Effects of Liver-Derived Dendritic Cell Progenitors on Th1- and Th2-Like Cytokine Responses In Vitro and In Vivo

Ajai Khanna, Adrian E. Morelli, Cuiping Zhong, Takuya Takayama, Lina Lu and Angus W. Thomson
J Immunol February 1, 2000, 164 (3) 1346-1354; DOI: https://doi.org/10.4049/jimmunol.164.3.1346

Abstract

There is evidence that donor-derived dendritic cells (DC), particularly those at a precursor/immature stage, may play a role in the immune privilege of liver allografts. Underlying mechanisms are poorly understood. We have examined the influence of in vitro generated mouse liver-derived DC progenitors (DCp) on proliferative, cytotoxic, and Th1/Th2 cytokine responses induced in allogeneic T cells. Liver DCp, propagated in GM-CSF from C57B10 mice (H2b), induced only minimal proliferation, and weak cytotoxic responses in allogeneic (C3H; H2k) T cells compared with mature bone marrow (BM)-derived DC. Flow-cytometric analysis of intracellular cytokine staining revealed that mature BM DC, but not liver DCp, elicited CD4+ T cell production of IFN-γ. Intracellular expression of IL-10 was very low in both BM DC- and liver DCp-stimulated CD4+ T cells. Only stimulation by liver DCp was associated with IL-10 secretion in primary MLR. Notably, these liver DCp cocultured with allogeneic T cells stained strongly for IL-10. Following local (s.c.) injection in allogeneic recipients, both BM DC and liver DCp homed to T cell areas of draining lymph nodes and spleen, where they were readily detected by immunohistochemistry up to 2 wk postinjection. Liver DCp induced clusters of IL-10- and IL-4-secreting mononuclear cells, whereas Th2 cytokine-secreting cells were not detected in mice injected with mature BM DC. By contrast, comparatively high numbers of IFN-γ+ cells were induced by BM DC. Modulation of Th2 cytokine production by donor-derived DCp may contribute to the comparative immune privilege of hepatic allografts.

The capacity of bone marrow (BM4)-derived dendritic cells (DC) to initiate or modulate immune responses appears to depend on their lineage development, microanatomic location, and their stage of phenotypic and functional maturation (1, 2, 3, 4, 5). Both myeloid and lymphoid DC (the latter derived from a precursor shared with T cells), with the apparent potential to differentially regulate immune responses, have been described (3, 4, 6, 7, 8). Although it is well established that classic mature myeloid DC in secondary lymphoid tissues are potent activators of naive T cells, immature myeloid DC resident within nonlymphoid organs (i.e., the liver or the respiratory tract) or in primary lymphoid tissue (i.e., BM) are deficient in cell surface costimulatory molecules, and can exhibit tolerogenic properties (2, 9, 10, 11).

Following organ transplantation, donor interstitial DC migrate to secondary lymphoid tissue, where they interact with specific, donor-reactive T cells (12, 13, 14, 15, 16, 17). Based on observations in rodent kidney or heart transplantation, these donor APC have been regarded historically, as instigators of rejection (14, 18). More recent evidence suggests that persistence of donor-derived DC in recipient lymphoid and nonlymphoid tissue, as occurs in nonimmunosuppressed murine liver allograft recipients, may be linked to the development of donor-specific tolerance (11, 15, 16, 17, 19). Immature myeloid DC propagated in GM-CSF from normal mouse liver, a hemopoietic organ, are deficient in cell surface costimulatory molecules. They migrate in vivo to T cell areas of secondary lymphoid tissue, where they persist for weeks in allogeneic recipients (20, 21). These liver-derived DC progenitors (DCp) can prolong allograft survival (22), a property shared with immature myeloid DC propagated from rodent BM (23, 24, 25).

DC tolerogenicity was first reported in the context of central (intrathymic) tolerance (26, 27, 28). More recently, evidence has accumulated for a role of DC in peripheral tolerance in various experimental models (2). In addition, several agents, including UVB radiation (29), IL-10 (30, 31, 32, 33), TGF-β (34, 35), or the chimeric fusion protein cytotoxic T lymphocyte Ag 4-Ig (36, 37), which blocks the B7-CD28 costimulatory pathway of T cell activation, have been shown to confer tolerogenic properties on DC. Mechanisms reported to underlie the capacity of DC to subvert T cell responses include the induction of anergy, activation-induced cell death, regulatory cells, veto function, and immune deviation toward a predominant Th2 cell response (2, 38). It is accepted that alloantigen-specific Th1 cells initiate allograft rejection, and that Th2 cells exert an inhibitory influence on the development of Th1 clones. It has thus been proposed that preferential induction of alloantigen-specific Th2 lymphocytes could suppress the development of Ag-specific Th1 cells, and as a consequence, inhibit allograft rejection. Evidence that supports or refutes this hypothesis has recently been reviewed (39, 40).

