SEARCH

SEARCH BY CITATION

Abstract

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

The immune response to foreign antigens in the liver is often suboptimal and this is clinically relevant in chronic persistence of hepatotropic viruses. In chronic infection with the hepatitis C virus, activated CD8+ T cells specific for viral epitopes are present in the peripheral blood and the liver, yet viral clearance is unusual. To define the fate of activated CD8+ entering the liver, we developed a mouse model of portal vein injection of activated CD8+ T cells in vivo. Activated CD8+ T cells are retained very efficiently by the liver and undergo an approximately 8-fold expansion in the first 48 hours. This expansion is followed by apoptosis and a decline in numbers of the retained cells over the next 4 days. The presence of high affinity (HA) antigen does not affect the initial retention by the liver but greatly limits the expansion in the first 48 hours by increasing apoptosis of the retained cells. In the absence of Kupffer cells, the initial retention and expansion are unchanged, but HA antigen does not limit the expansion of the liver CD8+ T cell pool. In conclusion, these data identify a previously unknown phase of CD8+ T cell expansion after entering the liver, demonstrate that HA antigen limits the hepatic CD8+ T cell pool by inducing apoptosis, and that this effect requires Kupffer cells. Interfering with antigen presentation by Kupffer cells may be a strategy to limit HA antigen-induced deletion of activated CD8+ T cells entering the liver. (HEPATOLOGY 2004;39;1017–1027.)

The healthy liver is at the interface between the external and internal environments and has a number of unique immunological features. It contains a large and diverse immune cell population, yet immune responses to foreign antigens in the liver are frequently suboptimal.1 Due to its large blood flow and unique anatomy, a large proportion of T cells activated in the lymph nodes (LN), and all T cells activated in the spleen, flow into the liver. Such activated T cells often mount an inadequate effector response to foreign antigens in the liver.2 The reasons for the poor effector response by activated T cells entering the liver are of great clinical relevance in understanding why infection with hepatitis B and C frequently progress to chronicity despite the presence of virus-specific T cells in the peripheral blood and liver.3, 4

The healthy liver is unusual in retaining activated CD8+ T cells entering it. Such retention is due to a combination of unique anatomy, with low flow rates, a very mobile system of tissue macrophages, and the presence of adhesion molecules ICAM-1 and VAP-1.5 Liver retention has a high degree of specificity for activated T cells, and particularly CD8+ rather than CD4+ T cells. On entering the liver, CD8+ T cells are in physical contact with Kupffer cells, and a large proportion undergo apoptosis.5–7 In between the initial retention and final apoptosis, very little is known about activated CD8+ T cells that enter the liver.

In this study, we quantify the expansion of a well-defined effector population of CD8+ T cells on entering the liver. Such a population of activated CD8+ T cells will expand approximately 8-fold in 48 hours, with a gradual reduction over the next 4 days. If the retained CD8+ T cells interact with high affinity (HA) peptide and major histocompatibility complex (MHC) complexes on liver cell populations, the expansion is severely limited, and the removal of the retained CD8+ T cells is accelerated. Previously, we have shown that activated T cells retained in the liver are in physical contact with Kupffer cells. By using mice deficient in mature tissue macrophages, we demonstrate here that the ability of HA peptide to limit the expansion of the effector pool of CD8+ T cells is dependent on mature Kupffer cells. Inhibition of antigen presentation by Kupffer cells may be a therapeutic strategy for minimizing HA antigen-induced apoptosis of activated CD8+ T cells in the liver.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Animals.

C57BL/6J mice and CSF-1 deficient mice (op/op mice)8 were purchased from The Jackson Laboratory (Bar Harbor, ME). A colony of OT-1 T cell receptor (TCR) transgenic mice was maintained on the CD45.1 allotype background. All animals were on the C57BL/6J background and housed in a specific pathogen-free environment, in accordance with institutional guidelines for animal care.

CD8+ OT-1 Lymphocyte Activation and Adoptive Transfer.

Axillary and inguinal LN and spleen were dissected from 6- to 8-week-old OT-1 mice. A cell suspension was obtained by mechanical homogenization under sterile condition. The cells were washed several times with 5% fetal bovine serum (FBS) in phosphate buffered saline (PBS) and resuspended in RPMI (Roswell Park Memorial Institute Media) 1640 (GIBCO, Grand Island, NY) supplemented with 10% FBS, 100 U/ml of penicillin G (SIGMA, St. Louis, MO), 100 μg/ml of streptomycin sulfate (SIGMA), and 50 μM of 2-ME (2-Mercaptoethanol, SIGMA) at the concentration of 2 × 106 cells/ml. HA peptide SIINFEKL (20 nM) was added and incubated under 37°C and 5% CO2. After 3 days, the cells were washed three times with PBS 5% FBS. In some experiments, the cells were incubated at 37°C with 2 μM carboxyfluorescein diacetate (CFSE) (Molecular Probes, Eugene, OR) for 20 minutes and washed for another three times. By day 3 of activation in vitro, more than 95% of these cells were CD8+, and more than 99% CD8+ cells were CD25 high. This confirmed that we had a very pure population of highly activated CD8+ T cells.

Age- and gender-matched C57BL/6J mice and CSF-1 deficient mice were used as recipients. Under ketamine/xylazine anesthesia, the abdominal cavity of the mice was exposed by a midline incision. Activated cells (5 × 106) prepared as described above were injected directly into the portal vein. After the injection, the abdominal cavity of mice was closed using proline sutures and Autoclips (Becton Dickinson, Parsippany, NJ). Twenty minutes before portal vein injection, each recipient mice received an intraperitoneal (ip) injection of either 25 nmol of the HA SIINFEKL in PBS, 25 nmol of the control peptide (ovalbumin 323-339), or PBS alone. Peptide or PBS were injected ip every 24 hours subsequently.

