgG2a, the hybridoma, was a kind gift from Dr. B.F. de St Groth, Centenary Institute of Cancer Medicine and Cell Biology, Sydney, Australia). Expression of B7-H1 on HSCs was identified using anti–B7-H1–specific mAb (rat IgG2a, eBioscience, San Diego, CA). The appropriate isotype control antibodies were used in the experiments.
Labeling of Cells With CFSE.
T cells (107/mL) were labeled with 5 μmol/L carboxyfluorescein diacetate succinimidyl ester (CFSE) (Molecular Probes, Eugene, OR) from a 10-mmol/L stock solution (in dimethylsulfoxide) for 10 minutes at 37°C, and washed before culture. The intensity of CFSE was determined via flow cytometric analysis.
T-Cell Proliferation Assay.
Proliferation of nylon wool-eluted C3H spleen T cells (2 × 105/well in 100 μL) was elicited via allogeneic DCs in a one-way mixed leukocyte reaction (MLR) or via anti-CD3 mAb. Cultures were performed in triplicate in 96-well round-bottom microculture plates (Corning, Corning, NY). Graded doses of gamma-irradiated (20 Gy; X-ray source) DCs from B10 mice were used as stimulators. Cultures were maintained in RPMI-1640 complete medium for 3 to 4 days in 5% CO2 in air. For stimulation with anti-CD3, anti-CD3ϵ mAb (BD PharMingen) was coated onto flat-bottom, 96-well plates (Corning) at 2 μg/mL in 100 μL phosphate-buffered saline (pH 7.2) at 4°C overnight. The plates were washed with phosphate-buffered saline twice before seeding the T cells in RPMI-1640 complete medium and were cultured for 2 days in 5% CO2 in air. [3H]-thymidine (1 μCi/well) was added for the final 18 hours, and incorporation of [3H]-thymidine into DNA was assessed via liquid scintillation counting. Results were expressed as mean counts per minute (cpm) ± 1 SD. To examine the effect of HSCs on T-cell proliferation, gamma-irradiated (50 Gy) HSCs were added into the cultures at the beginning of the culture unless otherwise indicated.
Cytotoxic T-Lymphocyte Assay.
C3H spleen T cells were cultured with gamma-irradiated (20 Gy) B10 allogeneic DCs at a ratio of 10:1 for 5 to 6 days, then used as effectors. EL4 (H2b), R1.1 (H2k) or P815 (H2d) lymphoma cells (4 × 106, all from American Type Culture Collection, Rockville, MD) were labeled with 100 μCi Na251CrO4 (NEN, Boston, MA) and used as donor-specific, syngeneic, or third-party targets. They were plated in 96-well round-bottom culture plates (5 × 103 cells/well, Corning). Serial, 2-fold dilutions of effector cells were added to give effector/target ratios of 100:1, 50:1, 25:1, and 12.5:1 in a total volume of 200 μL/well. The specific 51Cr release was determined after incubation for 4 hours at 37°C in RPMI-1640 complete medium in 5% CO2 in air. An aliquot (100 μL) of supernatant was recovered from each well after centrifugation at 300g for 1 minute. Maximum 51Cr release was determined by lysis of the target cells. The percent cytotoxicity was calculated using the following formula: % cytotoxicity = 100 × [(experimental cpm) − (spontaneous cpm)] / [(maximum cpm) − (spontaneous cpm)], and expressed as the mean ± SD of percentage-specific 51Cr release in triplicate cultures.
Apoptotic T cells were identified via double-staining with anti-CD3 mAb and terminal deoxynucleotidyl transferase–mediated dUTP nick end labeling (TUNEL). Following surface CD3 staining, cells were fixed in 4% paraformaldehyde and permeabilized with 0.1% Triton X-100 and 0.1% sodium citrate. The TUNEL reaction mixture of the Cell Death Detection kit (Roche Diagnostics, Indianapolis, IN) was then added according to the manufacturer's instructions. Cells incubated with the labeling solution in the absence of terminal transferase were used as negative controls. Cells were analyzed via flow cytometry with 10,000 events from each sample.
