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The Journal of Immunology, 2002, 169: 2407-2413.
Copyright © 2002 by The American Association of Immunologists

Immunomodulatory Effects of the Liver: Deletion of Activated CD4+ Effector Cells and Suppression of IFN-{gamma}-Producing Cells After Intravenous Protein Immunization1

Katja Klugewitz2,*,{dagger}, Friderike Blumenthal-Barby{dagger}, Arnhild Schrage{dagger}, Percy A. Knolle{ddagger}, Alf Hamann{dagger} and Ian Nicholas Crispe3,*

* Section of Immunobiology, Yale University Medical School, New Haven, CT 06520; {dagger} Experimentelle Rheumatologie, Charité, c/o Deutsches Rheumaforschungszentrum, Berlin, Germany; and {ddagger} Zentrum f. Molekulare Biologie, Heidelberg, Germany


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The liver is tolerogenic in many situations, including as an allograft and during the response to allogeneic MHC expressed on hepatocytes. The majority of data that address this issue focus on endogenous Ags. Little is known about CD4+ T cells and their fate under tolerizing conditions, especially with respect to fully differentiated CD4+ effector T cells. In this study, we used the adoptive transfer of populations of TCR-transgenic CD4+ T cells, skewed toward the Th1 or Th2 phenotype, to test whether either apoptotic or immune deviation mechanisms apply to cytokine-producing CD4+ T cells that enter the liver. After transfer, Th1 and Th2 cells could be detected up to 25 days in lymphoid organs and the liver. Intravenous high dose Ag application resulted in accumulation, proliferation, and subsequent deletion of effector cells within the liver. Th1 cells lost their capacity to produce cytokines, whereas IL-4 expression was sustained within Th2 cells from the liver. However, there was no evidence for a deviation of Th1-programmed cells toward a Th2 (IL-4) or regulatory T cell (IL-10) pattern of cytokine expression. We used isolated populations of liver-derived APCs to test whether the liver had the capacity to impose a bias toward IL-4 expression in T cells. These experiments showed that liver sinusoidal endothelial cells selectively suppress the expansion of IFN-{gamma}-producing cells, yet they promote the outgrowth of IL-4-expressing Th2 cells, creating an immune suppressive milieu within this organ. These data suggest that presentation of Ags in the liver leads to modulation of immune response in terms of quantity and quality.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The liver is tolerant as an allograft, and imposes tolerance on adoptively transferred CD8+ T cells that recognize allogeneic MHC class I Ags expressed on hepatocytes. In contrast, transplantation of other organs and MHC class I Ag expression in other tissues often lead to immunity (1, 2, 3, 4, 5). How does such tolerance operate? Presentation of Ag by liver-derived APC may lead to immune deviation, with the activation of anti-inflammatory T cells (6, 7, 8). However, the normal liver also contains a large population of dying T cells, suggesting that T cell apoptosis is one of its normal functions (9, 10). Either or both of these mechanisms could be important in liver tolerance. Previous investigations have shown that apart from these subsets, the liver also contains a relatively high frequency of cytokine-producing CD4+ effector T cells of both a Th1 and Th2 type (10, 11, 12). Th1 (inflammatory) cells express IFN-{gamma}, whereas Th2 (anti-inflammatory) cells synthesize mainly IL-4. Their mutual influence and their balance are crucial for the delivery of an immune reaction (13, 14, 15, 16). In short-term migration studies, these cells are more efficiently recruited to the liver than resting CD4+ T cells, with some preference for Th1 as opposed to Th2 cells (17, 18). In this study, we aimed to clarify what mechanisms of tolerance induction toward exogenous Ags apply to adoptively transferred Th1 and Th2 cells. We hypothesized that Th1 and Th2 cells undergo different fates within the liver, resulting in alterations in the balance of pro- and anti-inflammatory effector cells.