Cell Isolation.

At various time points between 10 minutes and 7 days after injection of activated CD8+ OT-1 cells, the mice were sacrificed and the lymphocytes from the liver, blood, spleen, and lungs were isolated. In brief, mice were anesthetized, heparinized, and exsanguinated by cutting the abdominal aorta and vena cava. The liver was perfused immediately with digestion buffer consisting of Bruff's medium containing 0.05% collagenase IV (SIGMA), 0.002% DNase I (SIGMA), and 5% FBS. After perfusion, the liver was dissected out of the abdominal cavity and homogenized. The homogenized liver was incubated in 10 ml digestion buffer at 37°C for 40 minutes in a shaking water bath. After removing hepatocytes and cell clumps by centrifugation at 20g, the supernatant was then centrifuged at 500g and a pellet was collected. The pellet was suspended in 22% Opti-prep (Axis-Shield, Oslo, Norway) and centrifuged at 1,500g. The cells at the interface were collected, washed, and analyzed. Lymphocytes from spleen, LN, and blood were obtained by mechanical homogenization. Red blood cells (RBC) were lysed using Ack Lysing Buffer (SIGMA) for spleen and blood.

Staining Reagents and Flow Cytometric Analysis.

The antibodies used for staining were anti-TCRαβ (clone H57-597), anti-CD8 (clone 53-6.7), anti-CD4 (clone H129.19), anti-CD45.1 (clone A20), and anti-CD25 (clone PC61). Avidin-fluorescein-5-isothiocyanate, phycoerythrin, or allophycocyanin was used when biotinylated antibody was used at primary staining. All these reagents were purchased from BD Biosciences (San Jose, CA). For terminal deoxynucleotidyl transferase mediated dUTP biotin nick end labeling (TUNEL) staining, Apop Tag Fluorescein Direct In Situ Apoptosis Detection Kit (Intergen, NY) was used according to the manufacturer's instructions. Cells (1–3 ×106) obtained from each organ were stained using reagents listed previously. After staining, the samples were fixed with 2% paraformaldehyde.

The samples were analyzed using FACS Calibur flow cytometer (BD Bioscience) and CellQuest software (BD Bioscience). Each sample data was first gated based on forward scatter and side scatter on lymphocytes and then further analyzed for fluorescence.

In Vivo Cytotoxic T Lymphocyte (CTL) Assay.

Target cells for in vivo evaluation of cytotoxic activity were prepared as described in detail elsewhere.9 Briefly, spleen and LN cells were isolated from age- and gender-matched C57BL/6 mice and RBC were lysed. The cells were washed and divided into two populations. One population was pulsed with 1 μM of OT-1 specific peptide SIINFEKL, whereas the other was left unpulsed at 37°C for 1 hour. The peptide-pulsed population was labeled with high (5 μM) CFSE concentration and the unpulsed population was labeled with low (0.5 μM) concentrations. Both population were then washed five times to remove the residual peptide and CFSE. The cells from each population were counted, mixed in a 1:1 ratio, and 2 × 107 of the cell mixture (in 0.5 ml PBS) was injected into the tail vein of mice that had portal vein injections of PBS (control group) or portal vein injections of activated CD8+ OT-1 cells (test group) 36 hours before. Specific in vivo cytotoxicity was determined by analyzing isolated lymphocytes from liver, spleen, and LN of the recipient mice 12 hours after injection of the target cells using FACS. The target and control cell population can be detected separately at different CFSE fluorescence intensities. Specific target cell lysis was described using a value estimated as (1 – % SIINFEKL pulsed cells / % non-pulsed cells) × 100. To minimize the subset difference among target cells in each organ and/or each mice, the isolated cells were further stained with anti-TCR antibody and gated on TCR positives at FACS analysis.

Statistical Analysis.

Mean values ± SE for various groups were calculated and statistical significance determined using the Student's t test.

Results

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Efficient Activation of OT-1 In Vitro.

Culture of naive OT-1 cells from the spleen and LN with the HA peptide (SIINFEKL) for 3 days results in efficient activation of the OT-1 CD8+ T cells (Fig. 1A and B). In addition, by day 3, the cultured cell population had a purity of greater than 96% activated OT-1 CD8+ T cells.

thumbnail image

Figure 1. In vitro activation of OT-1 CD8+ T cells. (A) FACS plots of CD8+ and CD25+ expression on OT-1 LN and spleen cells before activation. (B) FACS plots of the same population of OT-1 LN and spleen cells after 3 days of culture with HA peptide (SIINFEKL) at a concentration of 20 nM, under standard culture conditions. Note that after 3 days of culture the population is 97% pure for activated CD8+ T cells. After 3 days, the cells were washed to remove and residual peptide and adoptively transferred into control and test populations via the portal vein.

Download figure to PowerPoint

Activated CD8+ T Cells Are Efficiently Retained by the Healthy Liver.

We previously showed in a single pass perfusion model that the normal liver selectively retains activated CD8+ T cells in an antigen non-specific manner. To test the efficiency of activated CD8+ T cell retention in vivo, activated CD8+ T cells were directly injected into the portal vein of normal B6 mice. Ten minutes after injection, the percentages of donor CD8+ T cells in various organs were determined by cell isolation and FACS analysis.

