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Akt and NF-κB pathways
Molecular Immunology 44 (2007) 2686–2696
Triptolide impairs dendritic cell migration by inhibiting CCR7 and COX-2 expression through PI3-K/Akt and NF-B pathways Qiuyan Liu a,b , Taoyong Chen a , Guoyou Chen a , Xiaoli Shu b , Anna Sun b , Pengcheng Ma c , Liwei Lu d , Xuetao Cao a,b,∗ a
Institute of Immunology and National Key Laboratory of Medical Immunology, Second Military Medical University, Shanghai 200433, PR China b Institute of Immunology, Zhejiang University, Hangzhou 310031, PR China c Institute of Dermatology, Chinese Academy of Medical Sciences, Nanjing 210042, PR China d Department of Pathology, The University of Hong Kong, Hong Kong, PR China Received 20 October 2006; accepted 6 December 2006 Available online 16 January 2007
Abstract Inhibition of dendritic cell (DC) migration into tissues and secondary lymphoid organs is an efficient way to induce immunosuppression and tolerance. CCR7 and PGE2 are critical for DC migration to secondary lymphoid organs where DC initiate immune response. Triptolide, an active component purified from the medicinal plant Tripterygium Wilfordii Hook F., is a potent immunosuppressive drug capable of prolonging allograft survival in organ transplantation by inhibiting T cell activation and proliferation. Considering the essential role in T cell tolerance of DC migration to secondary lymphoid organs, here we demonstrate that triptolide can significantly inhibit LPS-triggered upregulation of CCR7 expression and PGE2 production by inhibiting cyclooxygenase-2 (COX-2) expression in DC, thus impairing DC migration towards CCR7 ligand CCL19/MIP-3 in vitro. Moreover, triptolide-treated DC display impaired migration into secondary lymphoid organs and in vivo administration of triptolide also inhibits DC migration. Further studies show that the triptolide-mediated inhibitory effects of LPS-induced activation of phosphatidylinositol-3 kinase (PI3-K)/Akt and nuclear NF-B activation are involved in down-regulation of COX-2 and CCR7 expression resulting in impaired migration to secondary lymphoid organs of DC. Therefore, inhibition of DC migration through decreasing COX-2 and CCR7 expression via PI3-K/Akt and NF-B signal pathways provides additional mechanistic explanation for triptolide’s immunosuppressive effect. © 2006 Elsevier Ltd. All rights reserved. Keywords: Dendritic cell; Migration; CCR7; Immunosuppression; Signal transduction
1. Introduction Triptolide, a diterpene triepoxide, is an active component of extracts derived from the medicinal plant Tripterygium Wilfordii Hook F. (TWHF), and has anti-inflammatory and immunosuppressive activities (Qiu and Kao, 2003). Tripterygium extracts have, for many years, been used widely to treat autoimmune dis-
Abbreviations: trip, triptolide; DC-trip1, DC were treated with triptolide 1 ng/ml; DC-trip1/LPS, DC were treated with triptolide 1 ng/ml and stimulated with LPS; DC-trip5, DC were treated with triptolide 5 ng/ml; DC-trip5/LPS, DC were treated with triptolide 5 ng/ml and stimulated with LPS ∗ Corresponding author at: Institute of Immunology and National Key Laboratory of Medical Immunology, Second Military Medical University, 800 Xiangyin Road, Shanghai 200433, PR China. Tel.: +86 21 55620605; fax: +86 21 65382502. E-mail address: [email protected] (X. Cao). 0161-5890/$ – see front matter © 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.molimm.2006.12.003
eases including rheumatoid arthritis, immune complex nephritis, and systemic lupus erythematosus in China (Ramgolam et al., 2000; Tao and Lipsky, 2000). Clinical and experimental studies have also demonstrated that Tripterygium extracts effectively prolong allograft survival in organ transplantation including bone marrow, cardiac, renal and skin transplantation (Chen et al., 2002; Fidler et al., 2002; Wang et al., 2000; Yang et al., 1992). The immunosuppressive effects by triptolide can be partly attributed to its potent inhibitory effects on T cell activation and interleukin-2 (IL-2) production (Chen et al., 2000; Qiu et al., 1999). Dendritic cells (DC), the most potent professional antigenpresenting cells (APC), are critical for the induction of both immune responses and immune tolerance (Banchereau and Steinman, 1998; Banchereau et al., 2000; Hackstein and Thomson, 2004; Sallusto and Lanzavecchia, 1999). DC originate from bone marrow precursors and migrate through the blood
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stream to almost every tissue, where they reside in an immature state. At the site of infection, DC recognize and respond to common pathogen-associated molecular patterns (PAMPs), including lipopeptides, lipopolysaccharides (LPS) and nucleic acids via Toll-like receptors (TLR) (Ricciardi-Castagnoli and Granucci, 2002). LPS activates DC and induces production of proinflammatory cytokines and other mediators via TLR4, thus triggering a complex phenotypic and functional maturation program of DC including up-regulation of the chemokine receptor CCR7 (Sallusto et al., 1998). The CCR7 upregulation enable DC more responsiveness to CCR7 ligands, which promotes DC migration from peripheral tissues to secondary lymphoid organs such as lymph nodes (LN) and spleen where DC present antigens to T cells and initiate immune response (MartIn-Fontecha et al., 2003; Randolph, 2001). The expression of CCR7 on mature DC is essential for their migration to the T-cell area of draining LN because this migration is guided by two ligands for CCR7, CCL19 (EBI1-ligand chemokine [ELC], macrophage inflammatory protein [MIP]-3), and CCL21 (secondary lymphoid-tissue chemokine [SLC], 6-Ckine). Both chemokines are expressed by stromal cells in the T-cell area of secondary lymphoid organs (Ohl et al., 2004). The essential role of CCR7 and its ligands in mature DC migration to LN was demonstrated in CCR7deficient mice (Beckmann et al., 2004) and plt/plt mice, which lack the ligands for CCR7 (Luther et al., 2000). Because the migration of DC into secondary lymphoid organs is required for the initiation of specific immune responses, so, inhibition of DC migration into secondary lymphoid organs by inhibiting CCR7 expression is an efficient way to induce immunosuppression and tolerance. PGE2 , derived from metabolism of free arachidonic acid, exerts an important immunomodulatory role in DC differentiation and function. Previous studies have shown that PGE2 is required for CCR7-expressed DC migration to its ligands CCL19 and CCL21 (Kabashima et al., 2003; Scandella et al., 2002; Scandella et al., 2004), the importance of PGE2 for DC migration in vivo has recently been shown in Ptger4−/− mice lacking the PGE2 receptor EP4. These mice display impaired migration of Langerhans cells and reduced skin immune responses (Legler et al., 2006). Besides its inhibitory effects on T cell function, triptolide may also affect other cells involved in immune responses. We and other researchers have shown that triptolide can affect DC functional maturation at lower or clinical concentrations (Chen et al., 2005; Liu et al., 2006; Zhu et al., 2005) and induce DC apoptosis at high concentrations (≥10 ng/ml) (Liu et al., 2004). Considering that the migration of DC into secondary lymphoid organs is required for the initiation of immune responses and CCR7 on DC is a potential target for immunosuppressive drug, in this study, we investigated the effect of triptolide, used at clinical concentration that does not induce DC apoptosis, on DC migration in response to LPS stimulation. We demonstrate that triptolide can significantly inhibit LPS-triggered upregulation of CCR7 expression and PGE2 production by inhibiting cyclooxygenase-2 (COX-2) expression in DC, thus impairing DC migration towards CCR7 ligand CCL19/MIP-3 in vitro, and in vivo administration of triptolide also inhibits DC migration to the spleen and lymphoid
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nodes. Furthermore, triptolide-mediated inhibitory effects of LPS-induced activation of PI3-K/Akt and nuclear NF-B activation are involved in down-regulation of COX-2 and CCR7 expression resulting in impaired migration to secondary lymphoid organs of DC. Therefore, inhibition of DC migration through decreasing COX-2 and CCR7 expression via PI3-K/Akt and NF-B signal pathways provides additional mechanistic explanation for triptolide’s immunosuppressive effect. 2. Materials and methods 2.1. Mice and reagents Male wild-type C57BL/6 mice, 5–6 weeks of age, were purchased from SIPPR-BK Experimental Animal Co. (Shanghai, China). GFP+/+ C57BL/6-TgN (ACTbEGFP) mice were obtained from the Jackson Laboratory (Bar Harbor, ME). All mice were housed in a pathogen-free facility for all experiments. Crystalline triptolide (PG490, molecular weight 360, purity of 99%) was obtained from the Institute of Dermatology, Chinese Academy of Medical Sciences (Nanjing, PR China) and prepared as previously described (Mao et al., 1998). Triptolide was reconstituted in dimethyl sulphoxide (DMSO) stock solutions (1 mg/ml) and stored at −20 ◦ C. Triptolide was freshly diluted to the indicated concentrations with culture medium before use. The DMSO concentration in all test conditions never exceed 0.002% (V/V). Recombinant mouse GM-CSF and IL-4 were purchased from R&D Systems (Minneapolis, MN). Chemokine receptor antibodies (CCR1, CCR5, CCR7 and CXCR4), polyclonal Abs against actin, nucleoporin p62, monoclonal Ab against COX-2 and monoclonal Ab for NF-B p65 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). PE-labeled CCR7 Ab was purchased from ebioscience (San Diego, CA). Phospho-Abs against extracellular signal-regulated kinase p44/p42 (ERK1/2, Thr202/Tyr204), c-Jun N-terminal kinase/stress associated protein kinase (JNK/SAPK, Thr183/Tyr185), p38 (Thr180/Tyr182), and corresponding Abs against non-phosphorylated signaling proteins were from Cell Signaling (Beverly, MA). ELISA kits for murine PGE2 , TNF-␣, IL-1 and IL-6 and recombinant murine MIP-1␣ and MIP-3 were purchased from R&D Systems (Minneapolis, MN). Lipopolysaccharide (LPS), PGE2 and PI3-K/Akt inhibitor Wortmannin and COX-2 specific inhibitor NS398 were from Calbiochem (San Diego, CA). 2.2. Generation of mouse bone marrow (BM)-derived DC Mouse bone marrow-derived DC were prepared from bone marrow by culturing with 10 ng/ml recombinant GM-CSF and 1 ng/ml IL-4, as described previously (Chen et al., 2004). Following 6 days of culture, DC were harvested and purified using anti-CD11c-coated microbeads (Miltenyi-Biotec, Auburn, CA), then a highly enriched population of DC (90–95% CD11c+ cells) was obtained. DC were pretreated with triptolide for 12 h, and then LPS or LPS and PGE2 treatments were performed as indicated. In some experiments, DC were pretreated with signal inhibitors for 30 min before LPS or LPS and PGE2 stimulation.