Our data showed that proinflammatory Th1 cells became cytokine nonsecretory, while Th2 cells maintained their capacity to secrete IL-4 within the liver. Both types of cells were subject to intrahepatic apoptosis. This supports the point of view that, while death is a common fate for activated CD4+ T cells that localize to the liver, the mechanisms of tolerance induction are not accelerated death of inflammatory cells, but selective suppression of their capacity to make Th1 cytokines.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Antibodies

Directly labeled Abs against surface markers and cytokines, and annexin V FITC were purchased from BD PharMingen (San Diego, CA). KJ1.26 digoxigenin, anti-digoxigenin Cy5, anti-IFN-{gamma} FITC (XMG6), unlabeled cytokine Abs for cell culture, anti-IL-4, 11B11, anti-IL-12, C15.6.7, C17.8, C17.15, anti-IFN-{gamma}, XMG6, AN18.17.24, R4.6A2, and anti-FcII/III (2.4G2) were kindly provided by the Deutsches Rheumaforschungszentrum (Berlin, Germany). The mouse endothelial cell mAb ME-9F1 was made, purified, and provided by A. Hamann’s group. KJ1.26 biotin was a gift from K. Bottomly (Section of Immunobiology, Yale University). Rat IgG was purchased from Sigma-Aldrich (St. Louis, MO).

Reagents

Saponin, paraformaldehyde, PMA, ionomycin, BSA, OVA, brefeldin A, collagenase IV, and DNase I were purchased from Sigma-Aldrich. Metrizamide was obtained from Nycomed (Oslo, Norway). The chicken OVA-derived peptide 323–339 (ISQAVHAAHAEINEAGR) and IL-12 were kindly provided by K. Bottomly. IL-4 was obtained from Collaborative Research (Bedford, MA). IL-2 was purchased from Boehringer Mannheim (Mannheim, Germany). Alternatively, for the in vitro experiments, IL-2, IL-4, IFN-{gamma}, and IL-12 were bought from BD PharMingen. In these experiments, the OVA peptide 323–339 was synthesized by the Biochemical Institute, Humboldt-Universität (Berlin, Germany). MACS reagents (columns and beads) were bought from Miltenyi Biotec (Bergisch-Gladbach, Germany). Acetylated low density lipoprotein-Bodipy and CFSE were obtained from MoBiTec (Göttingen, Germany).

Mice

BALB/c mice (8–10 wk of age) were purchased from The Jackson Laboratory (Bar Habor, ME), or from Charles River (Wilmington, MA). DO11.10 mice and DO11.10/TCR {alpha}-chain knockout mice, both expressing a TCR specific for the chicken OVA peptide 323–339 presented in the I-Ad context, were a gift from K. Bottomly (19). Mice were housed and bred at the Yale University Medical School animal facility under pathogen-free conditions in conformance with institutional guidelines for animal care. For the in vitro experiments, all above-mentioned mouse strains were obtained from the Bundesamt für gesundheitlichen Verbraucherschutz und Veterinärmedizin (Berlin, Germany). All animals received humane care according to the criteria outlined in the "Guide for the Care and Use of Laboratory Animals" prepared by the National Academy of Sciences and published by the National Institutes of Health.

Isolation of T cells from the liver, spleen, and lymph nodes

Livers were rinsed in situ with 2 ml prewarmed digestion medium (Bruff’s/5% FCS supplemented with 2 mg/ml collagenase IV and 0.2 mg/ml DNase I) by injection into the portal vein. Organs were removed and ground in a sieve, followed by enzymatic digestion (40 min, 37°C). Lymphoid cells were isolated from the tissue suspension by a one-step density gradient centrifugation in 24% metrizamide. Total numbers of cells isolated from a single liver ranged between 0.5 and 1.5 x 107. This isolation method did not alter the expression of surface markers nor the intracytoplasmic cytokine expression, as shown by control experiments with cells isolated exclusively by density gradient centrifugation (data not shown). Spleens and lymph nodes were disrupted and passed through a fine mesh, resulting in a yield of ~1–1.5 x 108 lymphoid cells/spleen and 3–7 x 107 from the lymph nodes.