In a direct confirmation of the single pass perfusion model, 10 minutes after injection, donor cells were found in large numbers in the liver and virtually none were present in other organs including the blood, lung, or spleen (Fig. 2A–D) and also draining or distal LN. This distribution of portal vein injected activated CD8+ T cells staying in the liver was essentially unchanged for 24 hours after cell injection (Fig. 2E–H).

thumbnail image

Figure 2. Distribution of activated CD8+ T cells 10 minutes and 24 hours after transfer via portal vein. The LN and spleen cells derived from OT-1 mice were stimulated in vitro for 3 days using a HA peptide SIINFEKL. Activated CD8+ OT-1 cells (5 × 106) were injected via the portal vein into wild type littermates. Ten minutes or 24 hours after transfer, various organs from these mice were removed and lymphocytes were isolated. Donor cells were distinguished from resident cells by staining with anti-CD45.1 antibody, antibody against the donor cell surface marker, and analyzed using flow cytometry (FACS). FACS plots indicating percentage of donor CD8+ T cells population (upper right) in comparison with resident CD8+ cells population (upper left) at 10 minutes and 24 hours after the injection are shown from liver, blood, lung, and spleen.

Download figure to PowerPoint

Increase in the OT-1 CD8+ T Cell Population Retained by the Liver.

By 48 hours, there is a substantial increase in the percentage of OT-1 CD8+ T cells in the liver compared to the initial retention at 10 minutes (Fig. 3). On average, the number of OT-1 CD8+ T cells increased 8-fold in 48 hours. This increase between the 10 minute and 48 hour time points could have occurred due to intra-hepatic division of OT-1 CD8+ T cells retained in the liver, or due to their exiting from the liver, dividing extrahepatically, and then trafficking back to the liver. But because only a very few donor cells were seen outside the liver during the early time course after portal vein injection (Fig. 2), the latter possibility is unlikely. Between days 2 and 7 after injection, the percentage of OT-1 CD8+ T cells in the liver decreased substantially.

thumbnail image

Figure 3. The percentage of activated donor CD8+ T cells in the liver increases during the first 48 hours after injection, but the presence of the HA peptide in liver reduces this phenomenon. Activated CD8+ OT-1 cells were injected into the portal vein as described in Fig. 2. Twenty minutes before portal vein injection and every 24 hours subsequently, recipient mice were given ip injections of 0.25 ml of 100 μM OT-1 HA peptide SIINFEKL in PBS or PBS alone as a control. At various time points the liver was removed and lymphocytes were isolated. The cells were stained with anti-CD45.1 and anti-CD8 antibodies and analyzed using FACS. FACS plots show percentage of donor and recipient CD8+ T cells within the liver at 10 minutes, 48 hours, and 96 hours time point.

Download figure to PowerPoint

Presence of HA Antigen in the Liver Severely Limits the Degree of Expansion of OT-1 CD8+ T Cells.

To study whether the presence of HA antigen in liver affects the fate of retained cells, recipient mice were injected with the HA peptide for the OT-1 T cell receptor (SIINFEKL) 20 minutes before injection of activated OT-1 CD8+ T cells into the portal vein. At 10 minutes after injection of OT-1 CD8+ T cells, there was no difference between the HA peptide-injected mice and the PBS-injected controls (Fig. 3A and D). This demonstrates that the presence of HA peptide does not affect the initial retention of activated OT-1 CD8+ T cells by the liver. By 48 hours after transfer of OT-1 CD8+ T cells there is a significant difference in the percentage of OT-1 CD8+ T cells in the livers with and without HA peptide (Fig. 3B and E). The increase in OT-1 CD8+ T cells in the first 48 hours after they enter a liver without HA peptide is severely limited in the presence of HA peptide. From 14 experiments and 5–15 animals for each time point, this phenomenon was confirmed and statistically evaluated (Fig. 4). There was no difference between the control ovalbumin peptide (ova 323–339) and the PBS-injected mice. Each value (mean ± SE) was 10 minutes; PBS: 4.214 ± 1.429 versus HA peptide: 4.970 ± 1.512 (NS), 48 hours; PBS: 35.797 ± 4.801 4 versus HA peptide: 15.242 ± 3.711 (P < .005), 96 hours; PBS: 16.772 ± 2.837 versus HA peptide: 2.200 ± 1.004 (P < .001), day 7; PBS: 10.220 ± 4.220 versus HA peptide: 1.737 ± 0.756 (NS). In summary, this shows that the approximately 8-fold increase that occurs in the pool of activated OT-1 CD8+ T cells in the first 48 hours is severely limited to an approximately 3-fold increase if HA peptide is present but is unaltered in the presence of a control ovalbumin peptide.

thumbnail image

Figure 4. (A) The in vivo dynamics of activated CD8+ T cells retained in livers injected with PBS, HA peptide, or control peptide. At various time point after portal vein injection of activated CD8+ T cells, each liver were taken out and number of donor CD8+ T cells within the liver were analyzed as in Fig. 3. Each plot shows mean ± SE of the percentages of donor CD8+ T cells among liver lymphocytes in either PBS, HA peptide, and control peptide group at each time point (10 minutes, 48 hours, 96 hours, and day 7 are shown). (B) OT-1 cell percentages in the spleens of the same animals as in Fig. A showing significant decrease in OT-1 cells in response to HA but not control peptide.

Download figure to PowerPoint

Transferred OT-1 CD8+ T Cells Are Functional CTLs In Vivo.