In this study, we have analyzed the T cell stimulatory capacity, tissue trafficking, and influence on Th1 and Th2 cytokine production of immature liver-DCp compared with functionally mature BM-derived DC (BM DC) propagated under similar conditions. The findings suggest that modulation of Th2 cytokine (IL-10) production by donor-derived DCp might be a mechanism underlying the capacity of liver allografts to subvert host immune responses, and contribute to tolerance induction.

Materials and Methods

Experimental animals

Ten- to 12-wk-old C57BL/10J (B10; H2Kb, I-Ab, I-E−) and C3H/HeJ (C3H; H2Kk, I-Ak, I-Ek) mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Animals were maintained in the specific pathogen-free facility of the University of Pittsburgh Medical Center (Pittsburgh, PA).

Cytokines

rmGM-CSF was a gift from the Schering-Plough Research Institute (Kenilworth, NJ).

Propagation and purification of DC populations

BM cell suspensions were prepared in RPMI 1640 (Life Technologies, Grand Island, NY), supplemented with 10% v/v heat-inactivated FCS (Life Technologies), glutamine, nonessential amino acids, sodium pyruvate, 2-ME, and penicillin/streptomycin (complete medium), using conventional procedures. The method for in vitro culture of DC was modified after that described initially by Inaba et al. (41). Briefly, 2 × 106 cells were cultured in 24-well plates in RPMI 1640 complete medium, supplemented with 1000 U/ml rmGM-CSF. Nonadherent cells released spontaneously from proliferating cell clusters were collected after 6–8 days of culture, and resuspended in complete medium. This low density cell population (DC) was washed twice before final resuspension in complete medium. Purity of the DC was verified by morphologic appearance (Giemsa staining), flow-cytometric analysis, and immunocytochemical staining of cytospin preparations. An extensive panel of mAbs was used, including those specific for leukocytic lineage (CD45), lymphoid, and myeloid cell markers, and mouse DC-restricted cell surface and intracellular Ags. Mature DC were also characterized by the absence of phagocytic activity, using both carboxylated fluorescence latex microspheres (Fluoresbrite Carboxy YG, 0.5–3.0 μm diameter; Polysciences, Warrington, PA) and opsonized SRBC (Remel, Lenexa, KS). Liver DCp were propagated in rmGM-CSF from nonparenchymal cells isolated from collagenase-digested normal liver tissue, as described (20), and characterized by mAb staining and flow-cytometric analysis, as outlined above.

Flow cytometry

Cells were incubated with the following primary mAbs (each from PharMingen, San Diego, CA, unless specified): mouse anti-H2Kb (clone AF6-88.5); rat anti-DEC 205/NLDC145 (42); hamster anti-mouse CD40 (clone HM40-3); rat anti-Gr-1 (clone RB6-8C5); rat anti-mouse CD11b (clone M1/70); hamster anti-mouse CD11c (clone HL3); rat anti-mouse CD80 (clone 1G10); and rat anti-mouse CD86 (clone GL1). Incubation with primary mAbs was followed by FITC-conjugated secondary Abs, as described (20). MHC class II Ag (I-Ab) expression was identified using biotin-conjugated mouse anti-mouse mAb (clone 25-9-3) with FITC streptavidin (Jackson ImmunoResearch, West Grove, PA) as the secondary reagent. After staining, the cells were fixed in 2% paraformaldehyde in PBS before analysis. Cytometric analysis was performed using an EPICS Elite flow cytometer (Coulter, Hialeah, FL). Appropriate fluorochrome-conjugated, isotype-matched, irrelevant mAbs were used as negative controls.

Intracellular cytokine staining

Intracellular cytokines were detected in CD4+ responder T cells after 72-h MLR. To increase the intracellular concentration of cytokines, cells were restimulated with plate-bound anti-CD3ε mAb (clone 145.2C11; PharMingen) and soluble anti-CD28 mAb (clone 37.51; PharMingen), for 5 h at 37°C, in the presence of Brefeldin A (Sigma, St. Louis, MO) (10 μg/ml; 5 h at 37°C) before staining. Thereafter, the cells were washed with 1% FCS/PBS, fixed in 4% paraformaldehyde (20 min, 4°C), and permeabilized with 0.1% saponin/1% FCS/PBS. Cells were incubated with FITC-conjugated rat anti-mouse CD4 mAb (clone GK1.5) and with PE-labeled anti-mouse IFN-γ (clone XMG1.2), anti-mouse IL-10 (clone JES5-16E3), or anti-mouse IL-4 (clone BVD4-1D11) (all mAbs from PharMingen). The cells were then washed in 1% FCS/PBS, resuspended in 1% formaldehyde, and analyzed by flow cytometry.

Allostimulatory activity (MLR)

The stimulatory activity of the DC in 72-h primary MLR was determined using purified naive allogeneic splenic T cells as responders (20).

Induction of CTLs

Generation of CTLs was quantified using 51Cr-labeled specific, third party, and syngeneic target cells, as described (24).

Cytokine quantitation

ELISA kits (Biosource International, Camarillo, CA) were used to quantify mouse IFN-γ, IL-4, and IL-10 in supernatants of cocultures of C3H T cells (responders) and either allogeneic liver DCp, allogeneic BM DC, or control splenocytes.