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2.3. ELISA assay Cytokines (TNF-␣, IL-1, IL-6) and PGE2 contained in the culture supernatants of DC treated with or without triptolide after LPS stimulated or not were measured by ELISA (R&D, Minneapolis, MN) according to the manufacturer’s instructions. 2.4. Assay for chemokine receptor expression For RT-PCR assay of chemokine receptor mRNA expression, total cellular RNA was isolated from DC following indicated stimulation with TRIzol reagent according to the manufacturer’s instructions (Life Technology). cDNA was synthesized from 1 g of total RNA by extension of oligo(dT)18 primer with 200 units AMV (Promega, Madison, WI). Primers used for chemokine receptors were as follows: 5 GTG TTC ATC ATT GGA GTG GTG 3 and 5 GGT TGA ACA GGT AGA TGC TGG TC 3 for CCR1; 5 CAT CGA TTA TGG TAT GTC AGC ACC 3 and 5 CAG AAT GGT AGT GTG AGC AGG AA 3 for CCR5; 5 CCA GGA AAA ACG TGC TGG TG 3 and 5 GGC CAG GTT GAG CAG GTA GG 3 for CCR7; 5 CGG CAA TGG ATT GGT GAT CCT GGT C 3 and 5 GAG GGC CTT GCG CTT CTG GTG GCC C 3 for CXCR4. Primers used for mouse -actin were 5 GTG GGA ATT CGT CAG AAG GAC TCC TAT GTG 3 and 5 GAA GTC TAG AGC AAC ATA GCA CAG CTT CTC 3 . Equal amounts of total cellular RNA were used for reverse transcription under the same conditions. cDNA levels were semi-quantified by inclusion of a -actin control. PCR was performed at 94 ◦ C 30 s, 55 ◦ C 30 s and 72 ◦ C 30 s for 25 cycles, followed by 10 min elongation at 72 ◦ C. PCR products were separated on a 1% agarose gel and analyzed using a Gel Doc 2000 system (Bio-Rad, Hercules, CA). Western blot was performed to analyze chemokine receptor expression in DC at the protein level as described previously (Chen et al., 2004). For FACS assays, DC were labeled with chemokine receptor primary Ab in PBS for 30 min, and then washed, stained with FITC-labeled secondary Ab, and then analyzed by FACS as described previously (Chen et al., 2004). 2.5. In vitro DC chemoattraction assay In vitro chemoattraction of DC in response to macrophage inflammatory protein (MIP)-1␣ or MIP-3 was measured as previously described (Chen et al., 2004). 2 × 105 DC were added to the upper chamber of 24-well transwell chambers with 5 mpore size polycarbonate filters (Corning Costar, Cambridge, MA) in a total volume of 100 l. The chambers were incubated for 4 h at 37 ◦ C. Cells migrating to the bottom chamber were labeled with FITC-CD11c (DC) Ab for 30 min at 4 ◦ C and then counted by flow cytometry. Each experiment was performed in triplicate. 2.6. In vivo DC migration assays To determine whether the migration of triptolide-treated DC to secondary lymphoid organs was impaired, we transplanted BALB/c skin sheets (10–15 mm pieces) to C57BL/6 mice.
Two days after transplantation, in vitro-cultured C57BL/6 DC, with or without triptolide treatment (5 ng/ml for 24 h), were labeled with the red fluorescence marker PKH-26 (Sigma) as described (Chen et al., 2004), and immediately injected into skin allografts (1 × 106 in 100 l). Six hours after injection, mice were killed and ipsilateral inguinal lymph nodes and spleens harvested, embedded in optimal cutting temperature (OCT) compound (IEC, Needham Heights, MA), and frozen immediately at −80 ◦ C. Cryostat sections (6 m) were placed on cover slides, fixed in 1% paraformaldehyde for 5 min, incubated with 0.4 g/ml Hoechst (Calbiochem) for 5 min, and subsequently examined by fluorescence microscopy. Cell suspensions from lymph nodes and spleen were also prepared and PKH26-positive cells contained in these secondary lymphoid organs counted by FACS. To exclude the influence of in vitro labeling on DC function, green DC derived from GFP+/+ (green fluorescent protein) C57BL/6 mice were also used instead of PKH-26-labeled DC. To mimic normal/physiological in vivo DC migration as accurately as possible, C57BL/6 mice were intraperitoneally injected with triptolide (5 g/kg) every day for a total of 7 days, and FITC was painted onto the abdomen as described previously (Macatonia et al., 1987), then the mice were intravenously injected with LPS (5 mg/kg) via the tail vein 12 h after the final injection. Spleen and draining lymph nodes were isolated, and cell suspensions were prepared. To exclude the influence of DC apoptosis induced by triptolide in vivo, DC in the cell suspensions were stained with PE-labeled CD11c and 7-AAD, and FITC/PE double-positive and 7-AAD negative cells detected by FACS. Results are representative of n = 4 mice in each group. 2.7. Assay of PI3-K activity PI3-K activity was assayed with PI3-K ELISA kit (Echelon Biosciences Inc., Salt Lake City, UT) according to manufacturer’s instructions. In brief, DC were pretreated with triptolide (5 ng/ml) for 12 h or pretreated with PI3-K/Akt inhibitors for 30 min and then stimulated with LPS for 15 min. Cell lysates were prepared, PI3-K was immunoprecipitated with antibody against the p85 subunit and incubated with PI(4,5)P2. The reaction products were incubated with a PI(3,4,5)P3 detector protein, then added to PI(3,4,5)P3-coated microplate for competitive binding. A peroxidase-linked secondary detection reagent and colorimetric detection was used to detect PI (3,4,5)P3 detector protein binding to the plate. The colorimetric signal was inversely proportional to the amount of PI(3,4,5)P3 produced by PI3-K activity. 2.8. Assay of Akt kinase activity Akt activity was detected using Akt kinase Assay kit (Cell Signaling Technology Inc., Beverly, MA) according to manufacturer’s instructions. Briefly, DC were treated as above, cell lysates were prepared and immobilized Akt primary antibody was used to immunoprecipitate Akt from the extracts. GSK-3 fusion protein (1 g) was incubated with the precipitated protein in kinase buffer and then detected with Phospho-GST-3 specific
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antibody by Western blot. The phosphorylation level of GSK-3 fusion protein was proportional to Akt activity. 2.9. Immunoprecipitation and Western blotting DC were lysed with M-PERTM Protein Extraction Reagent (Pierce, Rockford, IL) supplemented with protease inhibitor cocktail, and protein concentrations of the extracts measured by BCA assay (Pierce, Rockford, IL). Forty micrograms of the protein was used either for immunoprecipitation, or loaded per lane, subjected to SDS-PAGE, transferred onto nitrocellulose membranes, then blotted as described previously (An et al., 2002). For detection of the nuclear translocation of NF-B p65, cytoplasmic and nuclear extracts were prepared using NE-PER nuclear and cytoplasmic extraction reagents (Pierce, Rockford, IL) and protein concentration was determined with a BCA-200 protein assay kit. Nuclear NF-B p65 subunit was detected by Western blot using specific antibody. 2.10. Statistical analysis Comparisons between experimental groups and relevant controls were performed by Student’s t-test. P < 0.05 was considered a statistically significant difference. 3. Results 3.1. Triptolide inhibits CCL19/MIP-3β-induced DC migration both in vitro and in vivo One characteristic of mature DC is their ability to migrate to secondary lymphoid organs where DC present antigens to T cells and initiate immune response. So, inhibition of DC migration to secondary lymphoid organs is an efficient way to induce immunosuppression and tolerance. To confirm whether triptolide affects DC migration, we examined in vitro migration of DC toward MIP-1␣ and MIP-3, two chemokines chemotactic for immature and mature DC respectively. As expected, immature DC efficiently responded to MIP-1␣ while LPS-matured DC responded well to MIP-3 (Fig. 1A and B). In contrast, after pretreatment with triptolide, LPS-matured DC responded poorly to MIP-3 (Fig. 1B). Interestingly, these mature DC retained their capability to respond to MIP-1␣ (Fig. 1A). To further investigate the possibility that triptolide treatment impaired either the capacity of DC to emigrate out of skin epidermis or to immigrate into secondary lymphoid organs, we performed in vivo experiments to examine the migratory capacity of DC in mice treated with triptolide. DC were treated with triptolide, and then labeled with PKH-26, and subcutaneously injected into skin transplants of C57BL/6 mice derived from BALB/c mice (1 × 106 DC per mouse), they showed dramatically reduced immigration into the spleen and lymph nodes as compared with untreated control DC (Fig. 1C). PKH-26-labeled cell numbers in spleen and lymph nodes were also enumerated by FACS; PKH-26-positive cells in secondary lymphoid organs were reduced in the triptolide treatment group (Fig. 1D). When GFP-expressing DC derived from C57BL/6GFP+/+ mice