To establish that the portal vein injected CD25+ CD8+ OT-1 cells had cytotoxic function, an in vivo CTL assay was performed. Figure 5(A–C) shows data from control mice that did not receive any CD8+ OT-1 cells, but did receive CFSE labeled T cell populations (CFSE high loaded with HA peptide and CFSE low not loaded with peptide). There is no loss of peptide loaded CFSE high target cells. Figure 5(D–F) shows data from mice that received CD8+ OT-1 cells and CFSE labeled T cell populations (CFSE high loaded with HA peptide and CFSE low not loaded with peptide). There is loss of the peptide loaded CFSE high cell population, demonstrating cytotoxic function of the injected CD8+ OT-1 cells. Liver, spleen, and LN were analyzed separately and the degree of loss of target cells was similar and there was no difference in cytotoxic activity among these organs.

thumbnail image

Figure 5. Activated CD8+ OT-1 T cells possess CTL function in vivo. LN and spleen cells from wild type littermates were pulsed with OT-1 specific HA peptide, stained with a high concentration of CFSE, and used as a target cell population. A control cell population was prepared using the same source of cells, but without peptide pulsing, and with CFSE staining at a low concentration. The two populations were mixed in a 1:1 ratio, and injected into the tail vein of mice that had been injected with PBS in the portal vein (A–C), or with activated CD8+ OT-1 cells 36 hours before (D–F). The liver, spleen, and LN from mice were removed 48 hours after the portal vein injection of PBS or activated CD8+ OT-1 cells, and lymphocytes were isolated and stained with anti-TCR antibody and analyzed using FACS. The histogram shows the frequency of remaining target cells among T cells in each organ. In this figure, peptide-pulsed cell populations are CFSE high, while peptide non-pulsed cell populations are CFSE intermediate. The upper row (A–C) shows the target and control cell profile in mice without the transfer of activated CD8+ OT-1 cells (indicated as CTL –), while the lower row (D–F) is from the mice portal vein injected with activated CD8+ OT-1 cells (indicated as CTL +). Percentages of specific target cell lysis is estimated as (1 – % peptide pulsed cells / % peptide non-pulsed cells) × 100 in the activated cells transferred mice. Data show a result of one representative experiment from three.

Download figure to PowerPoint

OT-1 CD8+ T Cells Undergo Proliferation and Apoptosis, With Enhancement of the Apoptosis by HA Peptide.

To test that the increase in the number of OT-1 CD8+ T cells occurs via cell division, the cell membrane dye CFSE was used. CFSE binds to cell surface protein and decreases in intensity as the cells divide. Thus, proliferation in vivo can be demonstrated. Activated OT-1 CD8+ T cells were labeled with CFSE and injected into the portal vein of mice previously injected ip with PBS or HA peptide. After 48 hours, liver lymphocytes were isolated and the CFSE intensity of donor cells was analyzed by FACS analysis by gating on CD45.1+ and CD8+ cells. This revealed an approximately 30-fold decrease in CFSE intensity in the donor cells in the HA peptide- and PBS-injected groups during the first 48 hours (Fig. 6). This demonstrates that retained activated CD8+ T cells do proliferate in the liver and this is most likely the cause of the dramatic increase of donor cells seen in the liver after 48 hours. The decline of CFSE intensity was identical in PBS- and HA-peptide injected mice, suggesting that the degree of cell division is not significantly affected by the interaction of the retained cells with HA peptide.

thumbnail image

Figure 6. The increase in the number of activated CD8+ T cells retained in the liver is accompanied with a decline of CFSE level in these cells. CD8+ OT-1 cells were stimulated in vitro for 3 days using the HA peptide. They were stained with CFSE, washed, and 5 × 106 cells were injected via the portal vein into wild type littermates. Twenty minutes before portal vein injection and after 24 hours, each recipient mouse was given ip injection of HA peptide SIINFEKL or the same volume of PBS as a control. At 48 hours, livers were taken out and lymphocytes were isolated. The cells were stained with anti-CD45.1 and anti-CD8 antibodies and analyzed using FACS. The CFSE intensity of donor cells was viewed by gating on CD45.1+ CD8+ cells. Data are from one representative experiment of four, all of which showed the similar result.

Download figure to PowerPoint

Death of donor cells was quantitatively analyzed looking at apoptotic cells detected by TUNEL method. Figure 7 shows TUNEL staining for the CD45.1 donor T cells isolated from livers of mice injected with PBS or HA peptide. The presence of HA peptide in the liver resulted in significantly greater percentage of the donor cell being TUNEL positive 48 hours after injection (Fig. 7). The possibility that donor T cells escape more out of the liver when the HA peptide is present is unlikely because significantly fewer donor T cells were detected in the HA peptide given group in every extra-hepatic site at any time point (Fig. 8).

thumbnail image

Figure 7. High affinity (HA) peptide increases the apoptosis of CD8+ T cells in the liver. CD8+ OT-1 cells were stimulated in vitro for 3 days using the HA peptide. These cells (5 × 106) were injected from portal vein into wild type littermates. Twenty minutes before portal vein injection and after 24 hours, each recipient mouse was given ip injection of HA peptide SIINFEKL or the same volume of PBS as a control. At 48 hours, the liver was removed and lymphocytes were isolated. The cells were stained with anti-CD45.1 antibodies and TUNEL staining to detect apoptosis. The samples were analyzed using FACS. The TUNEL staining on donor cells was viewed by gating on CD45.1+ cells. (A) and (B) show one set of representative data out of 9 mice in four experiments. (C) shows a summary of all the data.