Immunohistochemistry and cytochemistry

Tissue samples were embedded in Tissue-Tek OCT (Miles Laboratories, Elkhart, IN), snap frozen in isopentane, chilled in liquid nitrogen, and stored at −80°C until use. Cryostat sections (8 μm) were mounted on slides treated with Vectabound (Vector, Burlingame, CA), air dried, and fixed in cold acetone (4°C) for 10 min. Cells harvested from cocultures of C3H T lymphocytes and either allogeneic liver DCp or allogeneic BM DC were spun onto glass slides (5 min at 230 × g) using a Shandon cytocentrifuge, then air dried and fixed in cold acetone for 5 min. For cytokine staining, sections or cytospins were incubated successively with: 1) normal goat serum (1/10; 20 min at RT) to inhibit nonspecific binding by blocking FcR; 2) avidin-blocking solution (Vector) (15 min at RT); 3) optimal concentrations (1/50–1/100) of primary rat mAbs (PharMingen) specific for mouse IFN-γ (clone R4-6A2), mouse IL-10 (clone JES5-16E3), or mouse IL-4 (clone BVD4-1D11) (2 h at RT); 4) biotinylated polyclonal anti-rat Ig (1/100; PharMingen) (30 min at RT); and 5) avidin-biotin complex-alkaline phosphatase (ABC-AP; Vector) (30 min at RT).

For detection of donor cells in recipient tissues, sections were incubated in a 1/100 dilution of biotinylated mouse anti-I-Ab β-chain mAb (clone 25-9-17; PharMingen), followed by ABC-peroxidase (ABC-Px; Vector). AP activity was detected by incubation with the substrate Vector Blue (Vector). To increase the sensitivity of the assay for detection of IL-10 and IL-4 in some samples, the AP activity was developed with the substrate BCIP/NBT (Vector). Peroxidase (Px) activity was developed with 3,3′-diaminobenzidine tetrahydrochloride (Sigma). Endogenous AP activity was inhibited by addition of levamisole (Vector) in the substrate solution. Endogenous Px activity was blocked by successive passages in 70% ethanol, 1% H2O2 in methanol, and 70% ethanol. Tissue sections immunostained with ABC-AP (end product blue) were counterstained with Fast Red (Vector), and mounted first in Crystal/mount (Biomeda, Foster City, CA), and then in Permount (Fisher Scientific Company, Pittsburgh, PA). Sections stained with ABC-Px (end product brown) were counterstained with hematoxylin, dehydrated, and mounted in Crystal/mount. Rat or mouse irrelevant Igs of the same isotype as the primary mAb were used as controls.

Statistical analysis

Results are expressed as means ± 1 SD. Comparisons between different means were performed by ANOVA, followed by the Student Newman Keuls test. Comparison between two means was performed by the Student t test. A p value <0.05 was considered significant.

Results

Phenotypic characteristics of BM DC and liver DCp

After 6–8 days of culture, most cells released from proliferating aggregates of GM-CSF-stimulated liver- or BM-derived cells exhibited typical DC morphology, with prominent cytoplasmic processes, and absence of prominent cytoplasmic granules. The surface phenotype of nonadherent or loosely adherent cells from liver or BM-derived cultures was analyzed by flow cytometry after 6–8 days of culture. Staining for cells of T (CD3), B (CD45/B220), NK (NK1.1), and granulocytic (Gr-1) lineages was absent. As detailed previously (20), liver DCp expressed CD45 (leukocyte common Ag), CD24 (heat-stable protein), CD54 (ICAM-1), CD11b (MAC-1), and CD44 (nonpolymorphic determinant of Pgp.1 glycoprotein). These liver-derived cells also showed weak positivity for the mouse DC-restricted markers CD11c, DEC205, and 33D1, and were positive for the macrophage marker F4/80 (data not shown). Similar results were obtained for BM DC, except that they expressed CD11c and DEC205 strongly. Whereas liver DCp displayed low amounts of MHC and costimulatory molecules (CD40, CD80, CD86), BM DC expressed moderate to high levels of these markers. As reported previously (20), the low expression of MHC Ag and the absence or low levels of costimulatory molecules on liver DCp, cultured under similar conditions to BM DC, were indicative of cells at an immature stage of differentiation.

Liver DCp induce only minimal T cell proliferation

To test their allostimulatory activity, in vitro generated B10 BM DC or liver DCp were irradiated, and set up in 72-h primary MLR cultures with naive C3H T cells. In comparison with BM DC, which were potent inducers of DNA synthesis, and consistent with their surface phenotype, the liver DCp induced only minimal levels of T cell proliferation (Fig. 1⇓). The poor stimulatory capacity of liver DCp remained unchanged after longer incubation times with allogeneic T cells (4- or 5-day MLR; data not shown).

FIGURE 1.
FIGURE 1.