In vitro differentiation of Th1 and Th2 cells

Th1 and Th2 effector cells were prepared according to established methods (20). In brief, lymph node cells from TCR-transgenic mice (DO11.10) were cultured on total spleen cells as APCs in complete RPMI 1640/10% FCS supplemented with 5 µg/ml OVA peptide, murine recombinant cytokines, and cytokine Abs. Th1 cells were generated by adding anti-IL-4 (5 µg/ml), IL-12 (10 ng/ml), and IFN-{gamma} (20 ng/ml) to the medium; Th2 cells were cultured in medium supplemented with anti-IL-12 (5 µg/ml), anti-IFN-{gamma} (5 µg/ml), IL-2 (5 ng/ml), and IL-4 (3 ng/ml). Th1 and Th2 cells harvested at day 6 of the culture were predominantly in a resting stage, as defined by an L-selectinhigh, small cell size phenotype (data not shown). Th1 cultures contained between 30 and 80% IFN-{gamma}-producing cells. The percentage of cells expressing large amounts of IL-4 in the Th2 cultures usually ranged between 5 and 25%. Double stains (IFN-{gamma} FITC and IL-4 PE) showed no IFN-{gamma}/IL-4 double-positive cells and virtually no IFN-{gamma} expression in Th2-polarized cells or IL-4 expression in Th1 cells (data not shown).

Ex vivo isolation of mouse sinusoidal endothelial cells by MACS

Liver cell suspension was prepared from 5–10 pooled organs, as described above. Total cells obtained (2–5 x 108/10 livers) were stained with digoxigenized ME-9F1. This mAb (rat IgG2a) binds to mouse endothelial cells, including liver sinusoidal cells, smooth muscle cells, and basal membrane (21). Cells were washed twice, subsequently counterstained with antidigoxigenin MACS beads, and passed over an LS+ MACS column. This sorting technique leads to a yield of 1–5 x 107 cells in the positive fraction and 0.5–1 x 108 negative cells. Sorted cells were cultured overnight, harvested, and used the next day. To control the purity of liver sinusoidal endothelial cells (LSECs),4 aliquots of the sorted fractions were cultured in the presence of acetylated low density lipoprotein-Bodipy. This fluorescence-labeled lipoprotein is exclusively taken up by endothelial cells such as LSECs. Positively sorted cells contained between 90 and 95% LSECs (see Fig. 6Go). Contamination with F4/80-positive v. Kupffer cells and CD11c-expressing dendritic cells was low (1–3 and 3–5%, respectively; data not shown). The negative fraction comprised mainly v. Kupffer cells (up to 50%).



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  		FIGURE 6. FACS staining and MACS sorting of liver-derived APCs with the murine endothelial cell Ab ME-9F1. Surface-stained total liver cell preparations were sorted by MACS, and both the positive as well as the negative fraction were cultured overnight in the presence of acetylated, Bodipy-labeled low density lipoprotein. After 12 h, the cells were additionally stained for the expression of F4/80 as a marker for v. Kupffer cells. A gate was set on vital cells, and 20,000 counts were acquired. Data shown derive from 1 representative experiment of 10.

 
In vitro restimulation of Th1 and Th2 cells on LSECs

In vitro differentiated Th1 and Th2 cells were harvested at day 6 of culture. APCs from the spleen, and LSECs were prepared as described above. A total of 2.5 x 106 T cells was cultured on 1 x 107 APCs in the presence of 5 µg/ml OVA peptide. No additional cytokines or cytokine Abs were added. Cells were harvested at day 6 of the secondary culture, counted, and analyzed for KJ1.26, CD4, annexin V binding, and intracytoplasmic cytokine expression.