Download figure to PowerPoint

thumbnail image

Figure 8. Few donor CD8+ OT-1 cells were present in extra-hepatic sites in the presence of HA peptide. CD8+ OT-1 were activated in vitro with the HA peptide and 5 × 106 cells were injected from portal vein into wild type littermates. Intraperitoneal injection of either HA peptide or PBS were given to each recipient mouse 20 minutes before portal vein injection and every 24 hours subsequently. At various time points, non-liver organs were taken out and lymphocytes were isolated. They were stained and the percentages of donor CD8+ T cells in those organs were analyzed by FACS. Each plot shows mean ± SE of the percentages of donor CD8+ T cells. The result from the blood, spleen, and lung of either peptide or PBS given group at indicated time point are shown.

Download figure to PowerPoint

HA Peptide-Induced Apoptosis of Intrahepatic CD8+ T Cells Is Dependent on Kupffer Cells.

We previously showed that the majority of activated CD8+ T cells retained in the liver are in contact with Kupffer cells. In addition, we also demonstrated that ICAM-1, which is present on Kupffer cells and sinusoidal endothelium, is required for efficient retention of activated CD8+ T cells by the liver. To test the hypothesis that Kupffer cells are required for the retention and the antigen-induced apoptosis of activated CD8+ T cells, we studied the retention and antigen-induced deletion of activated OT-1 CD8+ T cells in the livers of wild-type and colony stimulating factor 1 (CSF-1) deficient mice. CSF-1 deficient mice (op/op mice) lack CSF-1 due to an inactivating frameshift mutation in the coding region of the CSF-1 gene.8 CSF-1 is essential for development of tissue macrophages, and CSF-1 deficient mice have no mature functional tissue macrophages including Kupffer cells. Total Kupffer cell numbers are less than 30% compared to hemizygous littermates, and the Kupffer cells present are small and unable to phagocytose.8 Using the CSF-1 deficient mice as recipients, an experiment was designed in order to reveal the role of Kupffer cells in liver retention and deletion of activated CD8+ T cells.

Activated OT-1 CD8+ T cells were transferred via the portal vein into CSF-1 deficient mice or wild type littermates as control that had received ip injections of PBS or HA peptide. At 10 minutes, there was no difference between CSF-1 deficient mice and wild type littermates in the PBS- and peptide-injected groups (Fig. 9A), indicating no difference in the initial retention by liver. After 48 hours, there was no difference between CSF-1 deficient mice and wild type mice when the HA antigen was not present. Both showed a similar increase in the percentages of donor cells in the liver (Fig. 9A). This demonstrates that the initial retention and expansion is not dependent on Kupffer cells. In the HA peptide-injected mice there was significant difference between the wild type and CSF-1 deficient mice. In wild type mice, the presence of HA peptide severely limited the increase of the OT-1 CD8+ T cells at 48 hours (as demonstrated in Fig. 4A), but in the CSF-1 deficient mice, HA peptide was unable to limit the increase in OT-1 CD8+ T cells (Fig. 9A) (% of donor CD8+ T cells in the liver lymphocyte pool. HA peptide-injected wild type: mean ± SE, 5.4 ± 2.244, P < .02, HA peptide-injected CSF-1 deficient 28.259 ± 6.443). This difference strongly suggests that the mechanism of HA peptide-induced CD8+ T cell death is lacking in CSF-1 deficient mice, and we propose it is due to the absence of Kupffer cells in these mice. TUNEL staining of donor cells in the wild type and CSF-1 deficient mice receiving HA peptide did not reveal any significant difference between the two groups (data not shown). We cannot, therefore, demonstrate that the lack of peptide-induced reduction in the CSF-1 deficient mice injected with peptide was due to less apoptosis, but because the mechanism for phagocytosis of apoptotic cells may be also reduced in CSF-1 deficient mice, it is difficult to interpret TUNEL data.

thumbnail image

Figure 9. (A) Deletion of activated CD8+ OT-1 cells by HA peptide does not occur in CSF-1 deficient mice that lack Kupffer cells. CD8+ OT-1 cells were activated in vitro with the HA peptide SIINFEKL. These cells (5 × 106) were injected from portal vein into CSF-1 deficient mice or wild type littermates as control. The CSF-1 deficient and wild type mice received ip injections of PBS or HA peptide at 20 minutes before portal vein injection and every 24 hours subsequently. At various time points, the liver was taken out and lymphocytes were isolated. The cells were stained and analyzed using FACS. Each plot shows mean ± SE of percentage of donor CD8+ cells in the four groups (wild type mice with PBS, wild type mice with peptide, CSF-1 deficient mice with PBS, CSF-1 deficient mice with peptide) at 10 minutes and 48 hours after injection of activated CD8+ OT-1 cells. The result is from 4 individual experiments and 4–7 mice for each plots. (B) No escape of donor T cells from the liver of CSF-1 deficient mice was seen in the presence of HA antigen. Each plot shows mean ± SE of percentage of donor CD8+ T cells in the spleen of four groups at each time point (10 minutes and 48 hours are shown).

Download figure to PowerPoint

Almost no donor cells are detectable in the blood, lungs, or spleen at 10 minutes after injection (Fig. 2A–D). The distribution of donor cells in the blood, lungs, and spleen in CSF-1 deficient mice were identical to wild type, suggesting that the absence of functional Kupffer cells has minimal effect on the ability of the liver to retain activated CD8+ T cells entering it. At 48 hours, when the HA antigen is not present, some donor cells start to appear in other organs, including spleen (Fig. 8A–C), but there were no differences between CSF-1 deficient mice and wild type mice (Fig. 9B). The presence of HA peptide results in very few donor cells being detected in the blood, lungs, and spleen of wild type mice (Fig. 8). This may be because the hepatic retention and apoptosis in the presence of peptide is so efficient that they do not exit the liver, or because they undergo apoptosis outside of the liver due to interaction with HA peptide. In contrast to the liver, where there was a discordance in the percentage of donor cells in the wild type peptide-injected and CSF-1 deficient peptide-injected groups, in the blood, lungs, and spleen almost no donor cells were present in the wild type and CSF-1 deficient peptide-injected groups (Fig. 9B).