Allostimulatory activity of γ-irradiated, GM-CSF-stimulated immature B10 (H2b) liver DCp (▾), or mature B10 BM DC (▿), assessed using naive C3H (H2k) splenic T cells as responders. The stimulator cells were propagated as described in Materials and Methods, harvested after 6- to 8-day culture, and purified by metrizamide centrifugation. They were set up at various concentrations with 2 × 105 responder T cells, and the cultures maintained for 72 h. [3H]TdR was added 18 h before harvesting. The MLR-stimulatory activity of freshly isolated syngenic (C3H ○) or allogenic (B10 •) bulk spleen cells is also shown. The results are expressed as mean cpm ± 1 SD, and are representative of at least three separate experiments.

Liver DCp elicit low levels of CTL activity

We next determined the ability of the cells to induce generation of CTL in 6-day primary MLR. As shown in Fig. 2⇓, effector T cells induced by allogeneic BM DC exhibited a progressive increase in specific cytotoxic activity at increasing E:T ratios. By contrast, identical numbers of liver DCp induced only minimal levels of CTL activity, which were lower than those induced by bulk allogeneic spleen cells.

FIGURE 2.
FIGURE 2.

CTL activity elicited by liver DCp or BM DC in allogeneic MLR. GM-CSF-stimulated liver DCp or BM DC were propagated from B10 mice, and used as stimulators of C3H T cells, as described in Materials and Methods. B10 or C3H spleen cells served as positive (allogeneic) and negative (syngeneic) control stimulators, respectively. After 6 days, the cultures were harvested, T cells purified, and CTL activity determined against appropriate (H2b) target cells. CTL activity generated by C3H splenocytes as stimulators (negative control) was below 2% in all experiments (data not shown). Results are representative of three separate experiments performed using a 4-h 51Cr release assay.

Differential expression of intracellular cytokines in allogeneic T cells stimulated with liver DCp or BM DC

The intracytoplasmic expression of IFN-γ, IL-10, and IL-4 was analyzed by flow cytometry in responder CD4+ T cells (C3H) after in vitro stimulation in 3-day MLR with either allogeneic mature B10 BM DC, or immature liver DCp. Before staining for intracytoplasmic cytokines, T cells were restimulated by immobilized mAb to CD3ε and soluble mAb to CD28 in the presence of Brefeldin A, which causes newly synthesized proteins to accumulate within the endoplasmic reticulum. Without Brefeldin A, the expression of cytokines could not be detected by flow cytometry. When Brefeldin A was added to the cultures, without restimulation with anti-CD3ε and anti-CD28 mAbs, a similar pattern of cytokine expression was detected, but at a much lower level. The cells were harvested, fixed, permeabilized, and processed for double staining using primary PE- conjugated cytokine-specific mAb, followed by FITC-coupled anti-CD4 mAb. Based on their high SSC compared with T cells, and their lack of expression of CD4 (41), DC were gated out of the mixed cell population (composed of T cells and DC at a ratio 20:1) harvested from the MLR. The intracellular expression of cytokines in CD4+ T cells stimulated either with liver DCp or with BM DC is illustrated in Fig. 3⇓. In a typical experiment, a substantial fraction of CD4+ T cells (>15%) was induced to produce IFN-γ, but few cells expressed IL-10 or IL-4, when allogeneic BM DC are used as stimulators (Fig. 3⇓). By contrast, very few CD4+ T cells (≤2%) produced detectable levels of either IFN-γ, IL-10, or IL-4 after 3 days allostimulation with liver DCp (Fig. 3⇓). C3H T cells maintained for 72 h, with or without IL-2 (2 U/ml), then stimulated with anti-CD3ε mAb and anti- CD28 mAb, treated with Brefeldin A, and immunostained as described above, were included as controls. Less than 0.2% of these control CD4+ T cells were positive for the cytokines analyzed (data not shown).

FIGURE 3.
FIGURE 3.

Intracytoplasmic detection of IL-10, IFN-γ, and IL-4 in C3H CD4+ T cells after in vitro stimulation with either liver DCp (upper row) or mature B10 BM DC (lower row), as described in Materials and Methods. After 3-day MLR, the cells were harvested and restimulated with immobilized anti-CD3ε mAb and soluble anti-CD28 mAb in the presence of Brefeldin A for 5 h at 37°C. Thereafter, the cells were fixed, permeabilized, and processed for double immunocytochemical staining, using primary rat mAbs specific for murine cytokines, revealed by PE-conjugated anti-rat Igs, followed by FITC-conjugated anti-CD4. Stimulatory myeloid DC were gated out according to their high SSC and their lack of expression of CD4. The data are representative of three separate experiments.

