Anthrax Edema Toxin Induces Maturation of Dendritic Cells and Enhances Chemotaxis towards Macrophage Inflammatory Protein 3{beta}

Bacillus anthracis secretes two bipartite toxins, edema toxin(ET) and lethal toxin (LT), which impair immune responses andcontribute directly to the pathology associated with the diseaseanthrax. Edema factor, the catalytic subunit of ET, is an adenylatecyclase that impairs host defenses by raising cellular cyclicAMP (cAMP) levels. Synthetic cAMP analogues and compounds thatraise intracellular cAMP levels lead to phenotypic and functionalchanges in dendritic cells (DCs). Here, we demonstrate thatET induces a maturation state in human monocyte-derived DCs(MDDCs) similar to that induced by lipopolysaccharide (LPS).ET treatment results in downregulation of DC-SIGN, a markerof immature DCs, and upregulation of DC maturation markers CD83and CD86. Maturation of DCs by ET is accompanied by an increasedability to migrate toward the lymph node-homing chemokine macrophageinflammatory protein 3β, like LPS-matured DCs. Interestingly,cotreating with LT differentially affects the ET-induced maturationof MDDCs while not inhibiting ET-induced migration. These findingsreveal a mechanism by which ET impairs normal innate immunefunction and may explain the reported adjuvant effect of ET.

INTRODUCTION

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INTRODUCTION

Bacillus anthracis secretes three proteins that combine to formtwo distinct exotoxins, edema toxin (ET) and lethal toxin (LT)(13). These two exotoxins share a receptor-binding subunit,protective antigen (PA), but differ in their catalytic moieties,with the combination of PA plus edema factor (EF) forming ETand the combination of PA plus lethal factor (LF) forming LT.Following secretion, PA binds to host cells via one of two identifiedcell surface receptors, anthrax toxin receptor 1 (ANTXR1) andanthrax toxin receptor 2 (ANTXR2) (9, 54). PA must be proteolyticallyactivated by host proteases such as furin, which allows foroligomerization and subsequent binding of EF and/or LF (25,40, 42, 51). The toxin complex is then endocytosed and traffickedto an acidic endosomal compartment, where the low pH triggersa conformational change in PA, leading to an insertion in theendosomal membrane and translocation of EF and LF into the cytosol,where they induce their cytotoxic effects (1, 22, 25, 41, 66,67).

LF is a zinc-dependent metalloproteinase that cleaves and inactivatesmitogen-activated protein kinase kinases (MKKs), thereby blockingsignaling through the p38 mitogen-activated protein kinase,extracellular signal-regulated kinase, and Jun N-terminal proteinkinase pathways (16, 61, 62). LT induces cell death in macrophagesand dendritic cells (DCs) (3, 16, 22, 47, 49, 62). Independentof cell death, LT also impairs cellular responses such as cytokinesecretion, actin-based motility in neutrophils, and endothelialcell barrier function (2, 18, 64).

EF is a calcium- and calmodulin-dependent adenylate cyclasethat raises cyclic AMP (cAMP) levels (36, 37). Early work demonstratedthat ET inhibits the phagocytic process in neutrophils (45).ET has been shown to cooperate with LT to impair monocyte-derivedDC (MDDC) cytokine response and T-lymphocyte (T-cell) activationstate (46, 60). In addition, it has been hypothesized that ETacts synergistically with LT to promote death of the host (48,57).

It is becoming clear that a major role for anthrax toxins isto inhibit immune cell function during infection (8). DCs arepotentially early targets of anthrax toxins during the initialstages of B. anthracis infection, given their location at thesites of pathogen entry (11, 60). DCs are potent antigen-presentingcells (APCs) that bridge the innate and adaptive immune responsesthrough direct pathogen neutralization, cytokine production,and T-cell activation (7). These cells are present in most tissuesin an immature state, with an enhanced ability for antigen capture.Upon antigen capture, DCs undergo a maturation process and migrateto lymph nodes. Maturation is associated with reduced phagocyticand endocytic capacity, increased cytokine secretion, changesin cell surface markers, including increased membrane expressionof major histocompatibility complex class II and costimulatorymolecules, and increased T-cell stimulatory function (6, 7).Interestingly, DCs were suggested to contribute to the disseminationof spores through phagocytosis, leading to a systemic B. anthracisinfection (10, 12).

