Cholera toxin inhibits IL-12 production and CD8α⁺ dendritic cell differentiation by cAMP-mediated inhibition of IRF8 function

Prior studies demonstrated the ability of CT to inhibit the production of IL-12 from human monocytes and monocyte-derived DCs (5). We initially determined whether this also applied to freshly isolated mouse conventional DCs (cDCs), and whether it extended to other Gs protein agonists. CT, as well as agonists for β2-adrenergic receptor (salbutamol), and the adenosine A2a receptor (CGS 21680; 2-p-[2-Carboxyethyl] phenethylamino-5-N-ethylcarboxamidoadenosine hydrochloride) suppressed the production of both IL-12p40 and IL-12p70 from spleen cDCs stimulated with SAC and IFN-γ (Fig. S1), whereas IL-10 production was not suppressed (unpublished data).

CT has been shown to induce Th2 responses after oral administration with soluble protein antigens; however, because CT is normally given with inactivated proteins, the extent to which CT can modify the helper T (Th) cell phenotype during the induction of strong Th1 responses, as occurs after infection with intracellular pathogens, is not clear. Furthermore, when given intranasally, or orally with less purified antigens, CT has been reported to induce Th1 and Th2 responses (8, 9). Therefore, we decided to test the ability of CT to modify Th1 responses in mice during systemic infection of BALB/c mice with a high dose of the ME-49 strain of T. gondii. In this model, mice develop IL-12–dependent Th1 responses that clear the infection with 100% survival (10). As shown in Fig. 1, systemic administration of CT during the course of infection resulted in 100% mortality with nearly identical kinetics to that occurring after systemic neutralization of IL-12 (Fig. 1 A). In addition, parasite loads were high (Fig. 1 B) and serum levels of IL-12 and IFN-γ were suppressed in both CT and anti–IL-12–treated groups (Fig. 1 C), consistent with CT having a dominant inhibitory effect on IL-12 production and the induction of Th1 responses to this infection.

Although it is likely that a variety of cells produce IL-12 during infection with T. gondii in vivo, including macrophages (11) and neutrophils (12), CD8α⁺ cDCs are thought to be one major source of this essential cytokine (13). We therefore asked whether CT administration affects IL-12 production by cDC subsets in vivo. Although mice treated with CT showed a 20–30% reduction of total cell numbers in the spleen, there was a dramatic preferential reduction of CD11c⁺ CD8α⁺ cDCs compared with CD11b⁺ cDCs (Fig. 2 A) and other lymphocyte compartments (Fig. 2 B). The loss of CD8α⁺ cDCs was not caused by apoptosis (Fig. S2); was transient, with cell numbers returning to normal levels by day 6; and was dose dependent, with 5 µg of CT having maximal and 0.5 µg of CT having minimal effects (Fig. S3 B). Moreover, no difference in the viability of splenocytes or peripheral blood cells from CT-treated versus PBS-treated controls or CTB-treated mice was found (unpublished data). Treatment with CTB had no effect on CD8α⁺ cDC numbers (unpublished data), indicating that ADP ribosylate activity of CTA was required for the loss of CD8α⁺ cDCs.

The loss of IL-12–producing CD8α⁺ cDCs may contribute to the effects seen with CT in T. gondii infection. To address this issue more directly, we infected mice after treatment with 0.5 µg of CT, which has only minor effects on CD8α⁺ cDC numbers. At this dose, the effects of CT on the course of T. gondii infection were highly variable, yet in some experiments we observed a significant effect on survival and parasite load, despite only a minor change in CD8α⁺ cDC numbers (Fig. S3 and not depicted). To test the possibility that CT may affect IL-12 production independent of CD8α⁺ cDC loss, we gave graded doses of CT to mice, administered LPS systemically, and measured IL-12 and IFN-γ levels in the serum. Prior studies have shown that CD8α⁺ cDCs are a significant source of IL-12 under these conditions (13), and that IFN-γ production after LPS administration can be blocked with antibodies to IL-12p40 (5). We found that low doses of CT (0.5 µg) consistently suppressed both IL-12 and IFN-γ production in response to LPS (Fig. S3). Furthermore, the total percentages of CD4⁺ and CD8⁺ T cells, as well as CD19⁺ B cells, were unchanged after CT treatment (Fig. 2 B). These studies suggest that systemic administration of CT has two dose-dependent effects in vivo, one resulting in the loss of CD8α⁺ cDCs and the other resulting in the inhibition of IL-12 production by DCs and possibly other cells.

