Arch Dermatol Res (2000) 292 : 437–445
© Springer-Verlag 2000
O R I G I N A L PA P E R
Kerstin Steinbrink · Lydia Paragnik · Helmut Jonuleit · Thomas Tüting · Jürgen Knop · Alexander H. Enk
Induction of dendritic cell maturation and modulation of dendritic cell-induced immune responses by prostaglandins
Received: 3 February 2000 / Accepted: 8 June 2000
Abstract Dendritic cells (DC) are the most potent antigen-presenting cells of the immune system. In this study we investigated the effects of various prostaglandins (PG) on the stimulatory capacity of DC. DC were generated from peripheral progenitor cells in the presence of IL-4 and GM-CSF and stimulated with IL-1, IL-6 and TNF-α on day 7. Simultaneously, PG (PGD2, PGE1, PGE2, PGF2alpha, PGI2) were added at various concentrations (10–5 to 10–9 M) on day 7. In all experiments, PGE2 had the most potent influence on the maturation of the DC, followed by other PG in the order PGE1>PGD2>PGF2alpha>PGI2. In addition, the expression of the surface molecules CD40, CD54, CD58, CD80, CD83, CD86 and the MHC class II molecules was upregulated after stimulation with PG. Analysis of DC supernatants after treatment with PG demonstrated significantly higher amounts of the proinflammatory cytokines IL-1β, IL-6, TNF-α, and IL-12. Addition of PG to DC induced a markedly enhanced proliferation of both naive and activated CD4+ and CD8+ T cells in alloantigen-induced MLR assays. Assessment of coculture supernatants after restimulation revealed significantly higher amounts of the Th1cytokines IL-2 and IFN-γ and only minimal amounts of IL-4 compared to control cells. No production of IL10 was observed. The effects of PG on the maturation of DC and enhanced T-cell proliferation could be mimicked by db-cAMP and forskolin, indicating that they were due to elevated cAMP levels. Collectively, our data show that members of the PG family promote the
This work was supported by the DFG and the BMBF. Kerstin Steinbrink was supported by a fellowship of the Deutsche Forschungsgemeinschaft. K. Steinbrink () · L. Paragnik · H. Jonuleit · T. Tüting · J. Knop · A. H. Enk Department of Dermatology, University of Mainz, Langenbeckstr. 1, 55131 Mainz, Germany e-mail:
[email protected], Tel.: +49-6131-172297, Fax: +49-6131-176614
differentiation of DC and enhance their capacity to induce a Th1 immune response. Key words Prostaglandins · Dendritic cell · T cell immunity · Th1 response
Introduction Dendritic cells (DC) are a family of bone marrow-derived, highly specialized antigen-presenting cells (APC) of the immune system (Steinman 1991). The strategic positioning of DC in nonlymphoid tissue and their ability to circulate via blood and lymph to lymphoid organs after antigen stimulation demonstrate their important role in the induction of immune responses against invading pathogens (Austyn et al. 1988; Hoefsmit et al. 1982). Locally produced inflammatory cytokines and encounter with an antigen promote the migration of DC to regional lymph nodes. During this migration from peripheral tissues to lymphoid organs, DC undergo a process of maturation involving dramatic changes in phenotype and function from an “antigen-processing” to an “antigen-presenting” cell type, characterized by the expression of costimulatory molecules, cytokine production and a typical morphology (Schuler and Steinman 1985; Steinman and Young 1992; Young et al. 1992). In contrast to other types of APC, fully mature DC are potent activators of naive T cells and are regarded as important initiators of primary specific immune responses (Steinman 