Adoptive transfer and in vivo restimulation of cells

In vitro polarized Th1 or Th2 cells were injected into the tail vein of syngeneic BALB/c mice (1 x 107 Th1 or Th2 cells/mouse) at day 0. At day 1 following transfer, mice received a single i.v. injection of OVA (500 µg solubilized in 250 µl sterile PBS) or, in the case of control mice, no Ag. Intravenously administered OVA can be preferentially detected in the liver as compared with lymph nodes and spleen (15- to 20-fold more in the liver), as shown by control experiments with radioactively labeled (125I) OVA (data not shown). At days 1, 3, 5, 15, and 25 after transfer, mice were sacrificed, and lymph nodes (submandibular, axillar, inguinal, and mesenteric), spleen, and liver were removed. Cells isolated from these organs were counted, and the frequency of activated donor cells and transferred cells undergoing apoptosis was determined. Furthermore, cells were functionally analyzed for cytokine expression after Ag-specific restimulation.

Detection of in vivo proliferation by CFSE labeling

Th1 and Th2 cells were differentiated as described above, harvested at day 6, and labeled, as previously described (22). Briefly, cells were suspended in PBS at 1 x 107/ml. For fluorescence labeling, 1 µl CFSE stock solution (5 mM in DMSO) was incubated with 2.5 x 107 cells for 3 min at 37°C.

FACS analysis of surface markers

Aliquots of cells from the different organs were stained with KJ1.26 Cy5 (alternatively KJ1.26 allophycocyanin), CD4 PerCP, and B220 PE/annexin V FITC expression. Annexin V is bound by phosphatidylserine, which is translocated into the outer leaflet of the cell membrane of apoptosing cells. Nonspecific binding was blocked by preincubating the cells with rat IgG (10 µg/ml) and anti-FcII/III. Four-color stained cells were analyzed on a FACSCalibur (BD Biosciences, Heidelberg, Germany) by dual laser technique. Live cells were defined by forward and side scatter, and a combined gate was set on KJ1.26/CD4 double-positive cells. A total of 2,000–10,000 donor cells was acquired. Total counts ranged between 150,000 and 450,000 counts, depending on the frequency of donor cells in the sample. Data were analyzed using CellQuest software (BD Biosciences).

Intracytoplasmic cytokine staining

Aliquots of cells from the liver and lymph nodes were mixed with syngeneic BALB/c spleen cells as APCs (2:1, APCs to isolated cells) and restimulated with the OVA peptide (5 µg/ml). Spleens were restimulated without adding APCs. After 2 h, brefeldin A was added and the cells were incubated for another 4 h. After 6-h total restimulation time, cells were harvested, stained with KJ1.26 and CD4, and fixed in 2% paraformaldehyde for 20 min. Following surface stain and fixation, cells were permeabilized in 0.5% saponin/0.1% BSA and double stained for intracytoplasmic cytokine expression (IFN-{gamma} FITC/IL-4 PE, IL-2 FITC/IL-10 PE). Nonspecific binding was blocked by preincubating the cells with rat IgG (10 µg/ml) and anti-FcII/III. Cells were analyzed using a FACSCalibur (BD Biosciences).

Alternatively, cells were restimulated with PMA/ionomycin, brefeldin A was added after 45 min, and the cells were cultured for a total time span of 5 h.

Data analysis

Data are presented as mean ± SD of mean, and a paired t test was applied. Alternatively, p was determined by a Wilcoxon test. Statistical analyses were performed with Minitab software for MacIntosh (Minitab, State College, PA).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
The liver contains a relatively high frequency of Th1 and Th2 cells, corresponding to an efficient recruitment of activated and effector T cells from the circulation (11, 12, 18, 23). In this study, we aimed first to determine the distribution of resting and in vivo activated effector cells between liver and lymphoid compartments after adoptive transfer. Second, their fate after delivery of Ag by a tolerogenic route was investigated. In vitro polarized, transgenic Th1 and Th2 cells were transferred into syngeneic mice, and animals were i.v. injected 1 day later with a high dose of the respective Ag and subsequently sacrificed at various time points. Donor cells in lymph nodes, spleen, and liver were identified by the clonotype-specific Ab KJ1.26.