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were used instead of PKH-26-labeled DC, similar results were observed (Fig. 1E). In the FITC-painting assay, following LPS mobilization, FITC-positive DC numbers in lymph nodes and spleen of triptolide-treated mice were significantly reduced than that in normal mice (Fig. 1F). Our study also demonstrated that triptolide-pretreated DC exhibited a reduced allostimulatory activity (data not shown). Taken together, these results demonstrate that administration of triptolide not only impairs the migration of DC into secondary lymphoid organs, but also reduces allostimulatory activity of DC. 3.2. Triptolide inhibits LPS-induced CCR7 expression in DC It’s well known that LPS induces maturation of DC with upregulation of CCR7 which binds to its ligand CCL19/MIP3 and directs DC migration to secondary lymph organs, this process is critical for DC to initiate immune response. To confirm the reduced migration of DC by triptolide to secondary lymphoid organs is associated with the inhibited CCR7 expression, we first investigated the chemokine receptor expression in triptolide-treated DC with or without LPS stimulation. DC were pretreated with triptolide for 12 h and then stimulated with LPS (1 g/ml) for 12 h for RT-PCR and FACS analysis or 24 h for Western blot assay. We found that triptolide pretreatment decreased the LPS-induced upregulation of CCR7 and CXCR4 in DC both at the mRNA (Fig. 2A) and protein level (Fig. 2B and C). In contrast, expression of CCR1 and CCR5 in DC was not decreased by triptolide. These results demonstrate that triptolide can significantly inhibit LPS-induced CCR7 expression in DC. Furthermore, the impaired LPS-induced upregulation of CCR7 expression in triptolide-treated DC may account for their reduced responsiveness to MIP-3 while the constitutive expression of CCR1 and CCR5 in triptolide-treated DC may be responsible for their continued responsiveness to MIP-1␣. 3.3. Triptolide inhibits COX-2 expression and PGE2 production in DC PGE2 , derived from metabolism of free arachidonic acid, exerts an important immunomodulatory role in DC differentiation and function (Kabashima et al., 2003; Scandella et al., 2002, 2004). Cyclooxygenases (COX-1 and COX-2) are the major enzymes involved in PGE2 production (Dubois et al., 1998; Smith et al., 2000). COX-1 is constitutively expressed in most tissues and appears to mediate various physiological functions (Smith et al., 1996; Smith and DeWitt, 1995). By contrast, COX-2 is undetectable in most normal tissues, but is rapidly induced in response to inflammatory stimuli, including TNF␣, IL-1␣ and LPS (Harizi et al., 2002; Sharma et al., 2003). It was shown that both extracts of TWHF and triptolide could inhibit PGE2 production in a variety of human cells by blocking the upregulation of COX-2 (Tao et al., 1998; Yao et al., 2005). So we wondered whether triptolide impairs DC migration by inhibiting LPS-induced COX-2 expression and resultant PGE2 production in DC. As shown in Fig. 3A, triptolide treatment of DC resulted in a decrease of LPS-stimulated COX-2 expres-
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Fig. 1. Effect of triptolide on DC migration in vitro and in vivo. (A and B) Triptolide causes LPS-stimulated DC to retain responsiveness to MIP-1␣ (A) but inhibits responsiveness of LPS-stimulated DC to MIP-3 (B). DC were cultured and treated as described in Section 2, then evaluated for responsiveness to MIP-1␣ and MIP-3 by in vitro chemoattraction assay. 2 × 105 triptolide-treated DC were resuspended in 0.1 ml RPMI 1640 containing 0.5% BSA, and loaded into the upper well of a transwell chamber. Lower compartment were filled with RPMI 1640 containing 0.5% BSA, with or without MIP-1␣ (100 ng/ml) or MIP-3 (100 ng/ml), respectively. The chambers were incubated for 4 h at 37 ◦ C. Directed migration was expressed as the number of CD11c+ cells that had migrated to the lower chamber, counted by FACS analysis. Each experiment was performed in triplicate at least three times. Data is expressed as mean number of migrated CD11c+ cells ±S.D. * P < 0.05, ** P < 0.01. (C and D) Triptolide treatment impairs PKH-26-labeled murine DC migration into spleen and lymph nodes (LN). (C) Immunofluorescence microscopy of PKH-26-positive cells in spleen and LN. Results here are representative of sections derived from at least three samples. Red cells are PKH-26 labeled DC, and blue cell nucleus is stained with Hoechst. Original magnification, 400×. (D) FACS enumeration of PKH-26-positive cells in spleen and LN. (E) FACS analysis of DC GFP+ cells (R2 region) in spleen and LN following injection into skin transplants. Labels indicate percentage of GFP+ cells in cell suspensions. (F) Triptolide decreases the frequency of FITC+ CD11c+ DC in skin migration to spleen and LN. C57BL/6 mice were treated with triptolide (5 g/kg) for 7 days as described in Section 2, and then FITC was painted onto the abdomen, the mice were intravenously injected with LPS (5 mg/kg) via the tail vein 12 h after the final injection. Spleen and draining LN were isolated, and cell suspensions were prepared. DC in the cell suspensions were stained with PE-labeled CD11c and 7-AAD, and FITC/PE double-positive and 7-AAD negative cells detected by FACS. Results represent three independent experiments. * P < 0.05, ** P < 0.01.
sion at both the mRNA and protein level. Importantly, PGE2 levels in the culture supernatant were also reduced significantly (Fig. 3B), suggesting that triptolide inhibits PGE2 production by decreasing COX-2 expression in DC. 3.4. Exogenous PGE2 overcomes the inhibited DC migration and reverses CCR7 expression We and other researchers have shown that PGE2 promotes DC migration in response to chemotactic factors by modulating
chemokine receptor expression (Scandella et al., 2002; Chen et al., 2004; Luft et al., 2002). To further investigate the role of the reduced PGE2 in triptolide-mediated inhibition of DC migration, we added exogenous PGE2 into DC migration assay, and found addition of exogenous PGE2 (1 M) restored the migratory capacity of triptolide-treated DC in chemoattraction assays (Fig. 4A). We then examined whether PGE2 also restored CCR7 expression in triptolide-treated and LPS-stimulated DC. In the presence of exogenous PGE2 , LPS-stimulated DC expression of CCR7 was also restored (Fig. 4B and C). These results are con-
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Fig. 3. Triptolide inhibits LPS-induced COX-2 expression and PGE2 production in DC. (A) Triptolide inhibits LPS-induced COX-2 expression. DC were treated as indicated and COX-2 expression assayed by RT-PCR (upper two panels) and Western blot assays (lower two panels). (B) Supernatants from triptolide-treated or -untreated DC, stimulated or unstimulated with LPS (1 g/ml) for 24 h, were examined for PGE2 production by ParameterTM PGE2 Assay.
leading to the impaired migration of DC to secondary lymphoid organs.
Fig. 2. Effect of triptolide on LPS-induced chemokine receptor expression in DC. (A) RT-PCR and (B) Western blot assay of chemokine receptor expression in DC pretreated with triptolide for 12 h or then stimulated with 1 g/ml LPS for 12 h or 24 h, respectively. -Actin (RT-PCR) and actin (Western blot) were used as a quantitative control. (C) DC pretreated with triptolide for 12 h and then stimulated with 1 g/ml LPS for 12 h, then labeled with Abs against chemokine receptors in PBS for 30 min, washed, and finally stained with FITC-labeled secondary Ab and analyzed by FACS. Unshaded regions represent isotype controls and shaded regions corresponding Abs, as indicated. Results were expressed as fluorescence intensity (FI). Data were expressed as mean (n = 3). There are significant differences in expression of CCR1, CCR5, CCR7 and CXCR4 between triptolide-treated DC with or without LPS stimulation. * P < 0.05, ** P < 0.01.
sistent with reports showing that PGE2 regulates the expression of CCR7 on DC and their migration. These results suggest that the inhibition of DC migration by triptolide may be partially due to its inhibition of COX-2 expression, which in turn results in a defective production of PGE2 and CCR7 expression in DC, thus
3.5. Triptolide impairs LPS-induced proinflammatory cytokines production in DC It is well known that COX-2 is undetectable in most normal tissues but is rapidly induced in response to proinflammatory cytokines (Harizi et al., 2002; Sharma et al., 2003), LPS activates DC and triggers production of proinflammatory cytokines including TNF-␣, IL-1 and IL-6. To investigate whether triptolide inhibits LPS-induced proinflammatory cytokines production, 5 × 105 cells/ml DC were pretreated with triptolide (5 ng/ml) for 12 h, which were then stimulated with or without LPS (1 g/ml) for indicated time. RT-PCR and ELISA results showed that triptolide inhibited both mRNA (Fig. 5A) and protein (Fig. 5B) expression of proinflammatory cytokines including TNF-␣, IL-1 and IL-6 by DC. Moreover, triptolide inhibited proinflammatory cytokines production by DC in a dose-dependent manner, even at very low concentrations (1 ng/ml) (Fig. 5C).
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3.6. Inhibition of PI3-K/Akt kinase activity and NF-κB nuclear activation contributes to the impaired COX-2 and CCR7 expression in DC by triptolide
Fig. 4. Exogenous PGE2 rescues the triptolide-induced migratory impairment of DC and restores CCR7 expression. (A) Exogenous PGE2 rescues migration of triptolide-treated DC to CCL19/MIP-3. DC were pretreated with triptolide for 12 h, and then stimulated with LPS or LPS and PGE2 together for 24 h, evaluated for responsiveness to CCL19/MIP-3 by in vitro chemoattraction assay as above described. * P < 0.05, ** P < 0.01. (B and C) Exogenous PGE2 restores CCR7 expression. (B) RT-PCR (upper panels) and Western blot (lower panels) and (C) FACS analysis assay of CCR7 expression in DC following supplementation with PGE2 (1 M).