Following phagocytosis of B. anthracis spores, DCs initiatea maturation process that is counteracted through the activitiesof de novo-synthesized anthrax toxins (10-12). ET and LT eachtarget distinct DC cytokine pathways, cooperating to inhibitcytokine secretion (2, 11, 60). In addition, ET may alter DCmaturation by raising cAMP levels. Indeed, cAMP analogues oragents that raise cAMP levels (i.e., cholera toxin [CT]) leadto an aberrant maturation of DCs in which some functions associatedwith mature DCs are altered (23, 24, 33). Given the observationsthat cAMP-elevating agents induce an altered activation statein DCs, we hypothesized that ET might also modulate the functionof these cells. In this study we report on ET-induced phenotypicand functional changes in DCs, including migration of DCs towardthe lymph node-homing chemokine, macrophage inflammatory protein3β (MIP-3β). Given that ET is produced together withLT during infection, we explore how the changes induced by ETare affected by the presence of LT.

MATERIALS AND METHODS

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MATERIALS AND METHODS

Reagents and toxins. Dibutyryl cAMP (dcAMP), camptothecin, forskolin, lipopolysaccharide(LPS) from Escherichia coli, and polymyxin B were purchasedfrom Sigma-Aldrich (St. Louis, MO). CT was purchased from ListBiological Laboratories (Campbell, CA). Purified LF (30) wasprovided by J. Mogridge (University of Toronto, Toronto, Ontario,Canada). PA expression plasmid PA-pET22b was kindly providedby John Collier (Harvard Medical School, Cambridge, MA) andtransformed into E. coli BL21(DE3) cells. EF and LF(H719C) expressionplasmids EF-pET15b and LF(H719C)-pET15b were kindly providedby J. Ballard (Oklahoma University Health Sciences Center, OklahomaCity, OK) and transformed into E. coli BL21(DE3) cells. To producetoxin subunits, a fresh colony of the appropriate E. coli transformantwas inoculated into a 20-ml starter culture of Luria Bertani(LB) Lennox medium (EMD Biosciences, Inc., Darmstadt, Germany)with 100 µg/ml ampicillin and grown overnight at 37°C.The following day, a 1:50 dilution was made in a 2-liter baffledErlenmeyer flask of LB Lennox medium with 100 µg/ml ampicillin(EMD Biosciences, Inc.). The culture was allowed to grow at37°C with shaking at 250 rpm until an optical density of1.0 was reached. PA-expressing cultures were then induced witha final concentration of 1 mM isopropyl-β-D-1-thiogalactopyranoside(IPTG) (Gold Biotechnology, St. Louis, MO) and allowed to growat 30°C with shaking at 250 rpm for 4 h. EF- and LF(H719C)-expressingcultures were induced with a final concentration of 0.1 mM IPTGand allowed to grow at 16°C with shaking at 250 rpm for16 h. PA was isolated from the periplasm and purified over aMacro-Prep high Q column (Bio-Rad Laboratories, Hercules, CA).EF and LF(H719C) were purified using Ni⁺ affinity chromatography(Amersham Biosciences, Piscataway, NJ) as previously described(63). Endotoxin was removed from PA, EF, and LF(H719C) proteinpreparations by using the Detoxi-Gel endotoxin removing gel(Pierce, Rockford, IL). Purified proteins were assayed for endotoxinby using the Limulus amebocyte lysate kit (BioWhittaker, Cottonwood,AZ) that has a detection minimum of 0.03 endotoxin units/ml.

Generation of MDDCs. Monocytes were isolated from peripheral blood obtained fromhealthy donors through the Virology Core of the UCLA AIDS Institute(institutional review board approval no. 06-06-102). Monocyteswere purified using RosetteSep monocyte enrichment cocktail(StemCell Technologies, Vancouver, British Columbia, Canada)according to manufacturer's protocol. Approximately 5.0 x 10⁶cells were seeded onto a 100-mm dish and differentiated in RPMI1640 medium containing 10% fetal bovine serum (FBS) and penicillin-streptomycin-glutamine(PSG) cocktail (Invitrogen, Carlsbad, CA), plus 50 ng/ml granulocyte-macrophagecolony-stimulating factor (GM-CSF) and 100 ng/ml interleukin-4(IL-4) (PeproTech, Rocky Hill, NJ). Fresh media and cytokineswere added on day 4, and cultures were maintained for an additional3 days to complete differentiation. On day 7, nonadherent cellswere collected and combined with adherent cells released withcalcium- and magnesium-free Dulbecco's phosphate-buffered saline(PBS) (Cellgro, Manassas, VA) plus 1 mM EDTA. Approximately4.0 x 10⁵ cells per well were seeded onto a 12-well plate andtreated as indicated in the figure legends in the presence of50 ng/ml GM-CSF and 100 ng/ml IL-4.