1991). Prostaglandins (PG) are members of the arachidonic acid family (Goodwin and Cuppens 1983). They are inflammatory mediators, produced by stromal cells and infiltrating mononuclear cells and possess many functions in the regulation of the immune system, the regulation of the cardiovascular system and the modulation of the activity of smooth muscles and nerve cells (Bergström et al. 1968; Davies et al. 1984). The most intensively studied PG include those of the E series (e.g. PGE2). Cell activation by mitogens, endotoxins, IFNs, or antigen-antibody complexes increases PGE2 production (Goodwin and
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Cuppens 1983; Goodwin et al. 1977). Of central importance in most host defence and autoimmune responses is that PGE2 inhibits T-cell proliferation and migration and decreases the production of Th1 cytokines, such as IL-2 and IFN-γ, but not of Th2 cytokines by T cells (Betz and Fox 1991; Goodwin et al. 1977; Oppenheimer-Marks et al. 1994; Snijedewint et al. 1993). In vivo, various Th2 cell-related diseases are associated with elevated PGE2 production (Fogh et al. 1989; Misra et al. 1995). Furthermore, the stimulation of whole blood culture or monocytes with PG downregulates the production of IL-12, which raises the possibility that PGE2 affects the function of APC and thereby modifies the resulting T-cell response (Van der Pouw et al. 1995). Recently, it has been shown that PGE2 induces the maturation of human DC resulting in an increased stimulatory capacity of T cells (Jonuleit et al. 1997; Rieser et al. 1997). However, other groups have demonstrated a reduced production of IL-12 by PG-stimulated DC and the induction of a Th2 response (Kalinski et al. 1997; Kalinski et al. 1998). The aim of our study was to investigate the effect of various members of the PG family (PGD2, PGE1, PGE2, PGF2alpha, PGI2) on the characteristics and function of human monocyte-derived DC and the resulting T-cell response. We demonstrated that PGE2 had the most potent influence on the maturation of the DC in all experiments performed. Analysis of the DC showed an increased expression of the costimulatory molecules CD40, CD54, CD80, CD86 and the MHC class II molecules. Furthermore, the percentage of fully mature DC as characterized by the DC-specific antigen CD83 is enhanced after the addition of PG to the culture (Zhou and Tedder 1996). Investigations of cytokine production by stimulated DC revealed higher amounts of proinflammatory cytokines (IL1β, IL-6, TNF-α) and also of IL-12. After restimulation, analysis of the coculture supernatants showed an increased level of the Th1 cytokines IL-2 and IFN-γ but no IL-4 or IL-10 compared to control cells, indicating the promotion of a Th1 T cell response.
Materials and methods Preparation of DC Blood-derived DC were prepared according to modified protocols (Peters et al. 1993; Romani et al. 1994; Ruppert et al. 1993). Briefly, whole blood was heparinized and separated by a Ficoll gradient. PBMC fractions were then depleted of T and B cells using immunomagnetic beads coated with anti-CD2 and anti-CD19 monoclonal antibodies (mAb) (Dynal, Oslo, Norway). Remaining cells were cultured in X-VIVO 15 (BioWhittacker, Md.) in six-well plates (Costar, Cambridge, Mass.) for 7 days. Cultures were supplemented with 1000 U/ml hIL-4 (PBH, Hannover, Germany) and 800 U/ml hGM-CSF (Leukomax, Sandoz, Basel, Switzerland), as well as 1% autologous plasma. Cells were fed with fresh medium every 2 days. On day 7, nonadherent cells were rinsed off the plates and resuspended in fresh complete medium with GM-CSF and IL-4 and additionally stimulated with IL-1β (10 ng/ml), TNF-α (10 ng/ml) (PBH), IL-6 (1000 U/ml) (R & D Systems, Wiesbaden, Germany) to induce and stabilize the maturation of DC (Jonuleit et