Comparatively low levels of IFN-γ, but comparatively high levels of IL-10, are present in cocultures of liver-derived DCp and allogeneic T cells

To ascertain the influence of mature BM DC vs liver DCp on allogeneic Th0 differentiation into Th1 or Th2 clones, we quantified the production of IFN-γ, IL-10, and IL-4 by ELISA in supernatants obtained from 72-h cocultures of allogeneic naive T cells with either BM DC or liver DCp (DC:T cell ratio = 1:20). IFN-γ production was detected in BM DC-stimulated cultures, but was substantially lower (∼10% of the former) when liver-derived DCp were employed as stimulators (Fig. 4⇓A). To confirm whether DC or T cells were the source(s) of cytokine production, cytospin preparations of cells collected from MLRs were stained for IFN-γ by immunocytochemistry. Both BM DC and some T cells, grouped in small clusters, were positive for IFN-γ (data not shown). By contrast, IL-10 production was detected only in cocultures of liver DCp and allogeneic T cells. In these latter cultures, the concentration of IL-10 at 72 h was ∼25-fold that in cocultures of BM DC and allogeneic T cells (Fig. 4⇓B). Immunostained cytospins revealed that liver DCp were significant IL-10 producers (Fig. 5⇓A). In addition, very few clusters of T cells, most of them in close apposition to DC, were also positive for intracytoplasmic IL-10 (Fig. 5⇓A). There was much less evidence of IL-10 production in corresponding cytospin preparations from BM DC-T cell cultures (Fig. 5⇓B). IL-4 was not detected by ELISA, or by immunocytochemistry in cytospin preparations.

FIGURE 4.
FIGURE 4.

Secretion of IFN-γ (A) and IL-10 (B) in MLR culture supernatants using B10 BM DC or liver DCp as stimulators of naive C3H T cells. Bulk allogeneic B10 spleen cells or syngeneic C3H spleen cells were used as reference controls. IFN-γ and IL-10 were quantified by ELISA at 72 h. Results are representative of two separate experiments.

FIGURE 5.
FIGURE 5.

Immunocytochemical detection of intracytoplasmic IL-10 (positivity in blue) in cytospin preparations from 3-day MLR elicited using allogeneic liver DCp (A), or BM DC (B) as stimulators. Responder T cells or T cell blasts are indicated by arrows, and stimulatory DC by an asterisk (×400; ABC-AP, counterstained with Fast Red).

Trafficking of BM DC and liver DCp to T cell areas in allogeneic recipients

To assess their ability to home to recipient lymphoid tissues, and to survive in an allogeneic environment in vivo, 5 × 105 liver DCp or BM DC (B10) were injected s.c. in one hind footpad of normal allogeneic mice (C3H). At days 1, 2, 7, and 14, animals were sacrificed, and donor cells detected in cryostat sections of recipient lymphoid tissues (draining lymph nodes, spleen, and thymus) by staining for donor MHC class II (I-Ab). Donor cells with dendritic morphology were present in the subcapsular and paracapsular sinuses, and in the T-dependent areas (the interfollicular area of the cortex, and paracortex) of popliteal lymph nodes, 2 days after their administration, but could not be detected thereafter (Fig. 6⇓, A and D). No donor MHC class II+ cells were observed in the inguinal lymph nodes. Donor BM DC and liver DCp showed a similar pattern of tissue homing in the spleen and thymus (Fig. 6⇓, B, C, E, and F). During the first 2 days after injection, both types of donor DC were located principally in the marginal zones of the spleen (Fig. 7⇓A). At later time points (days 7 and 14), DC mobilized into the periarteriolar lymphatic sheaths (PALS, Fig. 6⇓, B and E). In general, both donor BM DC and liver DCp were detected as single cells; however, less frequently, liver DCp formed clusters of 5–10 cells. These clusters were located mainly in marginal zones (Fig. 7⇓A). The number of donor BM DC present in spleen peaked at day 7, and decreased significantly thereafter (p < 0.05). By contrast, the number of liver-derived DCp remained constant throughout the 14-day follow-up period (p = 0.65). In the thymus, a few donor BM DC or liver DCp were detected at the corticomedullary junction on days 1, 2, 7, and 14 (Fig. 6⇓, C and F).

FIGURE 6.
FIGURE 6.

Tissue trafficking of allogeneic donor BM DC (top; A, B, and C) and liver DCp (bottom; D, E, and F) to recipient popliteal lymph nodes at day 2 after DC administration (left column, A and D), spleen at day 7 (middle column, B and E), and thymus at day 7 (right column, C and F). Donor DC were detected in cryostat sections by immunohistochemistry, using a mAb directed against the donor MHC class II (anti-I-Ab β-chain). An ABC-Px technique was employed to detect donor DC in lymph node and thymus (positive cells in brown), and an ABC-AP procedure to stain donor cells in spleen (positive cells in blue). In lymph nodes, positive cells with DC morphology (arrows) were located in the interfollicular areas, or in the subcapsular and paracapsular sinuses. In spleen, donor cells (arrows and blue cells in the insets) were located in the PALS, close to arterioles (arrowheads). In the thymus, rare donor cells (arrows and brown cells in the insets) were found at the corticomedullary junction (arrowheads) (×100; counterstained with hematoxylin or Fast Red).

FIGURE 7.
FIGURE 7.