Ag delivery leads to accumulation, proliferation, and subsequent deletion of CD4+ effector cells within the liver

Adoptively transferred Th1 and Th2 cells could be detected up to 25 days after injection. Without Ag administration, comparable frequencies of donor cells were found in all of the tissue compartments investigated, and we did not observe consistent differences in the long-term persistence between Th1 and Th2 cells (Table IGo). In contrast, Ag administration at day 1 resulted in a significant increase in the frequencies of Th1 cells in spleen and liver at day 5 after transfer and concomitant depletion in lymph nodes at later time points. However, the effects on Th2 cells were smaller and not significant in the whole set of experiments (Table IGo). The changes in the frequencies of donor cells were accompanied by corresponding alterations in their absolute counts within the respective organs (Fig. 1Go).


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  		Table I. Frequencies of transferred Th1 and Th2 cells in various organs without administration and after injection of Aga

 


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  		FIGURE 1. Kinetics of KJ1.26/CD4-positive donor cells in various organs. In vitro polarized Th1 or Th2 cells were transferred into syngeneic BALB/c mice; at day 1 of transfer, animals received a single i.v. injection of OVA, and control mice received no Ag. Animals were sacrificed at different time points (days 1, 5, 15, and 25) following transfer, organs were removed, and the absolute number of KJ1.26 allophycocyanin/CD4 PerCP-positive donor cells was determined. Graphs in plots show mean ± SD, n = 2–8 mice/time point and organ. Data presented were taken from two to six representative experiments.

 
We then investigated whether i.v. injection of the respective Ag might induce proliferation and subsequent deletion of CD4+ effector T cells within the liver, as has been observed for naive CD4+ and CD8+ T cells (24, 25). By transferring CFSE-labeled effector cells, we observed that Ag administration promotes proliferation of specific Th1 (Fig. 2Go) as well as Th2 effector cells, with Th2 cells displaying a slightly higher proliferation rate as compared with Th2 cells (data not shown). The spontaneous mitosis rate as a parameter of activation as well as the proliferative response toward the Ag were in general higher in the liver as compared with spleen and lymph nodes (Fig. 2Go; data from lymph node not shown). Furthermore, reisolated donor cells were checked for annexin V binding as an early parameter for apoptosis. Apoptosis could be detected as early as 24 h after transfer with comparable rates at days 5 and 15 (data not shown). The frequency of T cells exhibiting signs of apoptosis was generally higher among donor cells isolated from the liver than among cells from lymph nodes or spleen. Ag administration resulted in a further, significant increase of Th1 as well as Th2 cells undergoing apoptosis within the liver (median of two experiments 1.8-fold among Th1 and 2.3-fold increase among Th2 cells, p <= 0.05, n = 2–3 mice/experiment; one single, representative experiment is shown in Fig. 3Go). Taken together, the liver functions as a site of proliferation and apoptosis not only for short-term activated naive T cells, but also for effector cells of both Th1 and Th2 phenotype.



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  		FIGURE 2. Ag-induced proliferation of Th1 cells reisolated from spleen and liver. In vitro differentiated Th1 cells were labeled with CFSE and adoptively transferred, as already described. Donor cells recovered from spleen and liver at day 3 following transfer were analyzed by flow cytometry, and the mean division rate was determined. Data shown derive from one representative experiment of four.

 


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  		FIGURE 3. Ag-induced apoptosis in donor cells reisolated from lymph nodes, spleen, and liver. Adoptive transfer experiments were performed, as already described. Donor cells recovered from various organs at day 15 following transfer were analyzed for the binding of annexin V as a marker for apoptosis. A gate was set on CD4/KJ1.26-positive cells; 2,500–10,000 positive cells were acquired, and the frequency of annexin V FITC-positive donor cells was determined. Data are derived from one representative experiment (day 15) of two and show mean ± SD (n = 2–3 mice/time point and organ; mice were analyzed separately).