To detect apoptosis in cell suspension by microscopy, 1 × 107 cells in a plate were fixed with 3.7% (W/V) formaldehyde in methanol for 10 minutes, then dropped onto 1-cm2 slides to dry at room temperature. The cells were stained via TUNEL using an in situ apoptosis detection kit (Intergen, Norcross, GA) according to the manufacturer's instructions.
The cell culture supernatants were harvested and analyzed for the presence of cytokines using commercial enzyme-linked immunosorbent assay kits (R&D Systems, Minneapolis, MN) according to the manufacturer's protocols.
Ribonuclease Protection Assay.
Total RNA was isolated using TRIzol reagent (Invitrogen, Carlsbad, CA). The assay was performed using the RiboQuant Multi-probe RNase Protection Assay kit (BD PharMingen). The probes were synthesized with T7 RNA polymerase with incorporation of α-32P-UTP. Five micrograms of RNA was treated overnight with radiolabeled probes (specific activity: 800 Ci/mmol/L) at 56°C, followed by treatment with ribonuclease A (80 μg/mL) and T1 (250 U/mL) for 45 minutes at 30°C. The murine L32 and glyceraldehyde-3-phosphate dehydrogenase riboprobes were used as controls. Protected fragments were subjected to electrophoresis through a 7.0 mol/L urea/5% polyacrylamide gel, then exposed to Kodak X-omat film for 72 hours.
The parametric data are given as the mean ± SD. Statistical significance was determined using the Student t test; P values of less than .05 were considered statistically significant.
Phenotype of HSCs.
Expression of key molecules on the surface of HSCs was examined using flow cytometry. Neither quiescent nor activated HSCs expressed markers for leukocytes (CD45), T cells (CD3), B cells (B220), or macrophages (CD14 and CD11b) (Fig. 1A). Quiescent HSCs expressed low levels of MHC class I and II, CD40, CD80, CD86, and CD54, while activation by culture on uncoated plastic for 7 days induced upregulated expression of MHC class I, CD40, CD80, CD86, and CD54 (intercellular adhesion molecule 1), and minimal increased expression of MHC class II (Fig. 1A). Further stimulation with interferon (IFN)-γ enhanced expression of MHC class I, CD86, and CD54 on activated HSCs (Fig. 1A).
Quiescent HSCs expressed low messenger RNA (mRNA) levels of IL-1Rα, transforming growth factor β1 (TGFβ1), and migration inhibitory factor. Activation of HSCs by 7-day culture induced upregulation of IL-6, TGFβ, and migration inhibitory factor mRNA transcription. Further stimulation by IFN-γ or syngeneic or naïve allogeneic T cells did not further upregulate cytokine expression in activated HSCs. Activated allogeneic T cells appeared to be the strong stimulators of HSC activation, resulting in significantly enhanced expression of inflammatory cytokines—including IL-1α, its receptor, and IL-6 (Fig. 1B).
HSCs Inhibit T-Cell Responses Elicited by Allogeneic DCs.
To address the effect of HSCs on T-cell responses, irradiated B10 HSCs were added to an MLR culture in which irradiated B10 DCs (H-2b) and C3H spleen T cells (H-2k) were used as stimulators and responders, respectively. Figure 2A demonstrates that the presence of activated B10 HSCs markedly inhibited T-cell proliferative responses in a dose-dependent manner. Seven-day cultured HSCs appeared to generate the greatest inhibitory activity, suppressing approximately 50% of thymidine uptake via DC-activated T cells at a HSC/T ratio of 1:40, and suppressing 90% at a ratio of 1:20. The inhibitory activity was dependent on the stage of HSC activation, because quiescent HSCs showed little inhibitory activity, and intermediately activated HSCs (5-day culture) exerted modest but significant inhibitory activity. Generation of specific cytotoxic T lymphocytes (CTLs) in these B10/DC-stimulated C3H spleen T cells was also markedly inhibited (Fig. 2B).