A, Tissue trafficking of donor liver DCp in allogeneic spleen. Donor DC were detected in cryostat sections by immunohistochemistry, 1, 2, 7, and 14 days after their injection, using a mAb directed against the donor MHC class II (I-Ab β-chain). Donor DC (blue cells, indicated by an arrow) were detected by the ABC-AP technique, as single cells, or more rarely, as small clusters (inset) in the T cell-dependent areas (PALS), or in the marginal zone. The arteriole of the PALS is indicated by an asterisk. (×100; counterstained with Fast Red). B, IL-10 expression in spleens of recipients of donor liver DCp injected 7 days previously. The black area (indicated by an arrow) represents a cluster of cells with lymphocyte morphology positive for IL-10. IL-10-secreting areas were detected only when liver DCp were injected, and were located in T cell-dependent areas of the spleen (×100, ABC-AP, counterstained with Fast Red).

BM DC and liver DCp differentially induce IFN-γ, IL-10, and IL-4 in allogeneic recipients

Analysis of in situ cytokine production by recipient lymphocytic cells in T cell areas of spleens of animals injected either with allogeneic liver DCp or BM DC was performed at different times (days 1, 2, 7, and 14) after their administration. The local synthesis of cytokines was quantified as the number of lymphocytic clusters (more than three positive cells with round cell morphology in close apposition) per cm2 of recipient spleen. As can be seen in Figs. 8⇓ and 9, the number of clusters producing IFN-γ at day 7 was significantly higher (p = 0.04) in those animals injected with BM DC compared with mice injected with liver DCp. Conversely, clusters expressing IL-10 or IL-4 were only detected in spleens of mice given liver DCp (Figs. 7⇑B and 8). Immunohistochemical staining of serial sections with a mAb specific for the T cell marker CD3ε demonstrated that the areas positive for specific cytokines corresponded to T cell areas of the spleen.

FIGURE 8.
FIGURE 8.

Kinetics of in situ IFN-γ, IL-10, and IL-4 production quantified by the number of cytokine-positive lymphocytic clusters (more than three cells) detected per cm2 in cryostat sections of recipient spleens at various times after s.c. administration of donor BM DC (♦), or liver DCp (▴, dotted line). Results are means ± 1 SD obtained from groups of three mice at each time point.

FIGURE 9.
FIGURE 9.

IFN-γ expression in spleens of recipients of donor BM DC (top; A, B, and C), or liver DCp (bottom; D, E, and F). IFN-γ-secreting areas in spleen were detected by immunohistochemistry on cryostat sections, at day 2 (A and D), day 7 (B and E), and day 14 (C and F) after DC administration. The blue areas (some indicated with arrows) represent foci of cells with lymphocytic morphology, positive for IFN-γ. Inset in B, detail of a cluster positive for IFN-γ. Most of the IFN-γ-producing areas were located in the PALS, indicated in some instances by a white arrow pointing to the arteriole (×100; ABC-AP, counterstained with Fast Red).

Discussion

After vascularized organ transplantation, donor passenger leukocytes (mainly interstitial DC) are mobilized out of the graft via peripheral blood to the recipient lymphoid and nonlymphoid tissues (13, 15, 16, 17, 19). For transplanted tissues such as skin, donor DC migrate from the graft via the dermal lymphatic vessels into draining lymph nodes (12). Improvement in graft survival following depletion of leukocytes from thyroid, pancreatic islet, skin, or kidney allografts supports the concept that trafficking and maturation of donor DC in recipient lymphoid organs lead to the activation of naive, alloreactive Th0 lymphocytes, and thus provides the primary stimulus for acute allograft rejection (18, 43, 44). By contrast, in the absence of immune suppression, liver allografts (that possess comparatively large numbers of passenger leukocytes) can induce donor-specific tolerance in fully MHC-mismatched mouse and certain rat strain combinations, and in a high proportion of outbred pigs (19, 45, 46, 47). Moreover, in humans, the liver is considered the least immunogenic of transplanted whole organs. In a tolerant rat strain combination, depletion of interstitial leukocytes from livers by pretransplant donor radiation prevents the tolerogenic effect, and results in acute rejection (48). Moreover, the tolerogenic ability of leukocyte-purged liver grafts can be restored by reconstituting the interstitial hemopoietic cell population by syngeneic liver or spleen leukocyte infusion, or after “parking” the leukocyte-depleted liver in a syngeneic recipient before allotransplantation (49).

Therefore, passenger leukocytes (most likely DC) may have a dualistic role with potential to elicit T cell activation and graft rejection, or induce T cell tolerance and graft acceptance (11, 14). The sustained release from the transplanted liver of immature APC, deficient in costimulatory molecules, with poor comparative ability to stimulate Th1 and the development of alloantigen-specific CTLs, delayed-type hypersensitivity, and IgG2a production (that promotes Ab-dependent cellular cytotoxicity), may contribute to tolerance induction. Alternatively, liver-derived DCp might induce the proliferation of Th2 clones with capacity to inhibit Th1 responses, or facilitate the development of Th cells with regulatory/suppressor functions, such as T regulatory-1 (Tr1) cells (50, 51). Recent findings suggest that in mice, lymphoid-related DC (CD11c+, CD11b−, CD8α+) elicit only Th1 responses, whereas myeloid DC (CD11c+, CD11b+, CD8α−) induce either Th1 or Th2 responses in vivo (52, 53).