It was reported that MAPK, PI3-K/Akt signal pathways and NF-B transcription regulate COX-2 and CCR7 expression in different cell lines (Ardeshna et al., 2000; Chang et al., 2004; Lee et al., 2003; St-Germain et al., 2004). We examined whether these signaling pathways are involved in the triptolide-mediated inhibitory effect on COX-2 and CCR7 expression in DC. Upon LPS stimulation, the ERK, JNK, and p38 pathways in DC were rapidly activated, and triptolide treatment did not significantly affect the activation of these pathways in DC (data not show), suggesting that triptolide does not interfere with the MAPK pathway in LPS-treated DC. We then examined whether PI3-K/Akt signaling pathway is involved in triptolide-mediated inhibition of COX-2 and CCR7 expression in DC. As shown in Fig. 6A, triptolide treatment decreased PI3-K and Akt/PKB kinase activity in LPS-stimulated DC. We further examined NF-B activation in triptolide-treated DC. Under normal conditions, the majority of NF-B subunits are sequestered in the cytoplasm by I-B␣ and translocated into nucleus following I-B␣ degradation. We found that total I-B␣ levels in cell lysates of LPS-stimulated control DC were rapidly decreased while that of the LPS-stimulated and triptolide-treated DC showed no significant changes (Fig. 6B), indicating that triptolide inhibits LPS-induced I-B␣ degradation in DC. In consistent with the inhibited degradation of I-B␣ in triptolidetreated DC in response to LPS, nuclear levels of NF-B p65 subunit in triptolide-treated and LPS-stimulated DC were also lower than that of LPS-treated DC without triptolide treatment (Fig. 6B). To further understand the role of the reduced PGE2 in triptolide-mediated inhibition of PI3-K/Akt and NF-B pathway, we added exogenous PGE2 together with LPS into culture system after DC were pretreated with triptolide for 12 h. As shown in Fig. 6C, exogenous PGE2 restores the triptolideinduced inhibition of Akt kinase activity and nuclear NF-B activaction in DC. Furthermore, we investigated the effects of inhibitors of PI3-K/Akt, NF-B and COX-2 pathways on DC migration toward MIP-3. As shown in Fig. 6D, inhibitor of PI3K/Akt, NF-B or COX-2, when used respectively, could inhibit DC migration significantly. However, combined used of any kind of these inhibitors could not further increase the inhibitory effect of triptolide on DC migration toward MIP-3, further confirming the triptolide-mediated inhibition of DC migration is through PI3-K/Akt, NF-B pathways. Taken together, these results suggest that triptolide-mediated inhibition of COX-2 and CCR7 expression by decreasing TLR4 ligand-induced PI3K/Akt activity and NF-B activation in DC, resulting in the impairment of DC migration to secondary lymphoid organs. 4. Discussion Although triptolide has been used extensively as an immunosuppressive agent for the treatment of various allograft organ transplantation and autoimmune diseases, the inhibitory effects
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Fig. 5. Triptolide inhibits LPS-induced proinflammatory cytokines production by DC. DC (5 × 105 /ml) were pretreated with triptolide (5 ng/ml) for 12 h, and then stimulated with (DC-LPS, DC-Trip/LPS) or without (DC, DC-Trip) 1 g/ml LPS at indicated time points and the culture supernatants of the resting and stimulated DC were collected. RT-PCR assay of cytokines (TNF-␣, IL-1, IL-6) mRNA expression (A) and ELISA assay (B) of the proinflammatory cytokines production were performed. (C) Dose-dependent inhibitory effects of triptolide on IL-6 production by DC in response to LPS stimulation. DC pretreated with various concentrations of triptolide as indicated for 12 h, and then stimulated with LPS (1 g/ml) for 24 h, and IL-6 production measured by ELISA. Results are representative of three independent experiments, and the data presented as mean ± S.D. * P < 0.05, ** P < 0.01.
of triptolide on the functions of DC, the key player of immune initiation and regulation, have not been clearly defined. DC migration, one critical step for mature DC-initiated immune response, is orchestrated by a complex interplay between chemokines and their receptors. During the differentiation and maturation of DC, sequential and orchestrated expressions of chemokine receptors are responsible for the proper migration of DC to immune sites (Sallusto et al., 1998). Upon DC maturation, CCR1, CCR2, and CCR5 that normally direct DC infiltrations into peripheral tissues are downregulated, accompanied by the upregulation of CCR7 that are responsive to chemokines (such as ELC/CCL19/MIP-3 or SLC/CCL21/6Ckine) expressed by secondary lymphoid organs, both of which work together to
direct the emigration of DC out of tissues and the immigration of DC into LN or spleen. The chemokine receptor CCR7 and its ligands CCL19/MIP-3 and CCL21/6Ckine control myeloid DC migration into afferent lymphatic vessels and the localization of these DC within LN (Sallusto et al., 1998; MartIn-Fontecha et al., 2003; Ohl et al., 2004). Our data show that triptolide can inhibit LPS-induced upregulation of CCR7 expression in DC, and accordingly, triptolide-treated DC exhibit the impaired migration to CCL19/MIP-3 in vitro and migration to secondary lymphoid organs in vivo. Recently, Chen et al. also reported that triptolide inhibits the chemotactic response of LPS-stimulated human monocyte-derived DC (MoDC) to CCL21/6Ckine by down-regulating CCR7 expression (Chen et al., 2005). It has
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been reported that the maturation-induced up-regulation of CCR7 expression on human monocyte-derived DC (MoDC) was insufficient to allow MoDC migration to CCL19/MIP-3 and CCL21/6Ckine. MoDC require a PGE2 -provided signals to acquire the potential to migrate effectively in response to LN-derived chemokines CCL19/MIP-3 and CCL21/6Ckine (Scandella et al., 2004; Liu et al., 2006; Randolph et al., 2005). So, we went further to investigate whether triptolide inhibits LPS-induced upregulation of COX-2 expression in DC. In agreement with previous reports showing triptolide inhibits COX expression in several other cell types (Tao et al., 1998; Yao et al., 2005), we demonstrate that triptolide inhibits PGE2 production by decreasing COX-2 gene expression in DC. It has been reported recently that stimulation of immature MoDC with PGE2 alone, or in association with CD40 ligand, TNF-␣, LPS or IL-1, can induce MoDC surface expression of CCR7 and acquisition of migratory responsiveness to CCL21/SLC (Luft et al., 2002; Ardeshna et al., 2000; Lee et al., 2003). This is consistent with our results that exogenous PGE2 can restore CCR7 expression and migratory capacity of triptolide treated and LPSstimulated-DC. Taken together, these data further confirm our conclusion that triptolide impairs the migratory capacity of DC, at least partially, by inhibiting the ability of DC to produce PGE2 in response to LPS. MAPK, PI3-K/Akt signal pathways and NF-B transcription regulate COX-2 and CCR7 expression in different cell lines. We also show that triptolide can inhibit LPS-activated NF-B and PI3-K/Akt pathways but not the MAPK pathway in DC, and the inhibited PI3-K/Akt and NF-B pathways may contribute to the decreased COX-2 and CCR7 expression in DC by triptolide. In addition, the inhibitory effect by triptolide on DC migration is not due to DC apoptosis since we did not detect either apoptosis, mitochondrial membrane potential decrease, or caspase 3 activation upon DC exposure to low concentrations of triptolide (1–5 ng/ml) (data not shown). In summary, our results demonstrate that triptolide potently inhibits LPS-induced CCR7 and COX-2 expression in DC leading to the impaired DC migration to secondary lymphoid organs both in vitro and in vivo. The impaired DC migration by triptolide may be due to the reduced COX-2 and CCR7 expression via the inhibition of PI3-K/Akt kinase activity and nuclear NF-B activation in DC. Therefore, our results have provided important
Fig. 6. Triptolide inhibits LPS-induced PI3-K/Akt kinase activity and nuclear activation of NF-B in DC. (A) Triptolide inhibits PI3-K and Akt/PKB kinase activity. DC were pretreated with triptolide for 12 h, or pretreated with PI3-K/Akt inhibitor for 30 min, and then stimulated with LPS for 15 min (PI3-K) or for indicated time (Akt kinase), cell lysates were prepared, PI3-K (upper panel) and Akt kinase (lower panels) activity was assayed respectively as
described in Section 2. Similar results were observed in three separate experiments. * P < 0.05, ** P < 0.01. (B) Triptolide inhibits LPS-induced NF-B nuclear activation. Nuclear extracts were prepared from triptolide-stimulated or LPSstimulated DC as indicated and probed for NF-B, nucleoporin p62 as a nuclear protein loading control. (C) Exogenous PGE2 rescues triptolide-mediated inhibition of Akt kinase activity and nuclear NF-B activation in LPS-stimulated DC. DC were pretreated with triptolide for 12 h, and then stimulated with LPS or LPS and PGE2 together for indicated time, Akt kinase activity and nuclear NF-B activation was assayed as described in Section 2. (D) The effect of combined use of signal inhibitor and triptolide on DC migration. DC were pretreated with inhibitor of PI3-K/Akt, NF-B or COX-2 respectively for 30 min, and then treated with or not triptolide for 24 h, then in vitro chemoattraction assay of DC toward MIP-3 was detected as described in Section 2. * P < 0.05, ** P < 0.01 (* compared with LPS group). P > 0.05 ( compared with triptolide treatment group).