Discussion

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

The liver contains a large and varied population of immune cells and has unique functional features.10, 11 During in vivo T cell activation, there is massive accumulation of the apoptosing T cells in the liver.12 This has been confirmed in a wide variety of models ranging from transgenic animals, to adoptive transfer studies, to viral infection of unmanipulated wild type mice.13–15 The demonstration in a single pass perfusion model, of the ability of the healthy liver to retain activated CD8+ T cells further suggested that the liver functions very differently from other non-lymphoid organs.5 Collectively, it has become clear that the healthy liver retains activated CD8+ T cells, and many or most of these cells undergo apoptosis. These two facts are consistent with a number of hypothesis, with two contrasting views being 1) the liver is trapping T cells that are programmed for apoptosis (elephants graveyard model) or 2) the liver is retaining T cells and inducing apoptosis of the T cells that interact with cognate antigen (responder trap).16

In this study, we provide detailed information on the fate of T cells retained by the liver. From the previous adoptive transfer of activated T cells into the systemic circulation, it has not been possible to address in detail issues of liver T cell retention, function, proliferation, and apoptosis because it is not known when the adoptive transferred population actually enters the liver.12, 17 To clarify this we used a strategy of portal vein injection, giving us an exact time of T cell entry into the liver. Our data confirm the previous observations that activated CD8+ T cells entering the liver are retained and undergo substantial apoptosis.5 In between these two events there is not uniform apoptosis and decline in the retained population. Rather, there is significant expansion that peaks at approximately 8-fold by day 2 and then a gradual decline (Fig. 4). The decline is associated with apoptosis of the retained cells, but the gradual increase in the percentage of donor cells in the LN and spleen as they are decreasing in the liver suggests that at least some of the retained cells escape from the liver. So it is a graveyard, but death is preceded by division and some cells escape the graveyard altogether.

The consequences for an activated CD8+ T cell entering a liver containing HA peptide are quite clear. There is no difference in retention, with virtually complete initial retention by the liver regardless of whether the cells entering the liver do or do not interact with HA peptide (Fig. 3). However the expansion of the retained population is significantly curtailed if the activated T cells interacts with HA peptide on a liver cell population. The presence of HA peptide diminishes the peak size of the retained cell population by approximately half, and we believe this is mostly due to increased apoptosis of the retained cells (Fig. 7). These data are consistent with the responder trap hypothesis. This mechanism will apply to all T cells activated in the spleen because they exit into the portal circulation. These findings were obtained using HA peptide, and the response to low affinity agonists, or partial antagonists, is yet to be determined.

By day 7, the presence of HA peptide results in approximately 6-fold fewer donor cells in the liver (Fig. 4). Similarly, in the blood, lung, and spleen, there are many fewer donor cells in the presence of HA peptide (Fig. 8). We infer that in the presence of HA peptide, activated CD8+ T cells are retained by the liver and undergo massive apoptosis, leaving very few cells to exit into the circulation. The proposed mechanism for the enhanced retention is the upregulation of LFA-1 after engagement of the T cell receptor.18 In fact, interaction of LFA-1 with ICAM-1, in the absence of adequate costimulation, may also result in apoptosis, particularly of CD8+ T cells.19–21 We, however, cannot exclude that the decrease of cells in the spleen, LN, and lungs of the peptide-injected animals occurs due to apoptosis at these sites rather than the liver. The numbers of donor cells outside the liver were too few to generate consistent TUNEL data. Our peptide-injected model does not have the tissue specificity of antigen presentation, but is a relevant model for many conditions, such as viral hepatitis, in which viral antigen is distributed widely throughout the body, and virus-specific T cells enter the liver.

It is of great interest to identify the cellular and molecular mechanism required for peptide-induced deletion of activated CD8+ T cells retained by the liver. From bone marrow chimera studies we have shown that antigen presentation on bone marrow derived cells enhances peptide-induced deletion of activated CD8+ T cells.14 To test if Kupffer cells are the bone marrow cell population enhancing apoptosis, it was necessary to use a mouse model lacking Kupffer Cells. A number of maneuvers have been used to impair Kupffer cell function in vivo, including silica and gadolinium chloride injections.22–24 These strategies have significant drawbacks. First, they are reasonably good at impairing Kupffer cell phagocytosis but do not physically remove mature Kupffer cells.25 Second, they are associated with Kupffer cell activation with release of cytokines, and this has unpredictable effects. To avoid these issues, we made use of the natural mutant mice lacking CSF-1.8 In these mice, the maturation of all tissue macrophages is grossly impaired and mature Kupffer cells are not present in the liver.26

In wild type mice, the presence of HA peptide dramatically reduces the number of OT-1 CD8+ T cells at day 2 by enhancing intrahepatic apoptosis of these cells (Figs. 3 and 4). In contrast to wild type mice, the presence of peptide in the liver of CSF-1 deficient mice does not result in any decrease in the number of OT-1 CD8+ T cells. This strongly supports the hypothesis that antigen presentation by Kupffer cells is required for peptide-induced deletion of the retained activated OT-1 CD8+ T cells. It may therefore be possible to selectively interfere with antigen presentation by Kupffer cells, thus blocking the peptide-induced CD8+ T cell deletion without interfering with the CD8+ T cell effector response against hepatocytes. This is therapeutically desirable in situations of suboptimal intrahepatic immune responses, such as chronic infection with hepatitis B and C viruses. The ability of Kupffer cells to induce apoptosis of T cells in vitro by a CD95 dependent mechanism, and the inability to induce oral tolerance in gadolinium chloride treated animals, is consistent with our hypothesis.27–29