In the present study, we have confirmed that, in comparison with mature BM DC, liver DCp display an immature phenotype with absence of CD40 and CD86 surface expression, low levels of MHC class I and II, and as a consequence, low stimulatory capacity for naive allogeneic T cells. Unlike mature BM DC, these liver DCp did not induce detectable levels of intracytoplasmic IFN-γ in allogeneic CD4+ cells in 72-h MLR, and elicited very low levels of CTLs in vitro. These observations suggest that liver DCp have a more restricted capacity than BM DC to stimulate the differentiation of Th0 cells to Th1 cells. Accumulating evidence indicates that the type of cytokine secreted during the early stages of a primary immune response determines the sort of T cell clone to be generated. Thus, in an IL-4-rich microenvironment, DC induce T cells to differentiate into IL-4/IL-5/IL-10/IL-13-producing Th2 cells (50). DC are able to produce and secrete IL-12, a cytokine that turns naive, CD4+ Th0 cells into IFN-γ-secreting Th1 lymphocytes (54, 55). Although we were able to detect secretion of IL-12 p70 by BM DC using ELISA (49.9 ± 2.4 pg/ml/24 h/106 DC), IL-12 p70 secretion by liver DCp could not be detected. This finding could explain the poor capacity of liver DCp to generate Th1 cells in vitro. Interestingly, we found that liver DCp produced IL-10, and induced a low incidence of IL-10+ allogeneic responder CD4+ cells, a phenomenon that may be related to the induction of either Th2 or Tr1 cell clones (50, 51). The very low incidence of IL-10+ T cells induced, and the low proliferative capacity of Th2 or Tr1 cell clones that has been reported (10, 51) could explain the poor incorporation of [3H]TdR observed in proliferation assays in this study. Stumbles et al. (10) have reported that in the rat respiratory tract, resident DC also show an immature phenotype (MHC class Ilow, class IIlow, CD80−, CD86−) and express high levels of IL-10 mRNA. Although freshly isolated respiratory tract immature DC are poor stimulators in primary MLR, they elicit a predominant Th2 response in vivo after being pulsed ex vivo with OVA (10). By contrast, maturation of respiratory tract DC with GM-CSF up-regulates the expression of MHC molecules, B7, and the capacity to stimulate both Th1 and Th2 responses, a phenomenon that seems to be associated with increased production of IL-12 p35 chain mRNA, and down-regulation of IL-10 mRNA synthesis (10).

Previous reports have demonstrated that IL-10-treated DC, or viral IL-10 gene-transduced DC, exhibit tolerogenic properties (30, 31, 32, 33). In the present study, we have confirmed that IL-10 is produced by putative tolerogenic liver-derived DCp, an observation that agrees with the high levels of IL-10 mRNA reported in immature DC in the rat respiratory tract (10). There is also recent evidence that in humans, the subpopulation of CD14+ CD1a− DC generated in vitro in the presence of GM-CSF and TNF-α produces IL-10, a fact suggesting that other subpopulations of myeloid DC that reside in peripheral tissues might secrete this cytokine (56). In this regard, it was demonstrated recently that DC isolated from Peyer’s patches secreted IL-10, and were able to induce differentiation of Th2 cells (57). IL-10, produced by immature DC, Th2, and Tr1 cells, down-modulates the expression of CD80 and CD86 on DC, accelerates their apoptotic death, and skews the Th1/Th2 balance to Th2 by inhibiting IL-12 synthesis by DC (30, 31, 32, 58, 59, 60, 61). Thus, IL-10 may play a key role in exhibition of the tolerogenic properties of liver DCp in vivo.

A specialized ability of DC is the capacity to migrate in vivo to T cell areas of peripheral lymphoid tissues (20, 21, 62). As reported herein, both donor BM DC and liver DCp were detected in the subcapsular and paracortical sinuses of the draining lymph node after s.c. administration, a fact that might represent donor DC in transit to the interfollicular T cell area where they were also found. Using a similar model of DC trafficking, others have observed that the number of donor DC in the popliteal lymph node decreases rapidly 48 h after their administration (63). In the spleen, donor DC were present 24 h after administration. Whereas the number of donor liver DCp detected in spleen remained constant throughout the 14-day follow-up period, the density of BM DC increased steadily until day 7, and decreased thereafter. A greater capacity of donor liver DCp, in comparison with BM DC, to shut down generation of a Th1 response, or alternatively, their inability to trigger a cellular immune response, might contribute to the extended survival of donor liver DC that has been observed after liver transplantation (16, 19).