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mechanistic insights on triptolide-medicated immunosuppression. Acknowledgments We sincerely appreciate Ms Rui Zhang, Ms Dandan Zhang and Ms Weiqin Ni for their excellent technical assistance. This work was supported by grants from the National Natural Science Foundation of China (30418017, 30490240, 30128022), and the National Key Basic Research Program of China (2003CB515503, 2001CB510002). References An, H., Yu, Y., Zhang, M., Xu, H., Qi, R., Yan, X., Liu, S., Wang, W., Guo, Z., Guo, J., Qin, Z., Cao, X., 2002. Involvement of ERK, p38 and NF-kappaB signal transduction in regulation of TLR2, TLR4 and TLR9 gene expression induced by lipopolysaccharide in mouse dendritic cells. Immunology 106, 38–45. Ardeshna, K.M., Pizzey, A.R., Devereux, S., Khwaja, A., 2000. The PI3 kinase, p38 SAP kinase, and NF-kappaB signal transduction pathways are involved in the survival and maturation of lipopolysaccharide-stimulated human monocyte-derived dendritic cells. Blood 96, 1039–1046. Banchereau, J., Steinman, R.M., 1998. Dendritic cells and the control of immunity. Nature 392, 245–252. Banchereau, J., Briere, F., Caux, C., Davoust, J., Lebecque, S., Liu, Y.J., Pulendran, B., Palucka, K., 2000. Immunobiology of dendritic cells. Annu. Rev. Immunol. 18, 767–811. Beckmann, J.H., Yan, S., Luhrs, H., Heid, B., Skubich, S., Forster, R., Hoffmann, M.W., 2004. Prolongation of allograft survival in ccr7-deficient mice. Transplantation 77, 1809–1814. Chang, M.S., Lee, W.S., Chen, B.C., Sheu, J.R., Lin, C.H., 2004. YC-1-induced cyclooxygenase-2 expression is mediated by cGMP-dependent activations of Ras, phosphoinositide-3-OH-kinase, Akt, and nuclear factor-kappaB in human pulmonary epithelial cells. Mol. Pharmacol. 66, 561–571. Chen, B.J., Liu, C., Cui, X., Fidler, J.M., Chao, N.J., 2000. Prevention of graft-versus-host disease by a novel immunosuppressant, PG490-88, through inhibition of alloreactive T cell expansion. Transplantation 70, 1442–1447. Chen, B.J., Chen, Y., Cui, X., Fidler, J.M., Chao, N.J., 2002. Mechanisms of tolerance induced by PG490-88 in a bone marrow transplantation model. Transplantation 73, 115–121. Chen, T., Guo, J., Yang, M., Han, C., Zhang, M., Chen, W., Liu, Q., Wang, J., Cao, X., 2004. Cyclosporin A impairs dendritic cell migration by regulating chemokine receptor expression and inhibiting cyclooxygenase-2 expression. Blood 103, 413–421. Chen, X., Murakami, T., Oppenheim, J.J., Howard, O.M., 2005. Triptolide, a constituent of immunosuppressive Chinese herbal medicine, is a potent suppressor of dendritic-cell maturation and trafficking. Blood 106, 2409–2416. Dubois, R.N., Abramson, S.B., Crofford, L., Gupta, R.A., Simon, L.S., Van, D., Lipsky, P.E., 1998. Cyclooxygenase in biology and disease. FASEB J. 12, 1063–1073. Fidler, J.M., Ku, G.Y., Piazza, D., Xu, R., Jin, R., Chen, Z., 2002. Immunosuppressive activity of the Chinese medicinal plant Tripterygium wilfordii. III. Suppression of graft-versus-host disease in murine allogeneic bone marrow transplantation by the PG27 extract. Transplantation 74, 445–457. Hackstein, H., Thomson, A.W., 2004. Dendritic cells: emerging pharmacological targets of immunosuppressive drugs. Nat. Rev. Immunol. 4, 24–34. Harizi, H., Juzan, M., Pitard, V., Moreau, J.F., Gualde, N., 2002. Cyclooxygenase-2-issued prostaglandin E2 enhances the production of endogenous IL-10, which down-regulates dendritic cell functions. J. Immunol. 168, 2255–2263. Kabashima, K., Sakata, D., Nagamachi, M., Miyachi, Y., Inaba, K., Narumiya, S., 2003. Prostaglandin E2-EP4 signaling initiates skin immune responses by promoting migration and maturation of Langerhans cells. Nat. Med. 9, 744–749.
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Triptolide impairs dendritic cell migration by inhibiting CCR7 and COX-2 expression through PI3-K/Akt and NF-B pathways Qiuyan Liu a,b , Taoyong Chen a , Guoyou Chen a , Xiaoli Shu b , Anna Sun b , Pengcheng Ma c , Liwei Lu d , Xuetao Cao a,b,∗ a
Institute of Immunology and National Key Laboratory of Medical Immunology, Second Military Medical University, Shanghai 200433, PR China b Institute of Immunology, Zhejiang University, Hangzhou 310031, PR China c Institute of Dermatology, Chinese Academy of Medical Sciences, Nanjing 210042, PR China d Department of Pathology, The University of Hong Kong, Hong Kong, PR China Received 20 October 2006; accepted 6 December 2006 Available online 16 January 2007
Abstract Inhibition of dendritic cell (DC) migration into tissues and secondary lymphoid organs is an efficient way to induce immunosuppression and tolerance. CCR7 and PGE2 are critical for DC migration to secondary lymphoid organs where DC initiate immune response. Triptolide, an active component purified from the medicinal plant Tripterygium Wilfordii Hook F., is a potent immunosuppressive drug capable of prolonging allograft survival in organ transplantation by inhibiting T cell activation and proliferation. Considering the essential role in T cell tolerance of DC migration to secondary lymphoid organs, here we demonstrate that triptolide can significantly inhibit LPS-triggered upregulation of CCR7 expression and PGE2 production by inhibiting cyclooxygenase-2 (COX-2) expression in DC, thus impairing DC migration towards CCR7 ligand CCL19/MIP-3 in vitro. Moreover, triptolide-treated DC display impaired migration into secondary lymphoid organs and in vivo administration of triptolide also inhibits DC migration. Further studies show that the triptolide-mediated inhibitory effects of LPS-induced activation of phosphatidylinositol-3 kinase (PI3-K)/Akt and nuclear NF-B activation are involved in down-regulation of COX-2 and CCR7 expression resulting in impaired migration to secondary lymphoid organs of DC. Therefore, inhibition of DC migration through decreasing COX-2 and CCR7 expression via PI3-K/Akt and NF-B signal pathways provides additional mechanistic explanation for triptolide’s immunosuppressive effect. © 2006 Elsevier Ltd. All rights reserved. Keywords: Dendritic cell; Migration; CCR7; Immunosuppression; Signal transduction
1. Introduction Triptolide, a diterpene triepoxide, is an active component of extracts derived from the medicinal plant Tripterygium Wilfordii Hook F. (TWHF), and has anti-inflammatory and immunosuppressive activities (Qiu and Kao, 2003). Tripterygium extracts have, for many years, been used widely to treat autoimmune dis-
Abbreviations: trip, triptolide; DC-trip1, DC were treated with triptolide 1 ng/ml; DC-trip1/LPS, DC were treated with triptolide 1 ng/ml and stimulated with LPS; DC-trip5, DC were treated with triptolide 5 ng/ml; DC-trip5/LPS, DC were treated with triptolide 5 ng/ml and stimulated with LPS ∗ Corresponding author at: Institute of Immunology and National Key Laboratory of Medical Immunology, Second Military Medical University, 800 Xiangyin Road, Shanghai 200433, PR China. Tel.: +86 21 55620605; fax: +86 21 65382502. E-mail address: [email protected] (X. Cao). 0161-5890/$ – see front matter © 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.molimm.2006.12.003
eases including rheumatoid arthritis, immune complex nephritis, and systemic lupus erythematosus in China (Ramgolam et al., 2000; Tao and Lipsky, 2000). Clinical and experimental studies have also demonstrated that Tripterygium extracts effectively prolong allograft survival in organ transplantation including bone marrow, cardiac, renal and skin transplantation (Chen et al., 2002; Fidler et al., 2002; Wang et al., 2000; Yang et al., 1992). The immunosuppressive effects by triptolide can be partly attributed to its potent inhibitory effects on T cell activation and interleukin-2 (IL-2) production (Chen et al., 2000; Qiu et al., 1999). Dendritic cells (DC), the most potent professional antigenpresenting cells (APC), are critical for the induction of both immune responses and immune tolerance (Banchereau and Steinman, 1998; Banchereau et al., 2000; Hackstein and Thomson, 2004; Sallusto and Lanzavecchia, 1999). DC originate from bone marrow precursors and migrate through the blood
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stream to almost every tissue, where they reside in an immature state. At the site of infection, DC recognize and respond to common pathogen-associated molecular patterns (PAMPs), including lipopeptides, lipopolysaccharides (LPS) and nucleic acids via Toll-like receptors (TLR) (Ricciardi-Castagnoli and Granucci, 2002). LPS activates DC and induces production of proinflammatory cytokines and other mediators via TLR4, thus triggering a complex phenotypic and functional maturation program of DC including up-regulation of the chemokine receptor CCR7 (Sallusto et al., 1998). The CCR7 upregulation enable DC more responsiveness to CCR7 ligands, which promotes DC migration from peripheral tissues to secondary lymphoid organs such as lymph nodes (LN) and spleen where DC present antigens to T cells and initiate immune response (MartIn-Fontecha et al., 2003; Randolph, 2001). The expression of CCR7 on mature DC is essential for their migration to the T-cell area of draining LN because this migration is guided by two ligands for CCR7, CCL19 (EBI1-ligand chemokine [ELC], macrophage inflammatory protein [MIP]-3), and CCL21 (secondary lymphoid-tissue chemokine [SLC], 6-Ckine). Both chemokines are expressed by stromal cells in the T-cell area of secondary lymphoid organs (Ohl et al., 2004). The essential role of CCR7 and its ligands in mature DC migration to LN was demonstrated in CCR7deficient mice (Beckmann et al., 2004) and plt/plt mice, which lack the ligands for CCR7 (Luther et al., 2000). Because the migration of DC into secondary lymphoid organs is required for the initiation of specific immune responses, so, inhibition of DC migration into secondary lymphoid organs by inhibiting CCR7 expression is an efficient way to induce immunosuppression and tolerance. PGE2 , derived from metabolism of free arachidonic acid, exerts an important immunomodulatory role in DC differentiation and function. Previous studies have shown that PGE2 is required for CCR7-expressed DC migration to its ligands CCL19 and CCL21 (Kabashima et al., 2003; Scandella et al., 2002; Scandella et al., 2004), the importance of PGE2 for DC migration in vivo has recently been shown in Ptger4−/− mice lacking the PGE2 receptor EP4. These mice display impaired migration of Langerhans cells and reduced skin immune responses (Legler et al., 2006). Besides its inhibitory effects on T cell function, triptolide may also affect other cells involved in immune responses. We and other researchers have shown that triptolide can affect DC functional maturation at lower or clinical concentrations (Chen et al., 2005; Liu et al., 2006; Zhu et al., 2005) and induce DC apoptosis at high concentrations (≥10 ng/ml) (Liu et al., 2004). Considering that the migration of DC into secondary lymphoid organs is required for the initiation of immune responses and CCR7 on DC is a potential target for immunosuppressive drug, in this study, we investigated the effect of triptolide, used at clinical concentration that does not induce DC apoptosis, on DC migration in response to LPS stimulation. We demonstrate that triptolide can significantly inhibit LPS-triggered upregulation of CCR7 expression and PGE2 production by inhibiting cyclooxygenase-2 (COX-2) expression in DC, thus impairing DC migration towards CCR7 ligand CCL19/MIP-3 in vitro, and in vivo administration of triptolide also inhibits DC migration to the spleen and lymphoid
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nodes. Furthermore, triptolide-mediated inhibitory effects of LPS-induced activation of PI3-K/Akt and nuclear NF-B activation are involved in down-regulation of COX-2 and CCR7 expression resulting in impaired migration to secondary lymphoid organs of DC. Therefore, inhibition of DC migration through decreasing COX-2 and CCR7 expression via PI3-K/Akt and NF-B signal pathways provides additional mechanistic explanation for triptolide’s immunosuppressive effect. 2. Materials and methods 2.1. Mice and reagents Male wild-type C57BL/6 mice, 5–6 weeks of age, were purchased from SIPPR-BK Experimental Animal Co. (Shanghai, China). GFP+/+ C57BL/6-TgN (ACTbEGFP) mice were obtained from the Jackson Laboratory (Bar Harbor, ME). All mice were housed in a pathogen-free facility for all experiments. Crystalline triptolide (PG490, molecular weight 360, purity of 99%) was obtained from the Institute of Dermatology, Chinese Academy of Medical Sciences (Nanjing, PR China) and prepared as previously described (Mao et al., 1998). Triptolide was reconstituted in dimethyl sulphoxide (DMSO) stock solutions (1 mg/ml) and stored at −20 ◦ C. Triptolide was freshly diluted to the indicated concentrations with culture medium before use. The DMSO concentration in all test conditions never exceed 0.002% (V/V). Recombinant mouse GM-CSF and IL-4 were purchased from R&D Systems (Minneapolis, MN). Chemokine receptor antibodies (CCR1, CCR5, CCR7 and CXCR4), polyclonal Abs against actin, nucleoporin p62, monoclonal Ab against COX-2 and monoclonal Ab for NF-B p65 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). PE-labeled CCR7 Ab was purchased from ebioscience (San Diego, CA). Phospho-Abs against extracellular signal-regulated kinase p44/p42 (ERK1/2, Thr202/Tyr204), c-Jun N-terminal kinase/stress associated protein kinase (JNK/SAPK, Thr183/Tyr185), p38 (Thr180/Tyr182), and corresponding Abs against non-phosphorylated signaling proteins were from Cell Signaling (Beverly, MA). ELISA kits for murine PGE2 , TNF-␣, IL-1 and IL-6 and recombinant murine MIP-1␣ and MIP-3 were purchased from R&D Systems (Minneapolis, MN). Lipopolysaccharide (LPS), PGE2 and PI3-K/Akt inhibitor Wortmannin and COX-2 specific inhibitor NS398 were from Calbiochem (San Diego, CA). 2.2. Generation of mouse bone marrow (BM)-derived DC Mouse bone marrow-derived DC were prepared from bone marrow by culturing with 10 ng/ml recombinant GM-CSF and 1 ng/ml IL-4, as described previously (Chen et al., 2004). Following 6 days of culture, DC were harvested and purified using anti-CD11c-coated microbeads (Miltenyi-Biotec, Auburn, CA), then a highly enriched population of DC (90–95% CD11c+ cells) was obtained. DC were pretreated with triptolide for 12 h, and then LPS or LPS and PGE2 treatments were performed as indicated. In some experiments, DC were pretreated with signal inhibitors for 30 min before LPS or LPS and PGE2 stimulation.