These data support a model of hepatic antigen presentation in which antigen presentation by hepatocytes and endothelium leads to conventional CD8+ T cell effector function with release of cytokines and cytotoxic function. Antigen presentation by Kupffer cells, however, enhances CD8+ T cell apoptosis, thus limiting the overall hepatic CD8+ T cell response. Kupffer cells have a number of unique features that makes them good candidates for inducing antigen-specific deletion in activated CD8+ T cells. They are highly mobile and phagocytic.30 In addition, Kupffer cells possess a variety of molecules that may induce CD8+ T cell apoptosis. These include the production of TNF-α, CD95-ligand, galectin-1, and indolomine deoxygenase (IDO).28, 31–33 Of these, at least TNF-α and CD95-ligand are upregulated by the interferon-gamma that is produced in abundance by activated CD8+ T cells. The role of antigen recognition by activated CD8+ T cell in inducing apoptosis may be to increase the affinity of LFA-1 and induce stronger binding to Kupffer cell ICAM-1, or it may be to increase production of interferon gamma by the CD8+ T cell, resulting in upregulation of the Kupffer cell apoptotic mechanisms mentioned previously. After induction of apoptosis, Kupffer cells are able to phagocytose apoptotic cells, thus ensuring efficient removal of the CD8+ T cell apoptotic bodies.34, 35 All these features ensure that Kupffer cells are uniquely poised to sample the entire immunogenic environment of the liver, present it to activated CD8+ T cells, induce their apoptosis, and also phagocytose the apoptotic debris.

The presence of multiple mechanisms on Kupffer cells for the induction of apoptosis complicates the prospect of inhibiting CD8+ T cell apoptosis. Because antigen presentation is required for Kupffer cell induced apoptosis, it may be possible to block CD8+ T cell apoptosis by interfering with class I MHC processing in Kupffer cells rather than trying to block all the pro-apoptotic signals. A number of stages of the class I processing pathway are potentially amenable to blockage. One approach is the delivery of inhibitors of proteasomes to Kupffer cells to block class I processing.

In summary, we have shown that between hepatic retention and apoptosis, the pool of retained activated CD8+ T cells undergoes significant expansion when they enter a liver that does not contain HA peptide. Such expansion does not occur in the presence of HA peptide due to a high degree of apoptosis. In the absence of Kupffer cells, the retention of activated CD8+ T cells by the liver is not altered, but HA peptide does not limit the size of the retained CD8+ T cell pool.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

The authors are grateful to Richard Flavell for invaluable mentorship, and Nicholas Crispe for helpful discussion.