 
IFN-{gamma}-producing cells exhibit a time-dependent decrease, whereas IL-4-expressing cells persist within the liver

In addition to deletion, Th1 and Th2 cells might undergo quantitative (anergy) or qualitative (immune deviation) alterations in their cytokine expression following in vivo restimulation under tolerizing conditions. To test this assumption, reisolated donor cells were functionally tested for their ability to produce cytokines in response to Ag. Among Th1 and Th2 cells differentiated in vitro in a single activation/polarization round, only a certain fraction is able to produce IFN-{gamma} or IL-4, respectively. Following adoptive transfer, the IFN-{gamma}-expressing subset among Th1 cells decreased significantly in all organs investigated, with or without Ag injected (Fig. 4Go). In contrast, the reduction of the IL-4-expressing cells was less pronounced among Th2 cells reisolated from lymph nodes and spleen, and not detectable at all among Th2 cells recovered from the liver (Fig. 4Go). In seven independent experiments, the higher level of IL-4- as compared with IFN-{gamma}-producing cells in the liver was highly significant especially after administration of Ag (Table IIGo). Thus, i.v. Ag stimulation results in a conservation of IL-4 production. In contrast, IFN-{gamma} is repressed by deletion and functional down-regulation independent of the Ag.



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  		FIGURE 4. Kinetics of Ag-specific cytokine production in recovered donor cells. Adoptively transferred Th1 and Th2 cells reisolated from various organs at day 5 or 25 after injection were analyzed for Ag-specific cytokine expression by intracytoplasmic staining after Ag-specific restimulation, as already described. Data shown are derived from one representative experiment of four and show mean ± SD (n = 2–3 mice/time point and organ; mice were analyzed separately).

 

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  		Table II. Absolute numbers of IFN-{gamma}- and IL-4-producing cells in various organs without administration and after injection of Aga

 
Furthermore, double staining for some cytokines showed that IL-4 as well as IL-10 expression is restricted to Th2 cells, whereas IFN-{gamma}-producing cells are preferentially found among Th1 cells. IL-2 was only produced by a very small subset of both Th1 and Th2 cells (Fig. 5Go). Thus, our data do not provide any evidence for the occurrence of qualitative changes in the cytokine expression in the sense of an immune deviation or switch to an IL-10-expressing, regulatory phenotype among differentiated Th1 cells.



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  		FIGURE 5. Coexpression of cytokines produced by recovered Th1 and Th2 donor cells. Adoptively transferred Th1 and Th2 cells reisolated from the liver at day 5 after injection were analyzed for Ag-specific cytokine expression by intracytoplasmic staining, as described above. Data shown are derived from one representative experiment of six.

 
LSECs suppress the expansion of IFN-{gamma}-producing cells

The above data suggest a role for the liver in support of IL-4-producing and down-regulation of IFN-{gamma}-expressing cells, but do not provide clues as to the mechanism. We hypothesized that Ag recognition of effector T cells on liver-specific MHCII-positive cells, such as LSECs and v. Kupffer cells, might result in increased apoptosis or down-regulation of IFN-{gamma}-producing cells and preferential survival of IL-4-expressing cells. To test these assumptions, we isolated LSECs ex vivo (Fig. 6Go) and restimulated in vitro differentiated Th1 and Th2 cells on these liver-derived APCs without supplying cytokines or growth factors in addition to the Ag. Spleen cells were used as control APCs. After 6 days, the effector cells were harvested and counted, and the absolute number of cytokine-expressing cells was determined. In these experiments, we observed that IL-4-producing cells grew significantly better on LSECs compared with IFN-{gamma}-positive cells, whereas the opposite could be shown for splenic APCs (p <= 0.05). Furthermore, more IFN-{gamma}-expressing cells could be harvested from cultures activated on spleen APCs than on LSECs (p <= 0.005). Conversly, IL-4-expressing cells exhibited a tendency to expand preferentially on LSECs, yet this effect was not statistically significant with p = 0.06 (Fig. 7GoA). Qualitative changes in the cytokine pattern of cells did not occur, analogous to the in vivo situation. Furthermore, we did not observe any differences in the frequency of annexin V binding, apoptosing cells among Th1 and Th2 cells cultured on either liver-derived or spleen APCs (Fig. 7GoB).