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2.3. ELISA assay Cytokines (TNF-␣, IL-1, IL-6) and PGE2 contained in the culture supernatants of DC treated with or without triptolide after LPS stimulated or not were measured by ELISA (R&D, Minneapolis, MN) according to the manufacturer’s instructions. 2.4. Assay for chemokine receptor expression For RT-PCR assay of chemokine receptor mRNA expression, total cellular RNA was isolated from DC following indicated stimulation with TRIzol reagent according to the manufacturer’s instructions (Life Technology). cDNA was synthesized from 1 g of total RNA by extension of oligo(dT)18 primer with 200 units AMV (Promega, Madison, WI). Primers used for chemokine receptors were as follows: 5 GTG TTC ATC ATT GGA GTG GTG 3 and 5 GGT TGA ACA GGT AGA TGC TGG TC 3 for CCR1; 5 CAT CGA TTA TGG TAT GTC AGC ACC 3 and 5 CAG AAT GGT AGT GTG AGC AGG AA 3 for CCR5; 5 CCA GGA AAA ACG TGC TGG TG 3 and 5 GGC CAG GTT GAG CAG GTA GG 3 for CCR7; 5 CGG CAA TGG ATT GGT GAT CCT GGT C 3 and 5 GAG GGC CTT GCG CTT CTG GTG GCC C 3 for CXCR4. Primers used for mouse -actin were 5 GTG GGA ATT CGT CAG AAG GAC TCC TAT GTG 3 and 5 GAA GTC TAG AGC AAC ATA GCA CAG CTT CTC 3 . Equal amounts of total cellular RNA were used for reverse transcription under the same conditions. cDNA levels were semi-quantified by inclusion of a -actin control. PCR was performed at 94 ◦ C 30 s, 55 ◦ C 30 s and 72 ◦ C 30 s for 25 cycles, followed by 10 min elongation at 72 ◦ C. PCR products were separated on a 1% agarose gel and analyzed using a Gel Doc 2000 system (Bio-Rad, Hercules, CA). Western blot was performed to analyze chemokine receptor expression in DC at the protein level as described previously (Chen et al., 2004). For FACS assays, DC were labeled with chemokine receptor primary Ab in PBS for 30 min, and then washed, stained with FITC-labeled secondary Ab, and then analyzed by FACS as described previously (Chen et al., 2004). 2.5. In vitro DC chemoattraction assay In vitro chemoattraction of DC in response to macrophage inflammatory protein (MIP)-1␣ or MIP-3 was measured as previously described (Chen et al., 2004). 2 × 105 DC were added to the upper chamber of 24-well transwell chambers with 5 mpore size polycarbonate filters (Corning Costar, Cambridge, MA) in a total volume of 100 l. The chambers were incubated for 4 h at 37 ◦ C. Cells migrating to the bottom chamber were labeled with FITC-CD11c (DC) Ab for 30 min at 4 ◦ C and then counted by flow cytometry. Each experiment was performed in triplicate. 2.6. In vivo DC migration assays To determine whether the migration of triptolide-treated DC to secondary lymphoid organs was impaired, we transplanted BALB/c skin sheets (10–15 mm pieces) to C57BL/6 mice.
Two days after transplantation, in vitro-cultured C57BL/6 DC, with or without triptolide treatment (5 ng/ml for 24 h), were labeled with the red fluorescence marker PKH-26 (Sigma) as described (Chen et al., 2004), and immediately injected into skin allografts (1 × 106 in 100 l). Six hours after injection, mice were killed and ipsilateral inguinal lymph nodes and spleens harvested, embedded in optimal cutting temperature (OCT) compound (IEC, Needham Heights, MA), and frozen immediately at −80 ◦ C. Cryostat sections (6 m) were placed on cover slides, fixed in 1% paraformaldehyde for 5 min, incubated with 0.4 g/ml Hoechst (Calbiochem) for 5 min, and subsequently examined by fluorescence microscopy. Cell suspensions from lymph nodes and spleen were also prepared and PKH26-positive cells contained in these secondary lymphoid organs counted by FACS. To exclude the influence of in vitro labeling on DC function, green DC derived from GFP+/+ (green fluorescent protein) C57BL/6 mice were also used instead of PKH-26-labeled DC. To mimic normal/physiological in vivo DC migration as accurately as possible, C57BL/6 mice were intraperitoneally injected with triptolide (5 g/kg) every day for a total of 7 days, and FITC was painted onto the abdomen as described previously (Macatonia et al., 1987), then the mice were intravenously injected with LPS (5 mg/kg) via the tail vein 12 h after the final injection. Spleen and draining lymph nodes were isolated, and cell suspensions were prepared. To exclude the influence of DC apoptosis induced by triptolide in vivo, DC in the cell suspensions were stained with PE-labeled CD11c and 7-AAD, and FITC/PE double-positive and 7-AAD negative cells detected by FACS. Results are representative of n = 4 mice in each group. 2.7. Assay of PI3-K activity PI3-K activity was assayed with PI3-K ELISA kit (Echelon Biosciences Inc., Salt Lake City, UT) according to manufacturer’s instructions. In brief, DC were pretreated with triptolide (5 ng/ml) for 12 h or pretreated with PI3-K/Akt inhibitors for 30 min and then stimulated with LPS for 15 min. Cell lysates were prepared, PI3-K was immunoprecipitated with antibody against the p85 subunit and incubated with PI(4,5)P2. The reaction products were incubated with a PI(3,4,5)P3 detector protein, then added to PI(3,4,5)P3-coated microplate for competitive binding. A peroxidase-linked secondary detection reagent and colorimetric detection was used to detect PI (3,4,5)P3 detector protein binding to the plate. The colorimetric signal was inversely proportional to the amount of PI(3,4,5)P3 produced by PI3-K activity. 2.8. Assay of Akt kinase activity Akt activity was detected using Akt kinase Assay kit (Cell Signaling Technology Inc., Beverly, MA) according to manufacturer’s instructions. Briefly, DC were treated as above, cell lysates were prepared and immobilized Akt primary antibody was used to immunoprecipitate Akt from the extracts. GSK-3 fusion protein (1 g) was incubated with the precipitated protein in kinase buffer and then detected with Phospho-GST-3 specific
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antibody by Western blot. The phosphorylation level of GSK-3 fusion protein was proportional to Akt activity. 2.9. Immunoprecipitation and Western blotting DC were lysed with M-PERTM Protein Extraction Reagent (Pierce, Rockford, IL) supplemented with protease inhibitor cocktail, and protein concentrations of the extracts measured by BCA assay (Pierce, Rockford, IL). Forty micrograms of the protein was used either for immunoprecipitation, or loaded per lane, subjected to SDS-PAGE, transferred onto nitrocellulose membranes, then blotted as described previously (An et al., 2002). For detection of the nuclear translocation of NF-B p65, cytoplasmic and nuclear extracts were prepared using NE-PER nuclear and cytoplasmic extraction reagents (Pierce, Rockford, IL) and protein concentration was determined with a BCA-200 protein assay kit. Nuclear NF-B p65 subunit was detected by Western blot using specific antibody. 2.10. Statistical analysis Comparisons between experimental groups and relevant controls were performed by Student’s t-test. P < 0.05 was considered a statistically significant difference. 3. Results 3.1. Triptolide inhibits CCL19/MIP-3β-induced DC migration both in vitro and in vivo One characteristic of mature DC is their ability to migrate to secondary lymphoid organs where DC present antigens to T cells and initiate immune response. So, inhibition of DC migration to secondary lymphoid organs is an efficient way to induce immunosuppression and tolerance. To confirm whether triptolide affects DC migration, we examined in vitro migration of DC toward MIP-1␣ and MIP-3, two chemokines chemotactic for immature and mature DC respectively. As expected, immature DC efficiently responded to MIP-1␣ while LPS-matured DC responded well to MIP-3 (Fig. 1A and B). In contrast, after pretreatment with triptolide, LPS-matured DC responded poorly to MIP-3 (Fig. 1B). Interestingly, these mature DC retained their capability to respond to MIP-1␣ (Fig. 1A). To further investigate the possibility that triptolide treatment impaired either the capacity of DC to emigrate out of skin epidermis or to immigrate into secondary lymphoid organs, we performed in vivo experiments to examine the migratory capacity of DC in mice treated with triptolide. DC were treated with triptolide, and then labeled with PKH-26, and subcutaneously injected into skin transplants of C57BL/6 mice derived from BALB/c mice (1 × 106 DC per mouse), they showed dramatically reduced immigration into the spleen and lymph nodes as compared with untreated control DC (Fig. 1C). PKH-26-labeled cell numbers in spleen and lymph nodes were also enumerated by FACS; PKH-26-positive cells in secondary lymphoid organs were reduced in the triptolide treatment group (Fig. 1D). When GFP-expressing DC derived from C57BL/6GFP+/+ mice