al. 1997). Simultaneously various members of the arachidonic acid family (PGE2, PGE1, PGD2, PGF2alpha, PGI2) (Sigma, Munich, Germany) were added to the cultures at various concentrations (10–5 to 10–9 M). In subsequent experiments appropriate concentrations of every PG were used as indicated. DC were harvested for FACS analysis or functional tests 3–5 days after resuspension and stimulation. To inhibit the effect of PG, polyclonal PG-antisera (anti-PGD2, anti-PGE2, anti-PGE1, anti-PGF2alpha; CellSystems, Remagen, Germany) were simultaneously added at a concentration of 20 µg/ml to the DC. Furthermore, in coculture experiments antisera were added to exclude a direct effect on T-cell function. For experiments with nonphysiological modulators of cAMP dibutyryl-cAMP (db-cAMP, 10–6 M; Sigma) and forskolin (50 µM; Alexis Biochemical, Grünberg, Germany) were added simultaneously with the stimulating cytokines to the DC on day 7. T cell purification, allogeneic proliferation T cells were prepared from human blood using Ficoll gradients and subsequent purification by antibody-coated immunomagnetic beads (MACS systems, Miltenyl, Bergisch Gladbach, Germany) according to standard protocols (purity >95% CD4+/CD8+ T cells, >90% CD45RA+ T cells). In some experiments cord blood was used as the source of naive T cells. Purity was tested by FACS analysis. DC were prepared as described above and cocultured with 2 × 105 T cells per well at ratios in the range 1 : 10–1 : 20 for restimulation experiments and a 1 : 2 titration for allogeneic MLR in 96well-plates (Costar, Cambridge, Mass.). After 3 days the cells were pulsed with 1 µCi [3H]TdR ([methyl-3H]thymidine) per well for 8 h, harvested and counted. Tests were carried out in triplicate, and the results are expressed as mean±SD counts per minute. FACS analysis After culture, cells were subjected to analysis by FACS (Becton Dickinson) using the following mAbs: anti-CD1a, anti-CD2, antiCD14, anti-CD16, anti-CD40, anti-CD54, anti-CD58, anti-CD80, anti-CD83 (all Immunotech, Berlin, Germany), anti-CD86 (Pharmingen, Palo Alto, Calif.), anti-HLA-DR (Serotec, Camon, USA). As secondary reagents, DTAF-conjugated goat anti-rat IgG (Jackson Immunoresearch, USA), and PE-conjugated donkey antimouse IgG (Jackson) were used. Cytokine analysis For the determination of cytokine production, DC were harvested on day 9 of culture, intensively washed and cultured for 1 day in medium without cytokines, supplemented only with 1% autologous plasma. Subsequently, the supernatants were collected on day 10 of culture and stored at –70 °C. For assessment of cytokine production of T cells after coculture with DC, supernatants were collected after 24 h of coculture. The T cells were restimulated after 7 and 14 days in a primary and secondary restimulation in a plate coated with anti-CD3 antibody (15 µg/ml) and soluble anti-CD28 antibody (10 µg/ml) for 24 h. For the expansion of the T cells, IL-2 (10 U/ml) was added to the culture 3 days after the restimulation. The supernatants were collected 24 h after restimulation and stored at –70°C. Amounts of IL1β, TNF-α, IL-2, IL-4, IL-6, IL-10, IL-12 (p40) and IFN-γ were measured by ELISA using commercially available antibodies and standards according to the manufacturers’ protocols (Pharmingen, Hamburg, Germany; R & D Systems, Wiesbaden, Germany).