HSCs Do Not Inhibit T-Cell Activation.
To determine whether HSCs directly inhibit the activation of T cells, the levels of IL-2, IFN-γ, and IL-10 in MLR culture in the presence or absence of HSCs were analyzed via enzyme-linked immunosorbent assay. These cytokines were predominantly produced by T cells, because the ratio of T cells/DCs/HSCs in the culture was 40:2:1. In addition, previous studies have shown that DCs do not produce IL-2 and secrete very low levels of IFN-γ and IL-10.31 There is no evidence that HSCs produce IL-232 or IFN-γ.33 Activated HSCs produce IL-10,34, 35 but they may not contribute to the IL-10 expression in these experiments considering their very small proportion in the coculture. Although the expression of both IL-2 and IL-10 was slightly but significantly inhibited, IFN-γ levels remained high (Fig. 2C), indicating that T-cell activation was not markedly affected. This was confirmed by the results of T-cell mRNA expression. Activated HSCs slightly inhibited the transcription of IL-2, but not of IFN-γ mRNA, whereas they enhanced expression of IL-10 and IL-6 mRNA. It was noted that mRNA of T cells for IL-4 and IL-9 was markedly suppressed in the presence of HSCs (Fig. 2D).
Suppression of T-Cell Responses by HSCs Is Not MHC Specific.
Activated HSCs from C3H (syngeneic to T cells) or B10 (allogeneic to T cells) mice were added into MLR culture, in which B10 DCs and C3H T cells were used as stimulators and responders, respectively. HSCs from either C3H or B10 mouse livers markedly suppressed thymidine uptake by T cells in a dose-dependent fashion (Fig. 3A), suggesting that the inhibitory effect of HSCs on T-cell response is not MHC specific. Activated HSCs also inhibited [3H]-thymidine incorporation of T cells that were stimulated through TCR ligation with anti-CD3 mAb (Fig. 3B).
Inhibitory Effect of HSCs Is Associated With T-Cell Apoptosis.
We speculated that HSC-induced T-cell hyporesponsiveness may result from apoptotic death of activated T cells. To address this, C3H T cells were cultured for 3 days with irradiated allogeneic (B10) DCs in the presence or absence of activated B10 HSCs (T-cell/DC/HSC ratio of 40:2:1). The cell suspension was then stained via TUNEL. Microscopic examination revealed that the presence of HSCs markedly increased TUNEL-positive cell number (Fig. 4A). Flow cytometric analysis of the cells that were double-stained with anti-CD3 mAb and TUNEL confirmed that the activated HSCs enhanced incidence of TUNEL-positive cells in the T-cell population (Fig. 4B). The addition of z-VAD-fmk—a common caspase pathway inhibitor—partially but significantly reversed HSC-induced suppression of T-cell thymidine uptake (Fig. 4C), indicating that caspase-dependent apoptotic death of T cells is involved. These data suggest that induction of T-cell death involving a caspase pathway may contribute to the inhibitory effect of activated HSCs on T-cell responses.
Effect of HSCs on the Responses of Antigen-Specific CD4+ and CD8+ T Cells.
To more precisely analyze the influence of HSCs on T-cell activation and survival, antigen-specific TCR transgenic CD4+ and CD8+ T cells were used. DO11.10 (H2d) mice express an OVA-specific, transgene-encoded TCR on CD4+ T cells that can be specifically identified with mAb KJ1.26. T cells isolated from the spleens and lymph nodes of DO11.10 mice were labeled with CFSE and cultured with irradiated BALB/c (H2d) DC plus OVA peptide for 3 days. As shown in Fig. 5A, DCs effectively presented the OVA peptide to OVA-specific T cells, as evidenced by a significant increase in the incidence of KJ1.26+ T cells and vigorous T-cell division in the KJ 1.26+ cell population detected via CFSE dilution. The addition of activated HSCs to the culture did not affect the pattern of KJ1.26+ T-cell division; however, the incidence of KJ1.26+ T cells was reduced. This was associated with high levels of apoptotic cells in the culture identified with TUNEL staining (Fig. 5B), suggesting that activated HSCs do not directly inhibit specific CD4+ T-cell proliferative responses to an antigen stimulation, but they may induce T-cell death.