References

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  • 1
    Calne RY, Sells RA, Pena JR, Davis DR, Millard PR, Herbertson BM, Binns RM, et al. Induction of immunological tolerance by porcine liver allografts. Nature 1969; 223: 47.
  • 2
    Bertolino P, Heath WR, Hardy CL, Morahan G, Miller JFAP. Peripheral deletion of autoreactive CD8+ T cells in transgenic mice expressing H-2Kb in the liver. Eur J Immunol 1995; 25: 19321942.
  • 3
    Lauer GM, Ouchi K, Chung RT, Nguyen TN, Day CL, Purkis DR, Reiser M, et al. Comprehensive analysis of CD8+-T-cell responses against hepatitis C virus reveals multiple unpredicted specificities. J Virol 2002; 76: 61046113.
  • 4
    Wong DK, Dudley DD, Afdhal NH, Dienstag J, Rice CM, Wang L, Houghton M, et al. Liver-derived CTL in hepatitis C virus infection: breadth and specificity of responses in a cohort of persons with chronic infection. J Immunol 1998; 160: 14791488.
  • 5
    Mehal WZ, Juedes AE, Crispe IN. Selective retention of activated CD8+ T cells by the normal liver. J Immunol 1999; 163: 3202.
  • 6
    Qian S, Lu L, Fu F, Li Y, Li W, Starzl TE, Fung JJ, et al. Apoptosis within spontaneously accepted mouse liver allografts. J Immunol 1997; 158: 465.
  • 7
    Iwakoshi NN, Goldschneider I, Tausche F, Mordes JP, Rossini AA, Greiner DL. High frequency apoptosis of recent thymic emigrants in the liver of lymphopenic diabetes-prone BioBreeding rats. J Immunol 1998; 160: 5838.
  • 8
    Yoshida H, Hayashi SI, Kunisada T, Ogawa M, Nishikawa S, Okamura H, Sudo T, et al. The murine mutation osteopetrosis is in the coding region of the macrophage colony stimulating factor gene. Nature 1990; 345: 442.
  • 9
    Nelson D, Bundell C, Robinson B. In-vivo cross-presentation of a soluble protein antigen: kinetics, distribution, and generation of effector CTL recognizing dominant and subdominant epitopes. J Immunol 2000; 165: 61236132.
  • 10
    Mehal WZ, Azzaroli F, Crispe IN. Immunology of the healthy liver: old questions and new insights. Gastroenterology 2001; 120: 250260.
  • 11
    Crispe IN. Hepatic T cells and liver tolerance. Nature Rev Immunol 2003; 3: 5162.
  • 12
    Huang L, Soldevila G, Leeker M, Flavell RA, Crispe IN. The liver eliminates T cells undergoing antigen-triggered apoptosis in vivo. Immunity 1994; 1: 741.
  • 13
    Huang L, Sye K, Crispe IN. Proliferation and apoptosis of B220+CD4-CD8-TCR alpha beta intermediate cells in the liver of normal adult mice: implications for lpr pathogensis. Int Immunol 1994; 39: 259267.
  • 14
    Mehal WZ, Azzaroli F, Crispe IN. Antigen presentation by liver cells controls intrahepatic T cell trapping, whereas bone marrow-derived cells preferentially promote intrahepatic T cell apoptosis. J Immunol 2001; 167: 667673.
  • 15
    Belz GT, Altman JD, Doherty PC. Characteristics of virus-specific CD8+ T cells in the liver during the control and resolution phases of influenza pneumonia. Proc Natl Acad Sci U S A 1998; 95: 13812.
  • 16
    Crispe IN, Mehal WZ. Strange brew: T cells in the liver. Immunol Today 1996; 17: 522.
  • 17
    Klugewitz K, Blumenthal-Barby F, Schrage A, Knolle PA, Hamann A, Crispe IN. Immunomodulatory effects of the liver: deletion of activated CD4+ effector cells and suppression of IFN-gamma-producing cells after intravenous protein immunization. J Immunol 2002; 169: 24072413.
  • 18
    Dustin ML, Springer TA. T-cell receptor cross-linking transiently stimulates adhesiveness through LFA-1. Nature 1989; 341: 619.
  • 19
    Deeths MJ, Mescher MF. ICAM-1 and B7-1 provide similar but distinct costimulation for CD8+ T cells, while CD4+ T cells are poorly costimulated by ICAM-1. Eur J Immunol 1999; 29: 45.
  • 20
    Bachmann MF, McKall-Faienza K, Schmits R, Bouchard D, Beach J, Speiser DE, Mak TW, et al. Distinct roles for LFA-1 and CD28 during activation of naive T cells, adhesion versus co-stimulation. Immunity 1997; 7: 549.
  • 21
    Kuhlman P, Moy VT, Lollo BA, Brian AA. The accessory function of murine intercellular adhesion molecule-1 in T lymphocyte activation: contributions of adhesion and co-activation. J Immunol 1991; 146: 1773.
  • 22
    Allison AC, Harington JS, Birbeck M. An examination of the cytotoxic effects of silica on macrophages. J Exp Med 1966; 124: 141161.
  • 23
    Deaciuc IV, D'Souza NB, Nikolova-Karakashian M, de Villiers W, Sarphie TG, Hill DB, McClain CJ. The regulation of Fas (CD95/Apo-1)-mediated liver apoptosis in Kupffer cell-depleted mice. Hepatol Res 2002; 24: 192204.
  • 24
    Van Rooijen N, Sanders A. Elimination, blocking, and activation of macrophages: three of a kind? J Leuko Biol 1997; 62: 702709.
  • 25
    Boulton RA, Alison MR, Golding M, Selden C, Hodgson HJ. Augmentation of the early phase of liver regeneration after 70% partial hepatectomy in rats following selective Kupffer cell depletion. J Hepatology 1998; 29: 271280.
  • 26
    Naito M, Umeda S, Takahashi K, Shultz LD. Macrophage differentiation and granulomatous inflammation in osteopetrotic mice (op/op) defective in the production of CSF-1. Molec Reprod Dev 1997; 46: 8591.
  • 27
    Panes J, Perry MA, Anderson DC, Manning A, Leone B, Cepinskas G, Rosenbloom CL, et al. Regional differences in constitutive and induced ICAM-1 expression in vivo. Am J Physiol 1995; 269: H1955.
  • 28
    Muschen M, Warskulat U, Peters-Regehr T, Bode JG, Kubitz R, Haussinger D. Involvement of CD95 (Apo-1/Fas) ligand expressed by rat Kupffer cells in hepatic immunoregulation. Gastroenterology 1999; 116: 666667.
  • 29
    Roland CR, Mangino MJ, Duffy BF, Flye MW. Lymphocyte suppression by Kupffer cells prevents portal venous tolerance induction: a study of macrophage function after intravenous gadolinium. Transplantation 1993; 55: 11511158.
  • 30
    MacPhee PJ, Schmidt EE, Groom AC. Intermittence of blood flow in liver sinusoids, studied by high resolution in vivo microscopy. Am J Physiol 1995; 269: G692.
  • 31
    Jonsson JR, Hogan PG, Balderson GA, Ooi LL, Lynch SV, Strong RW, Powell EE. Human liver transplant perfusate: an abundant source of donor liver-associated leukocytes. HEPATOLOGY 1997; 26: 11111114.
  • 32
    Bradham CA, Plumpe J, Manns MP, Brenner DA, Trautwein C. Mechanisms of hepatic toxicity. I. TNF-induced liver injury. Am J Physiol 1998; 275: G387G392.
  • 33
    Perrilo NL, Pace KE, Seilhamer JJ, Baum LG. Apoptosis of T cells mediated by galectin-1. Nature 1995; 387: 736739.
  • 34
    Petermann H, Heymann S, Vogl S, Dargel R. Phagocytic function and metabolite production in thioacetamide-induced liver cirrhosis: a comparative study in perfused livers and cultured Kupffer cells. J Hepatol 1996; 24: 468477.
  • 35
    Ruzittu M, Carla EC, Montinari MR, Maietta G, Dini L. Modulation of cell surface expression of liver carbohydrate receptors during in vivo induction of apoptosis with lead nitrate. Cell Tissue Res 1999; 298: 105112.