In our initial studies using human monocytes stimulated with LPS alone or LPS and IFN-γ, CT had no effect on the activation of signaling molecules positively regulating IL-12 production, such as NF-κB, STAT-1, or the mitogen-activated protein kinases (MAPKs) p38 and JNK (unpublished data). In addition, CT had no effect on the activation of extracellular signal-regulated kinase MAPK, which is a known negative regulator of IL-12 expression (unpublished data). Moreover, CT treatment did not affect the surface expression of Toll-like receptor (TLR)-4 or IFN-γ receptor on LPS-stimulated or unstimulated monocytes (unpublished data) or the IFN-γ receptor signaling pathway, as IFN-γ–induced up-regulation of HLA-DR and CD40 was unaffected (unpublished data).

We next focused our study on the effect of CT on the function of IRF8, which is a transcription factor expressed by DCs, macrophages, granulocytes, and B cells; it is a positive regulator of IL-12p35 and IL-12p40 transcription (14–17) and has a central role in the differentiation of CD8α⁺ cDC and plasmacytoid DC (pDC) populations. Furthermore, production of IL-12 in T. gondii infection is dependent on IRF8 (11). IRF8^−/− mice lack the CD8α⁺ and pDCs, although they have CD11c⁺CD8α⁻ cDCs (18–21). Moreover, in these mice, the residual IRF8^−/− cDCs display impaired responsiveness to TLR-mediated signals and fail to produce IL-12p40 (18). Thus, IRF8 is essential for both IL-12 production by myeloid cells including cDCs and the development of CD8α⁺ cDCs that likely represent an IL-12–producing DC subsets during T. gondii infection.

We initially studied the dynamic interaction of IRF8 with chromatin in live cells using fluorescence recovery after photobleaching (FRAP) analysis, a method that was recently applied to the study of IRF8–chromatin interactions (22). For these studies, IRF8^−/− Tot2 cells were stably transduced with IRF8-GFP that localizes to the nucleus. In the nucleus, IRF8-GFP is present in two functionally distinct forms: a fraction of low-mobility IRF8 molecules engaged in interactions with chromatin and a pool of highly mobile, unbound IRF8. As depicted in Fig. 3, in untreated IRF8-transduced Tot2 cells, a small fraction of immobile chromatin-bound IRF8 molecules was present, and 80% of the initial fluorescence signal is recovered within 4 s after photobleaching. When cells were stimulated with LPS and IFN-γ that favored IRF8–chromatin interaction, an increase in the immobile IRF8 pool, and a consequent reduction of the highly mobile unbound IRF8 fraction, occurred, resulting in significantly slower fluorescence recovery. In these samples, only 60% of the recovery occurred within the first 4 s of analysis, and the maximal recovery achieved was 70% of initial signal compared with 90% observed in untreated cells (Fig. 3, red vs. black line). Treatment with CT, but not CTB, before stimulation with LPS and IFN-γ produced a recovery curve similar to that of unstimulated cells (Fig. 3, B and C). Similar to what we observed with CT, cells pretreated with the G protein–independent adenylate cyclase activator forskolin or the membrane-permeable cAMP analogue dbcAMP and stimulated with LPS and IFN-γ displayed a quick recovery of fluorescence (Fig. 3, E and F). Importantly, treatment with CT, CTB, forskolin, or dbcAMP alone did not alter the kinetics of fluorescence recovery (Fig. 3, A and D). IRF8-GFP was not overexpressed in IRF8^−/− Tot2 cells because levels were below those of endogenous IRF8 in a macrophage cell line (22). Collectively, these data indicate that increased levels of cAMP, secondary to CT-induced Gs protein ADP ribosylation, impair the ability of IRF8 to bind to chromatin, and therefore to interact with its target promoters. Interestingly, IRF8 amplifies the transcription of its own gene (23). Thus, the cAMP-dependent inhibition of IRF8 binding to its own promotor would lead to reduced IRF8 mRNA expression, a concept that is consistent with prior studies showing cAMP-dependent IRF8 mRNA down-regulation (24, 25).