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were used instead of PKH-26-labeled DC, similar results were observed (Fig. 1E). In the FITC-painting assay, following LPS mobilization, FITC-positive DC numbers in lymph nodes and spleen of triptolide-treated mice were significantly reduced than that in normal mice (Fig. 1F). Our study also demonstrated that triptolide-pretreated DC exhibited a reduced allostimulatory activity (data not shown). Taken together, these results demonstrate that administration of triptolide not only impairs the migration of DC into secondary lymphoid organs, but also reduces allostimulatory activity of DC. 3.2. Triptolide inhibits LPS-induced CCR7 expression in DC It’s well known that LPS induces maturation of DC with upregulation of CCR7 which binds to its ligand CCL19/MIP3 and directs DC migration to secondary lymph organs, this process is critical for DC to initiate immune response. To confirm the reduced migration of DC by triptolide to secondary lymphoid organs is associated with the inhibited CCR7 expression, we first investigated the chemokine receptor expression in triptolide-treated DC with or without LPS stimulation. DC were pretreated with triptolide for 12 h and then stimulated with LPS (1 g/ml) for 12 h for RT-PCR and FACS analysis or 24 h for Western blot assay. We found that triptolide pretreatment decreased the LPS-induced upregulation of CCR7 and CXCR4 in DC both at the mRNA (Fig. 2A) and protein level (Fig. 2B and C). In contrast, expression of CCR1 and CCR5 in DC was not decreased by triptolide. These results demonstrate that triptolide can significantly inhibit LPS-induced CCR7 expression in DC. Furthermore, the impaired LPS-induced upregulation of CCR7 expression in triptolide-treated DC may account for their reduced responsiveness to MIP-3 while the constitutive expression of CCR1 and CCR5 in triptolide-treated DC may be responsible for their continued responsiveness to MIP-1␣. 3.3. Triptolide inhibits COX-2 expression and PGE2 production in DC PGE2 , derived from metabolism of free arachidonic acid, exerts an important immunomodulatory role in DC differentiation and function (Kabashima et al., 2003; Scandella et al., 2002, 2004). Cyclooxygenases (COX-1 and COX-2) are the major enzymes involved in PGE2 production (Dubois et al., 1998; Smith et al., 2000). COX-1 is constitutively expressed in most tissues and appears to mediate various physiological functions (Smith et al., 1996; Smith and DeWitt, 1995). By contrast, COX-2 is undetectable in most normal tissues, but is rapidly induced in response to inflammatory stimuli, including TNF␣, IL-1␣ and LPS (Harizi et al., 2002; Sharma et al., 2003). It was shown that both extracts of TWHF and triptolide could inhibit PGE2 production in a variety of human cells by blocking the upregulation of COX-2 (Tao et al., 1998; Yao et al., 2005). So we wondered whether triptolide impairs DC migration by inhibiting LPS-induced COX-2 expression and resultant PGE2 production in DC. As shown in Fig. 3A, triptolide treatment of DC resulted in a decrease of LPS-stimulated COX-2 expres-
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Fig. 1. Effect of triptolide on DC migration in vitro and in vivo. (A and B) Triptolide causes LPS-stimulated DC to retain responsiveness to MIP-1␣ (A) but inhibits responsiveness of LPS-stimulated DC to MIP-3 (B). DC were cultured and treated as described in Section 2, then evaluated for responsiveness to MIP-1␣ and MIP-3 by in vitro chemoattraction assay. 2 × 105 triptolide-treated DC were resuspended in 0.1 ml RPMI 1640 containing 0.5% BSA, and loaded into the upper well of a transwell chamber. Lower compartment were filled with RPMI 1640 containing 0.5% BSA, with or without MIP-1␣ (100 ng/ml) or MIP-3 (100 ng/ml), respectively. The chambers were incubated for 4 h at 37 ◦ C. Directed migration was expressed as the number of CD11c+ cells that had migrated to the lower chamber, counted by FACS analysis. Each experiment was performed in triplicate at least three times. Data is expressed as mean number of migrated CD11c+ cells ±S.D. * P < 0.05, ** P < 0.01. (C and D) Triptolide treatment impairs PKH-26-labeled murine DC migration into spleen and lymph nodes (LN). (C) Immunofluorescence microscopy of PKH-26-positive cells in spleen and LN. Results here are representative of sections derived from at least three samples. Red cells are PKH-26 labeled DC, and blue cell nucleus is stained with Hoechst. Original magnification, 400×. (D) FACS enumeration of PKH-26-positive cells in spleen and LN. (E) FACS analysis of DC GFP+ cells (R2 region) in spleen and LN following injection into skin transplants. Labels indicate percentage of GFP+ cells in cell suspensions. (F) Triptolide decreases the frequency of FITC+ CD11c+ DC in skin migration to spleen and LN. C57BL/6 mice were treated with triptolide (5 g/kg) for 7 days as described in Section 2, and then FITC was painted onto the abdomen, the mice were intravenously injected with LPS (5 mg/kg) via the tail vein 12 h after the final injection. Spleen and draining LN were isolated, and cell suspensions were prepared. DC in the cell suspensions were stained with PE-labeled CD11c and 7-AAD, and FITC/PE double-positive and 7-AAD negative cells detected by FACS. Results represent three independent experiments. * P < 0.05, ** P < 0.01.
sion at both the mRNA and protein level. Importantly, PGE2 levels in the culture supernatant were also reduced significantly (Fig. 3B), suggesting that triptolide inhibits PGE2 production by decreasing COX-2 expression in DC. 3.4. Exogenous PGE2 overcomes the inhibited DC migration and reverses CCR7 expression We and other researchers have shown that PGE2 promotes DC migration in response to chemotactic factors by modulating
chemokine receptor expression (Scandella et al., 2002; Chen et al., 2004; Luft et al., 2002). To further investigate the role of the reduced PGE2 in triptolide-mediated inhibition of DC migration, we added exogenous PGE2 into DC migration assay, and found addition of exogenous PGE2 (1 M) restored the migratory capacity of triptolide-treated DC in chemoattraction assays (Fig. 4A). We then examined whether PGE2 also restored CCR7 expression in triptolide-treated and LPS-stimulated DC. In the presence of exogenous PGE2 , LPS-stimulated DC expression of CCR7 was also restored (Fig. 4B and C). These results are con-
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Fig. 3. Triptolide inhibits LPS-induced COX-2 expression and PGE2 production in DC. (A) Triptolide inhibits LPS-induced COX-2 expression. DC were treated as indicated and COX-2 expression assayed by RT-PCR (upper two panels) and Western blot assays (lower two panels). (B) Supernatants from triptolide-treated or -untreated DC, stimulated or unstimulated with LPS (1 g/ml) for 24 h, were examined for PGE2 production by ParameterTM PGE2 Assay.
leading to the impaired migration of DC to secondary lymphoid organs.
Fig. 2. Effect of triptolide on LPS-induced chemokine receptor expression in DC. (A) RT-PCR and (B) Western blot assay of chemokine receptor expression in DC pretreated with triptolide for 12 h or then stimulated with 1 g/ml LPS for 12 h or 24 h, respectively. -Actin (RT-PCR) and actin (Western blot) were used as a quantitative control. (C) DC pretreated with triptolide for 12 h and then stimulated with 1 g/ml LPS for 12 h, then labeled with Abs against chemokine receptors in PBS for 30 min, washed, and finally stained with FITC-labeled secondary Ab and analyzed by FACS. Unshaded regions represent isotype controls and shaded regions corresponding Abs, as indicated. Results were expressed as fluorescence intensity (FI). Data were expressed as mean (n = 3). There are significant differences in expression of CCR1, CCR5, CCR7 and CXCR4 between triptolide-treated DC with or without LPS stimulation. * P < 0.05, ** P < 0.01.
sistent with reports showing that PGE2 regulates the expression of CCR7 on DC and their migration. These results suggest that the inhibition of DC migration by triptolide may be partially due to its inhibition of COX-2 expression, which in turn results in a defective production of PGE2 and CCR7 expression in DC, thus
3.5. Triptolide impairs LPS-induced proinflammatory cytokines production in DC It is well known that COX-2 is undetectable in most normal tissues but is rapidly induced in response to proinflammatory cytokines (Harizi et al., 2002; Sharma et al., 2003), LPS activates DC and triggers production of proinflammatory cytokines including TNF-␣, IL-1 and IL-6. To investigate whether triptolide inhibits LPS-induced proinflammatory cytokines production, 5 × 105 cells/ml DC were pretreated with triptolide (5 ng/ml) for 12 h, which were then stimulated with or without LPS (1 g/ml) for indicated time. RT-PCR and ELISA results showed that triptolide inhibited both mRNA (Fig. 5A) and protein (Fig. 5B) expression of proinflammatory cytokines including TNF-␣, IL-1 and IL-6 by DC. Moreover, triptolide inhibited proinflammatory cytokines production by DC in a dose-dependent manner, even at very low concentrations (1 ng/ml) (Fig. 5C).