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Results The effect of PG on DC is dose-dependent: enhancement of the accessory cell capacity of human DC DC were exposed to PGD2, PGE1, PGE2, PGF2alpha and PGI2 at various concentrations (10–5 to 10–9 M) in the presence of IL-1β, IL-6 and TNF-α on day 7 and the effect on the stimulatory capacity of the DC was tested in an allogeneic MLR with CD4+ T cells as responders. All members of the PG family tested enhanced the stimulatory capacity of DC compared to control DC cultured only in the presence of IL-1β, IL-6 and TNF-α, but to a somewhat different extent (Fig. 1 A). PGE2 was the most potent acti-
Fig. 1 A, B The effects of PG on the stimulatory effect of DC are dose-dependent. Allogeneic CD4+ T cells showed increased proliferation after coculture with PG-stimulated DC. On day 7 of culture, DC were cultured in the presence of IL-1β, IL-6 and TNF-α (mDC mature DC, A) or cultured in medium (GM-CSF and IL-4) alone (iDC immature DC, B) and additionally stimulated with various PG (PGD2, PGE1, PGE2, PGF2alpha, PGI2) for 2 days. The DC were harvested and cocultured with CD4+ T cells in an allogeneic MLR for 72 h as described in Materials and methods. Proliferation was measured by incorporation of 3[H]thymidine for 18 h. The results presented are from five (A) and three (B) experiments
A
B
vator of the APC function of DC as measured by proliferation of cocultured T cells in allogeneic MLR experiments, followed by other PG in the order: PGE1>PGD2> PGF2alpha>PGI2. Proliferation experiments also revealed the dose-dependency of the stimulatory effect of the PG, with different optimal concentrations for each PG: PGE2 10–6 M, PGE1 10–6 M, PGD2 10–7 M, PGF2alpha 10–7 M, PGI2 10–8 M. These optimal concentrations were used in the following experiments. At a concentration of 10–5 M, all PG were toxic to DC, inducing a reduced viability of the cells. The effect of PG was inhibited by the simultaneous addition of anti-PG antisera (20 µg/ml) to the DC cultures (data not shown). To exclude a direct effect of PG on the T cell response, antisera against PG were added to the co-
440 Fig. 2 A, B Enhanced proliferation of naive CD4+ and CD8+ T cells after coculture with PG-treated DC. DC were cultured for 7 days as described in Materials and methods and exposed to IL-1β, IL-6 or TNF-α (as controls) or additionally to various PG (PGD2, PGE1, PGE2, PGF2alpha, PGI2). After extensive washing, DC were cocultured with naive (CD45RA+) CD4+ (A) or CD8+ T cells (B) in an allogeneic MLR. The proliferation was measured in terms of 3[H]thymidine uptake after 72 h. The values presented are the means±SD of triplicate cultures of one representative experiment of four performed
A
B
culture experiments. Compared to control experiments, no effect on T-cell proliferation was observed (data not shown). In some experiments, DC were stimulated with PG on day 7 in the absence of the stimulatory cytokines IL-1β, IL-6 and TNF-α. Similar to the results described above, all PG-treated DC on day 9 of culture induced an increased proliferation of cocultured CD4+ T cells in an allogeneic MLR compared to control cells, indicating a direct effect on DC in the absence of exogenous cytokines (Fig. 1 B). In contrast to fully mature DC stimulated with
the cocktail of cytokines, immature control DC were cultured in the presence of only GM-CSF and IL-4. DC are required for the activation of naive T cells We therefore tested the ability of DC treated with PG to induce the proliferation of allogeneic naive (CD45RA+) CD4+ and CD8+ T cells (Fig. 2 A, B). Compared to control cells, DC exposed to PGE2, PGE1 and PGD2 induced a significantly higher proliferation of both naive CD4+ (Fig.