IRF8 together with IRF1 has been reported to enhance IL-12p40 gene transcription induced by LPS and IFN-γ by forming a heterocomplex that interacts with a cis-acting element resembling the ISRE (IFN-stimulated response element; ISRE-like) located in the IL-12p40 mouse promoter (15, 26). To assess if CT inhibits DNA–IRF8 interaction specifically at the level of the IL-12p40 promoter, we performed a DNA affinity binding assay using a biotinylated IL-12p40 promoter sequence containing the native ISRE-like element conjugated to streptavidin beads (Fig. 4 A). As shown in Fig. 4 B, IRF8 in nuclear extracts from RAW 264.7 cells stimulated with LPS and IFN-γ bound to the probe-conjugated beads, and this binding was completely abrogated when cells were pretreated with CT, but not CTB. Moreover, a similar inhibitory effect was exerted by dbcAMP. Significantly reduced binding of IRF8 to a probe carrying a mutation in the ISRE-like element was observed (Fig. 4 B, lane 6) indicating that IRF8 binding was dependent on the integrity of the ISRE-like element.

The same ISRE-like element is a consensus sequence for IRF1 (26), therefore we investigated the effect of CT or dbcAMP on the ability of IRF1 to interact with the IL-12p40 promoter. Similar to IRF8, stimulation with LPS and IFN-γ induced IRF1 binding; however, neither CT nor dbcAMP reduced such interaction. Moreover, binding of TFIIB, a component of the RNA polymerase II complex, to the TATA box element included in the probe sequence, was unaffected by CT treatment, indicating that the inhibition exerted by CT and dbcAMP was specific for IRF8. Moreover, similar levels of IRF8 and IRF1 were detected in nuclear extracts of all cells stimulated with LPS and IFN-γ, indicating that observed effects were not caused by reduced IRF8 expression in the nucleus (Fig. 4 C).

The concomitant ability of IRF1 and inability of IRF8 to bind to the IL-12p40 promoter in cells treated with CT or dbcAMP prompted us to determine whether CT treatment could inhibit the formation of the IRF1–IRF8 heterocomplex. Consistent with this possibility, CT markedly reduced the amount of nuclear IRF1–IRF8 complex induced by LPS and IFN-γ (Fig. 4 D). Tyrosine phosphorylation of IRF1 is a crucial event for IRF1–IRF8 complex formation (27). However, the amount of phosphorylated IRF1 induced by LPS and IFN-γ was not modified by CT (Fig. 4 E). This indicates that the ability of CT to block IRF1–IRF8 complex formation is not caused by impaired IRF1 phosphorylation.