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3.6. Inhibition of PI3-K/Akt kinase activity and NF-κB nuclear activation contributes to the impaired COX-2 and CCR7 expression in DC by triptolide
Fig. 4. Exogenous PGE2 rescues the triptolide-induced migratory impairment of DC and restores CCR7 expression. (A) Exogenous PGE2 rescues migration of triptolide-treated DC to CCL19/MIP-3. DC were pretreated with triptolide for 12 h, and then stimulated with LPS or LPS and PGE2 together for 24 h, evaluated for responsiveness to CCL19/MIP-3 by in vitro chemoattraction assay as above described. * P < 0.05, ** P < 0.01. (B and C) Exogenous PGE2 restores CCR7 expression. (B) RT-PCR (upper panels) and Western blot (lower panels) and (C) FACS analysis assay of CCR7 expression in DC following supplementation with PGE2 (1 M).
It was reported that MAPK, PI3-K/Akt signal pathways and NF-B transcription regulate COX-2 and CCR7 expression in different cell lines (Ardeshna et al., 2000; Chang et al., 2004; Lee et al., 2003; St-Germain et al., 2004). We examined whether these signaling pathways are involved in the triptolide-mediated inhibitory effect on COX-2 and CCR7 expression in DC. Upon LPS stimulation, the ERK, JNK, and p38 pathways in DC were rapidly activated, and triptolide treatment did not significantly affect the activation of these pathways in DC (data not show), suggesting that triptolide does not interfere with the MAPK pathway in LPS-treated DC. We then examined whether PI3-K/Akt signaling pathway is involved in triptolide-mediated inhibition of COX-2 and CCR7 expression in DC. As shown in Fig. 6A, triptolide treatment decreased PI3-K and Akt/PKB kinase activity in LPS-stimulated DC. We further examined NF-B activation in triptolide-treated DC. Under normal conditions, the majority of NF-B subunits are sequestered in the cytoplasm by I-B␣ and translocated into nucleus following I-B␣ degradation. We found that total I-B␣ levels in cell lysates of LPS-stimulated control DC were rapidly decreased while that of the LPS-stimulated and triptolide-treated DC showed no significant changes (Fig. 6B), indicating that triptolide inhibits LPS-induced I-B␣ degradation in DC. In consistent with the inhibited degradation of I-B␣ in triptolidetreated DC in response to LPS, nuclear levels of NF-B p65 subunit in triptolide-treated and LPS-stimulated DC were also lower than that of LPS-treated DC without triptolide treatment (Fig. 6B). To further understand the role of the reduced PGE2 in triptolide-mediated inhibition of PI3-K/Akt and NF-B pathway, we added exogenous PGE2 together with LPS into culture system after DC were pretreated with triptolide for 12 h. As shown in Fig. 6C, exogenous PGE2 restores the triptolideinduced inhibition of Akt kinase activity and nuclear NF-B activaction in DC. Furthermore, we investigated the effects of inhibitors of PI3-K/Akt, NF-B and COX-2 pathways on DC migration toward MIP-3. As shown in Fig. 6D, inhibitor of PI3K/Akt, NF-B or COX-2, when used respectively, could inhibit DC migration significantly. However, combined used of any kind of these inhibitors could not further increase the inhibitory effect of triptolide on DC migration toward MIP-3, further confirming the triptolide-mediated inhibition of DC migration is through PI3-K/Akt, NF-B pathways. Taken together, these results suggest that triptolide-mediated inhibition of COX-2 and CCR7 expression by decreasing TLR4 ligand-induced PI3K/Akt activity and NF-B activation in DC, resulting in the impairment of DC migration to secondary lymphoid organs. 4. Discussion Although triptolide has been used extensively as an immunosuppressive agent for the treatment of various allograft organ transplantation and autoimmune diseases, the inhibitory effects
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Fig. 5. Triptolide inhibits LPS-induced proinflammatory cytokines production by DC. DC (5 × 105 /ml) were pretreated with triptolide (5 ng/ml) for 12 h, and then stimulated with (DC-LPS, DC-Trip/LPS) or without (DC, DC-Trip) 1 g/ml LPS at indicated time points and the culture supernatants of the resting and stimulated DC were collected. RT-PCR assay of cytokines (TNF-␣, IL-1, IL-6) mRNA expression (A) and ELISA assay (B) of the proinflammatory cytokines production were performed. (C) Dose-dependent inhibitory effects of triptolide on IL-6 production by DC in response to LPS stimulation. DC pretreated with various concentrations of triptolide as indicated for 12 h, and then stimulated with LPS (1 g/ml) for 24 h, and IL-6 production measured by ELISA. Results are representative of three independent experiments, and the data presented as mean ± S.D. * P < 0.05, ** P < 0.01.
of triptolide on the functions of DC, the key player of immune initiation and regulation, have not been clearly defined. DC migration, one critical step for mature DC-initiated immune response, is orchestrated by a complex interplay between chemokines and their receptors. During the differentiation and maturation of DC, sequential and orchestrated expressions of chemokine receptors are responsible for the proper migration of DC to immune sites (Sallusto et al., 1998). Upon DC maturation, CCR1, CCR2, and CCR5 that normally direct DC infiltrations into peripheral tissues are downregulated, accompanied by the upregulation of CCR7 that are responsive to chemokines (such as ELC/CCL19/MIP-3 or SLC/CCL21/6Ckine) expressed by secondary lymphoid organs, both of which work together to
direct the emigration of DC out of tissues and the immigration of DC into LN or spleen. The chemokine receptor CCR7 and its ligands CCL19/MIP-3 and CCL21/6Ckine control myeloid DC migration into afferent lymphatic vessels and the localization of these DC within LN (Sallusto et al., 1998; MartIn-Fontecha et al., 2003; Ohl et al., 2004). Our data show that triptolide can inhibit LPS-induced upregulation of CCR7 expression in DC, and accordingly, triptolide-treated DC exhibit the impaired migration to CCL19/MIP-3 in vitro and migration to secondary lymphoid organs in vivo. Recently, Chen et al. also reported that triptolide inhibits the chemotactic response of LPS-stimulated human monocyte-derived DC (MoDC) to CCL21/6Ckine by down-regulating CCR7 expression (Chen et al., 2005). It has
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been reported that the maturation-induced up-regulation of CCR7 expression on human monocyte-derived DC (MoDC) was insufficient to allow MoDC migration to CCL19/MIP-3 and CCL21/6Ckine. MoDC require a PGE2 -provided signals to acquire the potential to migrate effectively in response to LN-derived chemokines CCL19/MIP-3 and CCL21/6Ckine (Scandella et al., 2004; Liu et al., 2006; Randolph et al., 2005). So, we went further to investigate whether triptolide inhibits LPS-induced upregulation of COX-2 expression in DC. In agreement with previous reports showing triptolide inhibits COX expression in several other cell types (Tao et al., 1998; Yao et al., 2005), we demonstrate that triptolide inhibits PGE2 production by decreasing COX-2 gene expression in DC. It has been reported recently that stimulation of immature MoDC with PGE2 alone, or in association with CD40 ligand, TNF-␣, LPS or IL-1, can induce MoDC surface expression of CCR7 and acquisition of migratory responsiveness to CCL21/SLC (Luft et al., 2002; Ardeshna et al., 2000; Lee et al., 2003). This is consistent with our results that exogenous PGE2 can restore CCR7 expression and migratory capacity of triptolide treated and LPSstimulated-DC. Taken together, these data further confirm our conclusion that triptolide impairs the migratory capacity of DC, at least partially, by inhibiting the ability of DC to produce PGE2 in response to LPS. MAPK, PI3-K/Akt signal pathways and NF-B transcription regulate COX-2 and CCR7 expression in different cell lines. We also show that triptolide can inhibit LPS-activated NF-B and PI3-K/Akt pathways but not the MAPK pathway in DC, and the inhibited PI3-K/Akt and NF-B pathways may contribute to the decreased COX-2 and CCR7 expression in DC by triptolide. In addition, the inhibitory effect by triptolide on DC migration is not due to DC apoptosis since we did not detect either apoptosis, mitochondrial membrane potential decrease, or caspase 3 activation upon DC exposure to low concentrations of triptolide (1–5 ng/ml) (data not shown). In summary, our results demonstrate that triptolide potently inhibits LPS-induced CCR7 and COX-2 expression in DC leading to the impaired DC migration to secondary lymphoid organs both in vitro and in vivo. The impaired DC migration by triptolide may be due to the reduced COX-2 and CCR7 expression via the inhibition of PI3-K/Akt kinase activity and nuclear NF-B activation in DC. Therefore, our results have provided important
Fig. 6. Triptolide inhibits LPS-induced PI3-K/Akt kinase activity and nuclear activation of NF-B in DC. (A) Triptolide inhibits PI3-K and Akt/PKB kinase activity. DC were pretreated with triptolide for 12 h, or pretreated with PI3-K/Akt inhibitor for 30 min, and then stimulated with LPS for 15 min (PI3-K) or for indicated time (Akt kinase), cell lysates were prepared, PI3-K (upper panel) and Akt kinase (lower panels) activity was assayed respectively as
described in Section 2. Similar results were observed in three separate experiments. * P < 0.05, ** P < 0.01. (B) Triptolide inhibits LPS-induced NF-B nuclear activation. Nuclear extracts were prepared from triptolide-stimulated or LPSstimulated DC as indicated and probed for NF-B, nucleoporin p62 as a nuclear protein loading control. (C) Exogenous PGE2 rescues triptolide-mediated inhibition of Akt kinase activity and nuclear NF-B activation in LPS-stimulated DC. DC were pretreated with triptolide for 12 h, and then stimulated with LPS or LPS and PGE2 together for indicated time, Akt kinase activity and nuclear NF-B activation was assayed as described in Section 2. (D) The effect of combined use of signal inhibitor and triptolide on DC migration. DC were pretreated with inhibitor of PI3-K/Akt, NF-B or COX-2 respectively for 30 min, and then treated with or not triptolide for 24 h, then in vitro chemoattraction assay of DC toward MIP-3 was detected as described in Section 2. * P < 0.05, ** P < 0.01 (* compared with LPS group). P > 0.05 ( compared with triptolide treatment group).
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