We examined the effect of HSCs on activation and survival of antigen-specific CD8+ T cells using Des mice whose CD8+ T cells with TCR transgene recognize an H2Kb class I MHC antigens, and can be identified by a specific anti–Des mAb. Activated HSCs inhibited expansion of antigen-specific CD8+ T cells elicited by B10 (H2Kb) DCs, but did not affect T-cell proliferation as determined by CFSE dilution (Fig. 5C). Taken together, these results indicate that both activated CD4+ and CD8+ T cells are sensitive to HSC-induced apoptosis.
A Role for B7-H1 in Immunosuppressive Activity of HSCs.
B7-H1 is a recently identified member of the B7 family that negatively regulates T-cell immune responses.36–39 Its receptor, PD-1, can be induced on T cells and B cells upon activation.40 We demonstrated that quiescent HSCs expressed very low levels of B7-H1, while its expression was enhanced following in vitro culture, and markedly augmented by further stimulation by IFN-γ, or activated allogeneic T cells (Fig. 6A), indicating an inducible nature on HSCs. To determine whether B7-H1 plays a critical role in the immunosuppressive function of activated HSCs, an anti–B7-H1 blocking mAb was added to T cells, the proliferation of which was elicited by CD3 mAb–mediated TCR ligation and inhibited by the presence of activated HSCs. We examined anti–B7-H1 mAb at concentrations of 3.75 μg/mL to 30 μg/mL and found that the antibody elicited maximum effects at 7.5 to 15 μg/mL; the blocking effects became less effective at higher concentrations. Thus, a concentration of 10 μg/mL was used in the following studies. Figure 6B shows that addition of anti–B7-H1 mAb partially but significantly reversed the inhibition of T-cell proliferative responses induced by activated HSCs; this was associated with the reduction of T-cell apoptosis (Fig. 6C).
The anatomy of liver allows interactions between hepatic cell populations such as lymphocytes, antigen-presenting cells, endothelial cells, and HSCs. In this study, we demonstrate that mouse liver HSCs markedly inhibit T-cell responses. The immunosuppressive activity of HSCs may represent one of the mechanisms that regulate immune responses in the liver favoring the induction of tolerance rather than the induction of immunity.41 The liver receives blood from the gastrointestinal tract, containing bacterial products and food-derived antigens. These antigens and microbiologically derived molecules impose constraints on immune responses in the liver, and it has been hypothesized that there are control mechanisms that determine whether antigen encounter will result in immunity or tolerance.22 HSCs may modulate liver immunity toward tolerant outcome in these sophisticated mechanisms. Our results also show that the immune inhibitory activity of HSCs is not MHC specific, because HSCs—whether they are syngeneic or allogeneic to T cells—can similarly suppress T-cell responses. TCR-mediated T-cell responses by anti-CD3 mAb are also significantly suppressed, suggesting an “innate” immunoregulatory nature of HSCs.
It has been shown that HSC function can be influenced by a variety of cytokines produced by T cells.27, 42 Thus, TGFβ and tumor necrosis factor exert stimulatory effects by activating HSCs.43 Tumor necrosis factor and IL-1 induce HSCs to express chemokines and adhesion molecules that enhance T-cell recruitment.32 In contrast, IFN-α and IFN-γ inhibit HSC activation.44, 45 Our results demonstrate a bidirectional interaction between HSCs and immune cells. HSCs freshly isolated from normal mouse livers express low levels of key molecules that are required for immunoregulation. The activated HSCs—but not quiescent HSCs—markedly inhibit T-cell responses, indicating that HSCs acquire immunoregulatory activity during activation. HSCs undergo activation through exposure to IFN-γ, an inflammatory cytokine released by activated T cells; this results in upregulation of both stimulatory (CD80, CD86, and CD54) and inhibitory (B7-H1) surface molecules, as well as enhancement of both inflammatory cytokines (IL-6 and IL-1) and suppressive cytokines (TGFβ). However, activated HSCs manifest profound immunosuppressive activity, indicating that the enhanced function of these inhibitory molecules overrides the stimulatory counterparts. Alloreactive T cells—in particular T cells activated by allogeneic DCs—appear to be potent stimulators of HSCs, suggesting that circulating leukocytes that infiltrate the liver may provide a broad spectrum of mediators that modulate HSC behavior.