Together with IL-12 suppression, CT can enhance the ability of TLR ligands to induce DC production of IL-6, IL-1β, and IL-10, while inhibiting their production of TNF-α (4). We confirmed the ability of CT and dbcAMP to inhibit the production of IL-12 and TNF-α, but not IL-10, in human peripheral blood monocytes, monocyte-derived DCs, and the RAW 264.7 cell line stimulated with LPS and IFN-γ (Fig. S4). Interestingly, recent data indicate that TNF-α gene transcription in mouse macrophages is also regulated by IRF1 and IRF8 (28). Therefore, it is feasible that the mechanism by which CT and cyclic AMP blocks TNF-α production is also caused by effects on IRF1–IRF8 interactions. The molecular events triggered by CT that lead to reduced IRF1–IRF8 interaction remain unclear. In addition to IRF1, IRF8 has been reported to form heterocomplexes with other binding partners, including NFAT and PU.1. Interference with physical interaction between these proteins and IRF8 might regulate its ability to associate with DNA and IRF1. In support of this possibility, increased cAMP levels have been reported to inhibit NFAT activity (29). PKA-dependent phosphorylation of NFAT increased 14–3-3 binding and reduced NFAT transcription activity at the IL-2 promotor. Whether CT and other cAMP inducers might affect such interactions is currently under investigation in our laboratories.

Results of the FRAP analysis indicate that CT induces a general inhibition of IRF8 function, suggesting that other IRF8-dependent processes might also be affected. IRF8 plays a key role not only in the activation of IL-12p35 and p40 genes, but also in the normal development of pDC and CD8α⁺ cDC populations in vivo (18–21). Moreover, IRF8-deficent bone marrow cells fail to differentiate into pDCs when cultured in the presence of FLT-3L in vitro (21). We therefore studied the effect of CT on the generation of pDCs from FLT-3L–stimulated bone marrow cultures. In keeping with previous reports, after culture with FLT-3L, one third of mouse bone marrow cells differentiated into CD11c⁺ CD11b⁻ cells that also expressed B220 and Ly6c and could therefore be identified as pDCs (Fig. 5 A). In the presence of CT, we observed a dramatic reduction of the CD11c⁺CD11b⁻ subset (from 34.6 to 4.28%), and a concomitant increase in the CD11c⁺CD11b⁺ population. In addition, only 50% of the remaining CD11c⁺CD11b⁻ cells expressed Ly6C (Fig. 5 B). Conversely, exposure to CTB did not alter FLT-3L–induced pDC differentiation (Fig. 5 C), indicating that this effect of CT is dependent on the CTA activity. Consistent with these results, and similar to our findings with CD8α⁺ DCs, pDCs were preferentially lost after CT treatment in vivo (Fig. 5 D). The ability of CT to inhibit the differentiation of pDCs from FLT-3L–treated bone marrow and to induce pDC depletion in vivo is consistent with the hypothesis that IRF8–DNA interaction is a major target of CT activity.

In summary, our findings indicate that CT can inhibit IL-12 production in vivo after infection with T. gondii, or systemic administration of LPS. In addition, we identify the inhibition of IRF8 function as a major molecular mechanism by which CT and other cAMP-inducing agents prevent IL-12 production in vitro and in vivo. Furthermore, we demonstrate that the CT-induced block of IRF8 function is associated with impaired IRF1–IRF8 heterocomplex formation that is independent of the phosphorylation state of IRF1. Finally, by blocking IRF8 function, CT may simultaneously inhibit IL-12p40 mRNA expression in macrophages and differentiated DCs and interfere with normal differentiation of CD8α⁺ DCs (and pDCs) that represent a primary source of IL-12, at least under certain stimulation conditions.

8–12-wk-old conventionally housed female BALB/c mice were obtained from the National Cancer Institute and were used as approved under animal study protocol LMI11E by the National Institute of Allergy and Infectious Disease/National Institutes of Health (NIH) Animal Care and Use Committee. The derivation and maintenance of IRF8^−/− Tot2 cells and their transfection with IRF8-GFP were previously described (30). The mouse macrophage cell line RAW 264.7 was purchased from American Type Culture Collection (ATCC).