In the present study, injection of liver DCp mimicked the trafficking pattern to the spleen described by Demetris et al. after orthotopic liver transplantation in rats (16). Donor DC were concentrated initially in the marginal zones of the spleen, and later mobilized to the T cell-dependent PALS. The marginal zone has been considered a site populated by immature myeloid DC, B cells, and specialized macrophages (64, 65). Specialized gates in the marginal zone appear to be the main routes to the white pulp employed by circulating DC in physiological conditions (66, 67). However, the present and previous trafficking studies (16) indicate that donor-derived DC may also enter the white pulp through the marginal zone after liver transplantation. Whether immature donor DC remain as precursors in the marginal zone, and replicate to maintain the population of immature DC long-term (late chimerism) remains unknown. However, the presence of donor MHC class II+ proliferating cells (detected by others using 5-bromo-2′-deoxyuridine incorporation, or by the presence of mitotic figures), and our observation of clusters of donor cells with DC morphology in the marginal zone, are evidence for the replicative potential of donor liver DCp that colonize the recipient spleen (16, 21). As reported previously after liver transplantation, or after administration of liver DCp, very few donor DC were detected in the recipient thymus throughout the 14-day follow-up (16, 21). Interestingly, those that were found were located at the corticomedullary junction. Whether these donor DC may play a role in acquired thymic tolerance to alloantigens (68) is still unknown.

Administration of mature BM DC was associated with a higher number of IFN-γ-positive cells in spleen than observed after injection of immature liver DCp. A certain degree of DCp maturation and limited IL-12p70 secretion may have occurred in vivo in response to proinflammatory cytokines released at the injection site, and following interaction of DCp with extracellular matrix proteins (20) during cell trafficking. Clusters of cells with lymphocyte morphology, located in T cell areas, and positive for IL-10 or IL-4, were only detected after injection of liver DCp. These results suggest that in vivo, allogeneic immature liver DCp have more restricted capacity than BM DC to induce the proliferation of IFN-γ-producing recipient T cells. On the other hand, unlike BM DC, they appear to promote the proliferation of recipient allogeneic T cells with the ability to produce IL-10 and IL-4. Such cells may either shift the Th balance toward a Th2 response, or induce the proliferation of regulatory T cell clones with the capability to produce IL-10 and/or IL-4, but comparatively little IFN-γ (50, 51). The very low number of IL-10- or IL-4-positive clusters detected in spleens of liver DCp recipients, in proportion to the number of IFN-γ-positive areas observed when BM DC were injected, might be ascribed to the lower replicative potential of Th2 and T regulatory cell clones compared with Th1 cells (10, 51).

Although generation of a Th1 response, determined by cytokine mRNA expression after liver transplantation (69, 70, 71), may appear to contradict the present observation that liver DCp preferentially induce Th2 cytokines in vivo, the two phenomena may represent different phases of the tolerance induction process. During the early phase (days 1 to 2), migration of mature DC from the liver may induce a Th1 response in recipient lymphoid tissue (peak of IL-2 and IFN-γ mRNA synthesis in spleen and celiac lymph nodes (69, 70, 72)). However, during the following 14 days, the number of infiltrating recipient T cells and their cytotoxic activity rapidly decrease, due to apoptosis (72). Meanwhile, migration of donor liver DCp to secondary lymphoid organs may promote, in a second phase, generation of Th2 and/or regulatory T cell clones with the ability to down-regulate the Th1 allo-response, and maintain the state of alloantigen-specific tolerance.

There is evidence to support or refute the hypothesis that organ transplant tolerance involves a dominant Th2 response (39, 40, 73, 74). Moreover, it has been observed that administration of IL-10 to heart allograft recipients can inhibit or exacerbate rejection, depending on cytokine dosage and timing (75, 76). The present observations suggest that immature and mature myeloid DC may differentially modulate Th1 and Th2 cytokine production in vitro and in vivo. An implication of the finding that immature donor (liver) DCp secrete IL-10 and/or modulate host responses toward Th2 cytokine predominance is that these cells may contribute toward the comparative immunologic privilege of hepatic allografts.

Acknowledgments

We thank Dr. Adriana Larregina for advice on statistical analyses, Jennifer Little and Allison Logar for skillful assistance with cell culture and flow cytometry, and the Schering-Plough Research Institute (Kenilworth, NJ) for gifts of cytokines.

Footnotes

  • ↵1 This study was supported by National Institutes of Health Grants DK 49745 and AI 41011 (to A.W.T.).

  • ↵2 A.K. and A.E.M. contributed equally to this work and should be considered as co-first authors.

  • ↵3 Address correspondence and reprint requests to Dr. Lina Lu or Dr. Angus W. Thomson, Department of Surgery, University of Pittsburgh Medical Center, W1540, Biomedical Science Tower, 200 Lothrop Street, Pittsburgh, PA 15213. E-mail addresses: thomsonaw@msx.upmc.edu or lul{at}msx.upmc.edu

  • ↵4 Abbreviations used in this paper: BM, bone marrow; ABC, avidin-biotin complex; AP, alkaline phosphatase; DC, dendritic cell; DCp, dendritic cell progenitor; m, mouse; PALS, periarteriolar lymphatic sheaths; Px, peroxidase; RT, room temperature; Tr1, T regulatory-1.

  • Received March 26, 1999.
  • Accepted November 18, 1999.

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The Journal of Immunology: 164 (3)
The Journal of Immunology
Vol. 164, Issue 3
1 Feb 2000
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