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  		FIGURE 7. In vitro differentiated Th1 and Th2 cells (2.5 x 106 total T cells) were restimulated with OVA peptide on spleen cells and LSECs (1 x 107 APCs) in the absence of added cytokines. After 6 days, the cells were harvested and counted, and the absolute number of IFN-{gamma}-expressing Th1 and IL-4-synthesizing Th2 cells was determined by intracytoplasmic staining. The open bars indicate the absolute number of IFN-{gamma}-positive, the filled bars the absolute number of IL-4-producing cells given into coculture (day 0) and harvested at day 6 after coculture on spleen APCs or LSECs, respectively. Data derive from three to seven independent experiments. *, Difference significant with p <= 0.05; **, p <= 0.005.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In adoptive transfer models with naive CD4+ and CD8+ T cells, tolerance toward endogenous, MHC class I-restricted (viral) Ags can be mediated by apoptosis or induction of an anergic phenotype within the liver (26, 27). Furthermore, previous investigations suggested that the liver might be a site for tolerance induction to exogenous, MHCII-restricted (food) Ags entering the organ in large amounts via the portal circulation from the gut. Yet, in this case mechanisms are less well defined (28, 29, 30). Because CD4+ cytokine-producing effector cells are highly represented within the intrahepatic lymphocyte population and can become preferentially trapped as compared with naive CD4+ T cells (11, 12, 18), we hypothesized that the liver might be a major checkpoint for modulation or deletion of effector cells. To investigate the response of adoptively transferred CD4+ effector cells of Th1 and Th2 phenotype toward tolerance induction, we used an established model for high dose tolerance induction by i.v. injection of Ag (24, 31, 32).

Our results show that at 5 and more days after adoptive transfer, total Th1- and Th2-polarized cells had equally populated lymph nodes, the spleen, and the liver, and remained detectable at least up to day 25, even in the absence of Ag. This basal distribution pattern of adoptively transferred effector T cells was significantly altered by Ag: injected protein increased the number of transgenic T cells at early time points in spleen and liver, predominantly in the case of Th1 cells. At later time points, effector cells became deleted from lymphoid tissues upon exposition to Ag, but were still found in increased numbers in the liver, especially activated effector cells enriched there.

Is accumulation of activated cells within the liver linked to proliferation and deletion of these cells? Evidence for these assumptions is given by our findings that effector cells retrieved from the liver expressed the highest rate of proliferation as compared with lymphoid organs. Conversly, the liver contained the largest percentage of effector cells undergoing programmed cell death without Ag, and strongly increased after i.v. Ag administration. Analogous to what has previously been demonstrated for naive CD4+ and CD8+ T cells after i.v. application of Ag, CD4+ effector cells also undergo an abortive expansion in response to the Ag (25, 32, 33).

To determine the potential of intrahepatic APCs to induce CD4+ T cell proliferation and apoptosis, in vitro experiments were conducted. By restimulating Th1 and Th2 cells on LSECs and splenic APCs, we observed that LSECs are capable of Ag presentation to effector cells, resulting in expansion and concomitant apoptosis. Yet, effector cells cultured on LSECs did not exhibit an increased rate of cell death as compared with splenic APCs. Thus, presentation of Ag on LSECs does not seem to contribute to induction of apoptosis. It is feasible that CD4+ effector cells are driven into apotosis systemically, but accumulate in the liver due to preferential uptake of activated and perhaps apoptosing cells in this organ. Alternatively, it cannot be excluded that LSECs show differential properties under in situ conditions or that other cells of the liver are involved in apoptosis induction. In preliminary experiments, the stimulation of Th1 and Th2 cells on v. Kupffer cells appeared to result in similar rates of expansion and apoptosis compared with LSECs (K. Klugewitz, unpublished data).