CD11c⁺ splenocytes were isolated using anti–mouse-CD11c immunomagnetic beads (Miltenyi Biotech). Human monocytes were obtained by counterflow centrifugation-elutriation of blood collected from normal blood donors under the NIH Institutional Review Board–approved human subjects protocol 99-cc-0168. Monocyte-derived DCs were obtained by culturing purified human monocytes in GMCSF and IL-4, as previously described (5). The mouse macrophage cell line RAW 264.7 was and stimulated at 80% confluence. CD11c⁺ splenocytes, human blood monocytes, monocyte-derived DCs (Mo-DC), or the RAW 264.7 cells were cultured at 1–2 × 10⁶ cells/ml in RPMI-1640 and supplemented with 10% FCS, 100 µg/ml penicillin, 100 µg/ml streptomycin, 50 µg/ml gentamicin, 5% NCTC-109 media (all from Invitrogen), 15 mM Hepes, and 200 mM glutamine (cRPMI) at 37°C and 5% CO₂ with various stimuli as indicated. CT was purchased from List Biological Laboratories. Staphylococcus aureus, Cowan's strain I (SAC; Pansorbin) was obtained from Calbiochem. Recombinant mouse IFN-γ was obtained from BD. LPS (from Escherichia coli, serotype 0127:B8) and salbutamol were purchased from Sigma-Aldrich. Anti–mouse–IL-12 (clone C17.6) was originally provided by G. Trinchieri (National Cancer Institute, Bethesda, MD). CGS21680 and ZM241385 were obtained from Tocris Bioscience. Cytokine levels in cell culture supernatants and mouse sera were measured by ELISA using standard kits (BD). ELISA lower limit of detection was 50 pg/ml for IFN-γ and IL-12 p70 and 30 pg/ml for IL-12 p40.

T. gondii cysts (ME-49 strain) were prepared from brains of infected mice as previously described (31). BALB/c mice were untreated or treated with two doses of CT (5 µg) or anti–IL-12 antibody (1.8 mg) given i.p. at 3-d intervals. Mice were infected with 100 cysts given i.p. 1 d after the first dose of CT or anti–IL-12. The percentage of infected peritoneal exudate cells was determined using stained cytospin preparations. 500 cells were counted for each sample. 5 d after infection with T. gondii, blood was collected from individual mice for cytokine quantification.

Spleen cells, or CD11c⁺ enriched splenocytes were analyzed by flow cytometry using antibodies for cDCs (anti-CD11c, HL3), B cells (anti-CD19, 1D3), T cells (anti-CD3, 145-2C11, anti-CD4, GK1.5, anti-CD8, Ly-2), and pDCs (anti-B220, RA3-6B2, anti-Ly-6C, AL-21, mPDCA-1, JF05-1C2.4.1; all from BD). Labeled cells were analyzed using a FACSCalibur flow cytometer (BD), and data was analyzed using FlowJo software (Tree Star, Inc.).

FRAP for IRF8-chromatin interactions was performed on IRF8-GFP–transduced IRF8^−/− Tot2 cells as previously described (22). Tot2 cells plated on a chambered coverslip in 1% methylcellulose (MethoCult; Stem Cell Technologies) were kept at 37°C by using an air stream stage incubator (ASI 400; Nevtek). Live-cell imaging was performed on a 510 confocal microscope (Carl Zeiss, Inc.) by using the 488-nm line of an Ar laser with a 100×, 1.3-numerical aperture oil immersion objective. Photobleaching was performed on a small circular area (0.5-µm radius, 25 pixels) in the nucleus at the maximum laser power. 30 prebleach images were acquired before a bleach pulse of 115 ms. Fluorescence recovery was monitored at low laser intensity (0.2% of a 45-mW laser) at 45-ms intervals for 16 s. FRAP experiments were performed on at least 15 independent cells, and data were averaged to generate a single FRAP curve. FRAP data obtained using cells before and after stimulation with LPS+IFN-γ were analyzed to fit to the recently developed mathematical binding models to confirm and support our initial hypothesis on the two fractions of IRF8 (32).