441 Table 1 Expression of surface molecules by DC cultured as described in Materials and methods. On day 7, the cells were exposed to IL-1β, IL-6, TNF-α (control) or additionally to various PG at appropriate concentrations. After 2 days of stimulation DC were harvested and the expression of surface antigens was measured by FACS analysis. The data shown are the results of five independent experiments, and the values are percentages (values in parentheses are mean fluorescence indicating the expression of HLA-DR)
Treatment
CD1A CD14 CD40 CD54 CD58 CD80 CD83 CD86 HLA-DR
Control (IL-1β, IL-6, TNF-α)
+PGE2 (10–6 M)
+PGE1 (10–6 M)
+PGD2 (10–7 M)
+PGF2alpha (10–7 M)
+PGI2 (10–8 M)
0.8 ± 1.0 2.7 ± 2.1 56.3 ± 10.1 63.2 ± 10.3 54.9 ± 9.6 79.9 ± 11.4 60.7 ± 9.9 76.7 ± 6.9 98.7 (874)
0.5 ± 1.9 1.9 ± 0.5 88.8 ± 12.7 84.3 ± 6.5 88.9 ± 10.2 98.7 ± 9.0 98.7 ± 8.9 98.5 ± 10.5 99.4 (1256)
1.2 ± 1.1 1.0 ± 2.1 85.9 ± 9.5 87.3 ± 11.7 80.8 ± 10.1 97.1 ± 6.8 88.4 ± 10.3 96.3 ± 8.9 98.4 (1065)
3.5 ± 1.2 1.2 ± 0.1 64.8 ± 5.4 80.9 ± 11.7 81.3 ± 9.6 91.0 ± 8.9 85.7 ± 7.5 89.4 ± 10.6 99.5 (1100)
3.7 ± 1.4 2.9 ± 0.7 62.0 ± 10.4 79.3 ± 9.7 81.0 ± 6.5 85.9 ± 6.4 87.4 ± 10.3 84.5 ± 9.8 97.3 (954)
2.5 ± 0.4 3.6 ± 1.5 62.9 ± 9.6 73.5 ± 10.5 85.9 ± 9.7 86.8 ± 6.5 70.6 ± 9.8 88.3 ± 4.6 98.9 (899)
2A) and CD8+ (Fig. 2 B) T cells in all experiments. Also after culture of DC in the presence of PG2alpha and PGI2, an increased T-cell response was observed. However, only a few experiments revealed statistically relevant differences compared to control cells (stimulated with IL-1β, IL-6 and TNF-α alone).
Table 2 Expression of surface molecules by immature DC cultured as described in Materials and methods. On day 7, the cells were cultured in medium alone (with GM-CSF and IL-4) or additionally stimulated with various PG at appropriate concentrations. After 2 days of stimulation DC were harvested and the expression of surface antigens was measured by FACS analysis. The data shown are the results of three independent experiments, and the values are percentages (values in parentheses are mean fluorescence indicating the expression of HLA-DR) Treatment
PG induce maturation of DC To investigate the influence of the PG on the phenotype and the morphology of DC, FACS analysis was performed. Compared to control cells, DC exposed to PG showed a higher expression of costimulatory and adhesion molecules CD40, CD54, CD58, CD80, CD86 and of MHC class II molecules (Table 1). These surface molecules characterized the cells as fully mature DC with a potent capacity for T-cell stimulation. Furthermore, an elevated expression of the human DCspecific antigen CD83 (up to 98%) was found on the surface of PG-exposed DC (Table 1). This result corresponded to the morphological analysis of DC exposed to PG, which revealed the typical morphology of mature DC with distinctive and motile cytoplasmic veils surrounding the cells (data not shown). As in the proliferation experiments, PGE2 was the most potent inducer of a mature phenotype, followed by the other PG in the order PGE1> PGD2>PGF2alpha>PGI2. No immature CD83–/CD1A+ DC or CD14+ macrophages were found in control or PGtreated DC. The addition of PG in the absence of IL-1β, IL-6 and TNF-α led to a higher expression of surface molecules compared to immature control DC, indicating induction of maturation even in the absence of exogenous cytokines (Table 2). PG-treated cells were compared with mature DC (stimulated with IL-1β, IL-6 and TNF-α) and immature control DC cultured only in the presence of GM-CSF and IL-4. The results after addition of PGE1 and PGD2 are shown in Table 2). As in the experiments described above, PGE2 was the most potent inducer of mature DC, followed by the other PG in the order PGE1 >PGD2 >PGF2alpha >PGI2 (data not shown). No significant differences were observed after stimulation with PGF2alpha or PGI2.