Another important finding is that the inhibition of T-cell responses by activated HSCs is not a result of direct inhibition of T-cell activation. Thus, although thymidine uptake of activated T cells was dramatically inhibited by HSCs, IFN-γ production remains high. The effect of activated HSCs on the division and expansion of antigen-specific T cells in response to antigen stimulation confirms the fact that HSCs suppress T-cell responses by causing their apoptosis without inhibiting their activation. Therefore, HSCs may contribute to hepatic tolerance induction by deletion of activated T cells through induction of apoptosis. These observations are consistent with a recent report of a concanavalin A–induced hepatitis model in which T-cell apoptosis was preferentially distributed in the hepatic portal triad rather than the parenchyma, where apoptotic cells were spatially associated with α smooth muscle actin–positive myofibroblast-like HSCs.46In vitro experiments in this same study also demonstrated that lymphocyte apoptosis was more frequently induced in the coculture of concanavalin A–activated splenic T cells/activated HSCs compared with quiescent cells.
The partial reversal of HSC-suppressed T-cell thymidine uptake by the caspase inhibitor z-VAD-fmk suggests the involvement of both caspase-dependent and -independent mechanisms. In addition, we demonstrate that quiescent HSCs express very low levels of B7-H1, whereas B7-H1 expression on HSCs is markedly increased by exposure to various stimuli. Most importantly, inhibition of B7-H1 by a blocking mAb partially but significantly reverses the HCS-induced suppressive effect on T-cell responses via reduction of T-cell apoptotic death, suggesting a role of inducible B7-H1 on HSCs. These observations support recent reports that PD-1 ligation negatively regulates the immune system and plays an important role in T-cell downregulation at the effector phases.24, 36 PD-1 contains an immune receptor tyrosine-based inhibitory motif that negatively regulates cytokine synthesis.37, 38 Ligation of PD-1 and TCR leads to rapid phosphorylation of SHP-2, a phosphatase that attenuates TCR signaling.39 B7-H1 expressed on tumor cells actively inhibits immune responses by promoting the apoptosis of effector CTLs.47 A recent study demonstrates that the deficiency of B7-H1 results in accumulation of CD8+ T cells but not CD4+ T cells in the liver, suggesting a role of B7-H1 in regulating T-cell homeostasis.48 PD-1 ligation inhibits antiviral immunity in the liver, and the absence of PD-1 results in rapid clearance of the infected adenovirus.49 Our data suggest that the inducible expression of B7-H1 on HSCs may reflect a new crucial paradigm of B7 inhibitory family-mediated immunoregulation in the liver. However, compared with the controls, addition of anti–B7-H1 mAb only partially reverses the inhibition of T-cell proliferative responses and apoptotic activity, suggesting that factors other than B7-H1 ligation may be involved in these inhibitory processes. We used anti–B7-H1 mAb instead of anti–PD-1 mAb or fusion protein in this study because it has been demonstrated that B7-H1 may have receptors other than PD-1 on activated T cells.50–52 Thus, the B7-H1–mediated apoptosis of CTLs could not be consistently abrogated by a decoy PD-1.37, 38 In addition, there was no significant decrease of apoptosis in both activated CD4+ and CD8+ T cells in the liver of PD-1 knockout mice, suggesting that the deleterious effect of PDL-1 (B7-H1) on activated T cells is mediated through receptors other than PD-1.48
The authors thank Dr. Ngoc Thai for his critical review and Alison Logar for her assistance with flow cytometry.