Nuclear extracts from RAW 264.7 cells were obtained using NE-PER extraction kit following manufacturer's instructions. 5′-Biotin-labeled DNA sequence corresponding to the −81 to −24 region of the murine IL-12p40 promoter containing a known ISRE-like element, as well as a control sequence containing a substitution mutation in the region of the ISRE-like element were purchased from Invitrogen. Biotin-labeled probes were conjugated to 200 µg of streptavidin-coated magnetic beads (Dynabeads M-280 streptavidin; Dynal) in Tris-EDTA buffer (pH 7.9) and were blocked with 0.5% bovine serum albumin in TGED buffer (10 mM Tris–HCl, pH 8.0, 10% vol/vol glycerol, 0.1 mM ethylenediaminetetraacetic acid, and 0.01% Triton X-100) overnight at 4°C. The mixtures of probe-conjugated beads and 500 µg of nuclear extracts in 500 µl of TGED with the final concentration of NaCl at 100 mM were subsequently incubated at 4°C for 4 h. Beads were extensively washed with TGED buffer, and DNA-bound proteins were eluted using TGED with 1 M NaCl. Proteins were separated by SDS-PAGE for immunoblot detection with rabbit anti-IRF8 antibody. For coimmunoprecipitations, 500 µg of nuclear extracts from RAW264.7 cells diluted 1:3 with wash buffer (50 mM Tris-HCl, 1 mM PMSF, Complete protease inhibitor cocktail [Roche], and phosphatase inhibitor cocktail 1 and 2 [Sigma-Aldrich]) were incubated with mouse monoclonal anti-IRF1 (2.5 µg/sample; Santa Cruz Biotechnology, Inc.) or irrelevant IgG over night at 4°C in a rotator and recovered with 25 µl of G protein–conjugated magnetic beads (Active Motifs). Samples were subjected to SDS-PAGE and Western blot analysis with rabbit anti-IRF8 or rabbit anti-IRF1 (Santa Cruz Biotechnology). For determining IRF1 phosphorylation, mouse anti-phosphotyrosine monoclonal cocktail (5 µl/sample, 4G10; Millipore) or 2.5 µg of irrelevant IgG were incubated overnight at 4°C with 25 µl G protein–conjugated magnetic beads. Beads were then washed twice with ice-cold PBS and incubated (90 min at 4°C in a rotator) with 1 mg of nuclear extract. After extensive washing, samples were subjected to SDS-PAGE and Western blot analysis with rabbit anti-IRF1 antibody (Santa Cruz Biotechnology, Inc.).

Bone marrow from 6-wk-old mice was cultured at 5 × 10⁵ cells/ml in D-MEM (Biofluids, Inc.) supplemented with 10% fetal bovine serum and containing 100 ng/ml recombinant human FLT-3L (PeproTech). Medium together with FLT-3L was changed at days 3 and 6 of culture. CT or CTB was added at 20 ng/ml final concentration at the beginning of the culture and at every medium change. After 9 d, cells were analyzed by flow cytometry using fluorochrome-conjugated monoclonal antibodies against CD11c, CD11b, B220, and Ly6C (BD).

Results represent the mean ± SD where applicable. Statistical significance of differences was determined by the Student's t test.

Fig. S1 demonstrates the regulation of IL-12 production from murine cDCs by the β₂ adrenergic agonist Salbutamol, the adenosine A2a receptor agonist CGS21680, the adenosine A2a receptor antagonist ZM241385 and CT. Fig. S2 shows the time course of CD8α⁺ cDC loss as well as annexin V staining of CD8α⁺ cDCs after CT administration in vivo. Fig. S3 demonstrates the effect of CT dose on CD8α⁺ DC loss, and the effect on IL-12 and IFN-γ production in vivo. Fig. S4 demonstrates the effect of CT and dbcAMP on IL-12, TNF-α, and IL-10 production from human monocytes, monocyte-derived DCs, and RAW 264.7 cells.

This work was supported by research funds from the Division of Intramural Research, National Institutes of Allergy and Infectious Diseases, National Institutes of Health.

The authors have no conflicting financial interests.