CD1A CD14 CD40 CD54 CD58 CD80 CD83 CD86 HLA-DR
Control (GM-CSF+ IL-4)
+PGE2 (10–6 M)
+PGE1 (10–7 M)
+PGD2 (10–7 M)
1.5 ± 3.9 12.9 ± 3.5 32.0 ± 2.4 47.9 ± 9.8 37.7 ± 10.5 69.4 ± 9.9 33.2 ± 5.8 79.4 ± 12.5 88.4 (809)
1.5 ± 3.1 2.9 ± 2.2 59.9 ± 2.5 59.4 ± 1.7 56.9 ± 8.5 85.9 ± 6.3 58.4 ± 7.4 89.3 ± 6.9 97.3 (965)
1.9 ± 2.1 0.9 ± 2.7 50.2 ± 4.7 57.4 ± 1.9 52.9 ± 1.2 80.9 ± 3.1 55.1 ± 3.2 80.3 ± 1.4 97.3 (901)
3.5 ± 1.3 2.2 ± 1.1 44.8 ± 2.2 50.9 ± 11.7 45.6 ± 9.6 77.9 ± 1.9 48.9 ± 5.4 79.7 ± 2.1 96.5 (876)
PG-treated DC produce higher amounts of proinflammatory cytokines and IL-12 To analyse the cytokine production by DC, the cells were exposed to stimulatory cytokines and PG for 2 days as described above. The cells were then washed intensively and cultured for 24 h without any cytokine or PG in medium supplemented only with 1% autologous plasma. Supernatants of DC exposed to PG contained higher amounts of the proinflammatory cytokines IL-1β, IL-6 and TNF-α compared to control cells (Table 3). Furthermore, DC matured in the presence of PG showed a higher capacity to produce IL-12 (p40) compared to control cells (Table 3). IL-10 was not detected in any experiment.
442 Table 3 Production of cytokines by PG-treated DC. DC were generated as described in Materials and methods. On day 7, the cells were exposed to IL-1β, IL-6 and TNF-α (control) or additionally to various PG for 2 days. After extensive washing, DC were cultured for 24 h in medium without any cytokines or PG,
supplemented only with 1% autologous plasma. Subsequently, the supernatants were harvested and cytokine production was measured by ELISA in duplicate. The data show the results of three independent experiments (n.d. not detectable)
Treatment
IL-1β (pg/ml)
IL-6 (pg/ml)
TNF-α (pg/ml)
IL-12 (p40) (pg/ml)
IL-10 (pg/ml)
Control +PGE2 +PGE1 +PGD2 +PGF2alpha +PGI2
2432 ± 765 7765 ± 342 5432 ± 345 4929 ± 563 3432 ± 231 3195 ± 564
7243 ± 487 14543 ± 376 11043 ± 876 9543 ± 432 8932 ± 342 7918 ± 364
5054 ± 432 10764 ± 965 10432 ± 543 8653 ± 476 6943 ± 489 4953 ± 375
3076 ± 234 7654 ± 765 7343 ± 234 6732 ± 362 5914 ± 131 3420 ± 124
n.d. n.d. n.d. n.d. n.d. n.d.
Fig 3 A, B The effects of PG on DC are mimicked by nonphysiological modulators of cAMP. On day 7, DC were incubated with IL-1β, IL-6 and TNF-α (as controls), PGE2 or db-cAMP (10–6 M) or forskolin (50 µM). After 2 days, expression of CD83 on DC was investigated by FACS analysis (A). After coculture with CD4+ T cells in an allogeneic MLR, performed as described above, proliferation was measured in terms of 3[H]thymidine uptake after 72 h. The results presented are representative of three separate experiments (B)
A
443 Fig. 3 B
Table 4 PG-promoted DC maturation amplifies a Th1 cell bias. DC were cultured for 7 days as described, rinsed off the plates and stimulated with IL-1β, IL-6 and TNF-α (control) and additionally exposed to the various PG used in this study at appropriate concentrations (see Results). DC were harvested and cocultured with naive CD4+ T cells. T cells were expanded as described in MateriTreatment
Control +PGE2 +PGE1 +PGD2 +PGF2alpha +PGI2
IL-2 (pg/ml)
als and methods and restimulated with DC (DC/TC ratio 1 : 20). A second restimulation was done with mAb to CD3 and CD28. Supernatants were analysed for cytokine production 24 h after restimulation by ELISA (in duplicate). The results were similar in three independent experiments (n.d. not detectable)
IFN-γ (pg/ml)
IL-4 (pg/ml)
IL-10 (pg/ml)
1. Restimulation with DC
2. Restimulation with mAb
1. Restimulation with DC
2. Restimulation with mAb
1. Re2. Restimulation stimulation with DC with mAb
1. Re2. Restimulation stimulation with DC with mAb
2109 ± 234 7932 ± 165 6987 ± 567 5532 ± 348 4487 ± 438 3987 ± 543
5876 ± 435 16987 ± 897 13876 ± 547 12654 ± 326 9087 ± 548 6973 ± 371
7065 ± 987 21543 ± 997 19876 ± 543 17874 ± 421 14943 ± 231 7943 ± 532
12986 ± 986 29954 ± 937 28987 ± 265 20999 ± 673 19564 ± 754 13987 ± 564
55 ± 25 n.d. n.d. n.d. n.d. 64 ± 43
n.d. n.d. n.d. n.d. n.d. n.d.