Induction of peripheral tolerance is not exclusively dependent on deletion. To investigate whether effector cells undergo functional alterations, namely acquisition of hyporesponsiveness or immune deviation, we followed the ability of the transferred Th1 and Th2 cells to synthesize cytokines for longer periods after transfer. Our data show a strong decrease of the IFN-{gamma}-producing subset among total transferred Th1 cells in all compartments. These data support findings from other authors who have interpreted this effect as a partial reversion of an effector cell to a naive phenotype (34, 35). An alternative hypothesis is that IFN-{gamma}-expressing and nonproducing cells differ in the rate of apoptosis or proliferation, leading to a preferential survival of the nonproducing cells. This interpretation seems to be less likely because preliminary data from our group obtained with IFN-{gamma}-enriched populations (90% IFN-{gamma}-positive cells) show that even under these conditions cytokine production is strongly decreased and that both negative and positive subsets express comparable proliferation rates (K. Klugewitz, unpublished data).

Apart from these quantitative changes, we found no evidence for alterations in the cytokine phenotype of the transferred effector cells as would be predicted by the immune deviation model: transferred Th1 cells in the liver or other compartments had lost the capacity to produce IFN-{gamma} on the long term, but were not found to express IL-4 or IL-10 instead or in addition. These findings are in line with studies showing that the T1 or T2 profile of an ongoing immune reaction toward an endogenous, viral Ag does not undergo any repolarization (36).

In lymphoid tissues, the fraction of IL-4-expressing cells among the total transferred Th2 population also declined, yet to a lesser degree. Strikingly, however, IL-4 production in CD4+ T cells from the liver remained essentially unchanged. Therefore, the ratio of IL-4- to IFN-{gamma}-expressing cells increased steadily in this organ, especially after i.v. Ag delivery. As a consequence, anti-inflammatory phenotypes accumulate and survive, creating an immunosuppressive milieu within the liver, whereas proinflammatory subsets are silenced or die.

Because these effects were focused on the liver, we hypothesized that Ag recognition on liver-derived APCs might result in suppression of IFN-{gamma}-expressing subsets and induce or favor an anti-inflammatory phenotype within recruited effector cells. By restimulating Th1 and Th2 cells on various APCs, we showed that LSECs suppress the expansion of IFN-{gamma}-producing cells compared with splenic APCs. In contrast, LSECs rather promote growth of IL-4-positive cells. Therefore, our data allow the interpretation that loss of IFN-{gamma}-producing Th1 cells and accumulation of anti-inflammatory subsets within the liver might be a locally induced effect. In this respect, our point of view is in line with data from other authors showing that liver-derived APCs such as LSECs and dendritic cells promote an anti-inflammatory phenotype within naive CD4+ T cells, whereas they do not support a Th1 polarization (6, 37). The role of v. Kupffer cells, another major MHCII-positive population that might be potentially involved in the promotion of anti-inflammatory cells, is under current investigation.

In this study, we have defined distinct roles for apoptosis and immune deviation in CD4+ effector T cell liver tolerance. Although activated effector T cells undergo accelerated apoptosis in the liver in the presence of systemically administered tolerogenic Ag, their apoptosis rate is not different between inflammatory and anti-inflammatory subsets. However, in addition to apoptosis, there is a selective loss of effector function in inflammatory Th1 cells located in the liver. This may be induced by interaction of the T cells with LSECs.


   Footnotes
 
1 This work was supported by grants from the Deutsche Forschungsgemeinschaft, Germany, awarded to K.K. (KL 1183/1-1 and 1183/1-2) and A.H. (HA 1505/9-1), and by Grant AI37554 from the National Institutes of Health, awarded to I.N.C. Back

2 Address correspondence and reprint requests to Dr. Katja Klugewitz, Experimentelle Rheumatologie, Charité, c/o Deutsches Rheumaforschungszentrum, Schumannstrasse 21/22, 10117 Berlin, Germany. E-mail address: klugewitz{at}drfz.de Back

3 Current address: David H. Smith Center for Vaccine Biology and Immunology, Aab Institute of Biomedical Sciences, 601 Elmwood Avenue, Box 609, Rochester, NY 14642. Back

4 Abbreviation used in this paper: LSEC, liver sinusoidal endothelial cell. Back

Received for publication March 6, 2002. Accepted for publication June 26, 2002.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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