85 ± 35 n.d. n.d. n.d. 65 ± 45 73 ± 27
n.d. n.d. n.d. n.d. n.d. n.d.
The effects of PG on DC are mimicked by nonphysiological modulators of cAMP
PG-promoted DC maturation amplifies a Th1 cell bias in the resulting T-cell response
Most of the effects of PG are mediated by the intracellular second messenger cAMP (Phipps et al. 1991). Therefore, we sought to determine whether nonphysiological modulators of cAMP, such as db-cAMP or forskolin, could mimic the capacity of PG to stimulate DC. The effect on the expression of CD83 and the proliferation of CD4+ T cells in an allogeneic MLR compared to PGE2-treated or control cells was analysed (Fig. 3 A, B). Both db-cAMP (10–6 M) and forskolin (50 µM) induced a higher expression of CD83 and an increased T-cell response than in control cells. This effect was dose-dependent for both modulators and limited by the toxicity of the compounds (data not shown). The effect of PGE2 was similar to the effects induced by the nonphysiological modulators. These findings indicate that the effect of PG on DC is mediated by an increase in the intracellular amount of cAMP.
To study whether the maturation of DC in a PG-rich environment affects the type of immune response initiated by these cells, we compared the ability of PG-exposed DC and control DC to induce the production of Th1 (IFN-γ and IL-2) or Th2 (IL-4 and IL-10) cytokines in naive T cells in an allogeneic culture system. T cells were subsequently restimulated with DC generated from the same donor and in a second restimulation with mAb to CD3 and CD28 (Table 4). T cells primed with both control and PG-exposed DC displayed a Th1 cytokine profile after the first and second restimulation. The addition of PG resulted in higher levels of IL-2 and IFN-γ (Table 4). In contrast, only minimal or no amounts of the Th2 cytokines IL-4 and IL-10 were detected. As in the experiments described above, PGE2, PGE1 and PGD2, PGF2alpha were potent inducers of a Th1
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cytokine production, whereas PGI2 promoted only a small increase in Th1 cytokine production compared to control cells (Table 4). Data are shown for experiments with naive CD4+ T cells. Similar results were obtained after coculture with naive CD8+ T cells (data not shown).
Discussion In the present study, we demonstrated that various PG promote the maturation and stimulatory capacity of human DC and enhance the development of a Th1 immune response. Among the PG used in our system, PGE2 was the most potent modulator of APC. With regard to the effect of PG on the function of APC, we analysed the properties of human DC as the most potent APC of the immune system after stimulation with PG. Our results demonstrate that all members of the PG family are able to induce the differentiation and maturation of human DC and to affect the resulting T-cell response by increasing the production of IL-2 and IFN-γ. Analysis of cytokine production by activated DC showed an elevated level of proinflammatory cytokines (IL-1β, IL-6 and TNF-α) which by themselves are known to induce DC maturation (Jonuleit et al. 1997). Furthermore, an increased amount of IL-12 was observed in the supernatants of DC stimulated with PG. This might explain the resulting Th-biased T-cell response because IL-12 is a crucial factor in the generation of a Th1-type response (Trinchieri and Scott 1994). Our results are supported by the work of Rieser et al. (1997). These authors demonstrated that PGE2 is capable of inducing the maturation of human DC generated from blood progenitors resulting in higher amounts of IL-12 production compared to control cells (Rieser et al. 1997). Furthermore, these investigators showed that after deactivation of DC with endotoxin, PGE2 restores IL-12 production and DC maturation (Rieser et al. 1998). Our results are in contrast to those reported by Kalinski et al. These investigators have reported a reduced production of IL-12 by human DC cultured in the presence of PGE2 (and IL-1β and TNF-α) compared to control cells (Kalinski et al. 1997; Kalinski et al. 1998). In this study, investigations of the supernatants after coculture with naive T cells revealed reduced levels of IL-2 and IFN-γ and increased amounts of IL-4 and IL-5, indicating a Th2 bias of the immune response (Kalinski et al. 1997). Several aspects might explain these differences. Kalinski et al. used RPMI-1640 complemented with FCS, generating immature DC characterized by a strong expression of CD1a and a weak expression of CD83. These subpopulations of DC are known as immature DC with high antigen-processing but low antigen-presenting abilities. In our experiments X-VIVO15 with 1% autologous plasma was used resulting in a high percentage of mature CD83+/ CD1a– DC. Thus two different populations of human DC were generated, which appeared to differ in their response to PG. A further explanation for the different results may be the various cytokines used for the generation of mature
DC. Whereas all groups used GM-CSF and IL-4 to generate immature DC, the additional stimulation was performed with TNF-α, IL-1β and PGE2 (Kalinski et al.), TNF-α, and PGE2 (Rieser et al.) or TNF-α, IL-1β, IL-6 and PGE2 (our laboratory) to obtain fully mature DC (Kalinski et al. 1997; Kalinski et al. 1998; Rieser et al. 1997). Also, Kalinski et al. performed allogeneic MLR experiments with DC/TC ratios of 1 : 1 or 1 : 2, whereas we used ratios varying between 1 : 10 and 1 : 40, reflecting the biological conditions in lymphatic organs as the site of antigen-presentation and T-cell stimulation. The differences in the properties and function of the two populations of DC also may be due to a different pattern of PG receptors on their surfaces. Five types of receptors (DP, EP, FP, IP, TP) have been described, with various cell distribution and biological functions, acting as receptors with a high rate of crossreactivity to prostaglandins of other families (Coleman et al. 1994). Nothing is yet known about the expression of prostanoid receptors on DC. Our findings suggest the involvement of the cAMP signaling pathway in DC maturation because the addition of db-cAMP and forskolin, which is known to increase intracellular levels of cAMP (Phipps et al. 1991), mimicked the effects of the PG. In contrast to the findings of Kalinski et al., the addition of PGE2 increased the production of the proinflammatory cytokines IL-1β, IL-6 and TNF-α in DC in our system. Similarly, it has been demonstrated that PGE1 and PGE2 enhance the levels of cAMP and cGMP in human macrophages, which is paralleled by an increased activation and TNF-α production (Gemsa et al. 1977; Renz et al. 1988). Direct stimulation with db-cAMP converts human macrophages to highly active accessory cells with a high level of HLA-DR expression, downregulation of macrophage markers but sustained CD14 expression (Peters et al. 1990). Furthermore, in human monocytes, cAMP synergizes with TNF-α to upregulate the synthesis of IL-1β, suggesting that TNF signals and the PG pathway cooperate via elevation of the intracellular second messenger cAMP to induce cytokine production by APC (Lorenz et al. 1995). The results presented in this report show that all PG of the arachidonic family are able to induce the maturation of human DC and to enhance the capacity to promote a Th1 response. This effect was demonstrated in the presence of proinflammatory cytokines indicating a synergistic influence of PG and cytokines to support the physiological role of activated DC in an inflammatory environment induced by pathogens, allergens, injuries or autoimmune diseases.
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