Analysis of cell surface marker expression. MDDCs were treated as indicated in the figure legends for 48h. To measure cell surface marker expression, MDDCs were harvestedby scraping and washing with Dulbecco's PBS and were then incubatedon ice for 1 h with the following fluorophore-conjugated antibodies:anti-DC-SIGN-fluorescein isothiocyanate (FITC) (R&D Systems,Minneapolis, MN), anti-CD14-phycoerythrin (Caltag, Burlingame,CA), anti-CD83-APC (Caltag), anti-HLA-DR-APC (Caltag), anti-CD86-FITC(Caltag), and anti-CCR7-phycoerythrin (R&D Systems) dilutedin media per manufacturer's suggestions. Cells were then washedtwice with PBS and fixed in 1% paraformaldehyde. MDDC surfacemarker expression was analyzed using FACSCalibur (BD Biosciences,San Jose, CA). Data were analyzed using Cell Quest (BD) andFlowJo (Tree Star, Inc.) software.

Chemotaxis and invasion assays. MDDCs were treated as indicated in the figure legends for 48h. Following treatment, cells were collected from wells andcounted. For the chemotaxis assay, RPMI 1640 medium containing3% FBS and PSG, plus 50 ng/ml GM-CSF, 100 ng/ml IL-4, and 200ng/ml CCL19 (MIP-3β) (PeproTech), was added to the bottomof a 24-well plate. MDDCs (2.5 x 10⁴ per well) under each treatmentcondition (in triplicate) were added to the top of an uncoatedinsert with a pore size of 5 µm (Costar, Lowell, MA) inserum-free RPMI 1640 containing PSG, plus 50 ng/ml GM-CSF and100 ng/ml IL-4. Cells were allowed to migrate for 4 h at 37°C.For the invasion assay, RPMI 1640 medium containing 3% FBS andPSG, plus 50 ng/ml GM-CSF, 100 ng/ml IL-4, and 200 ng/ml CCL19,was added to the bottom of a 24-well plate. MDDCs ( 2.5 x 10⁴per well) under each treatment condition (in triplicate) wereadded to the top of a Matrigel-coated insert with a pore sizeof 8 µm (BD Biosciences) in RPMI 1640 medium containing3% FBS and PSG, plus 50 ng/ml GM-CSF and 100 ng/ml IL-4. Cellswere allowed to migrate for 22 h at 37°C. Inserts were thenremoved, cells that had not migrated were aspirated off thetop, and cells that had migrated and were adherent to the bottomof inserts were removed with trypsin (Invitrogen). Cells thatmigrated to the bottom well were collected and pooled with thetrypsinized cells. Pooled migrated cells were stained with trypanblue (Invitrogen) and counted using a hemocytometer. Statisticalanalysis was performed as described in the figure legends usingGraphPad PRISM 4 software (GraphPad Software, Inc., La Jolla,CA).

WCE and immunoblotting. Cells were treated with the indicated toxin and/or drug forthe indicated times and washed three times with cold PBS. Cellpellets were lysed on ice for 30 min with MCF7 lysis buffer(150 mM NaCl, 50 mM Tris-HCl [pH 8], 0.1% sodium dodecyl sulfate,1% Triton X-100, 5 mM MgCl₂) containing complete mini proteaseinhibitor cocktail (Roche, Mannheim, Germany). After centrifugation,the supernatants (whole-cell extracts [WCE]) were saved. Proteinconcentrations were determined using the Bio-Rad Bradford proteinassay reagent (Bio-Rad Laboratories). Sodium dodecyl sulfate-polyacrylamidegel electrophoresis and immunoblotting were performed using5 µg of WCE. Antibodies used were reactive against MEK-2(N-20) (Santa Cruz Biotechnology, Santa Cruz, CA), ${alpha}$ -tubulin(Sigma), procaspase-3 (BD Biosciences) or cleaved caspase-3(Cell Signaling Technology, Danvers, MA) (procaspase-3 and caspase-3antibodies were both kindly provided by F. Tamanoi [UCLA]).Secondary antibodies for immunoblotting were horseradish peroxidase-conjugatedgoat anti-rabbit immunoglobulin G (IgG; Zymed, San Francisco,CA) or horseradish peroxidase-conjugated goat anti-mouse IgG(H+L)(AnaSpec Inc.; San Jose, CA).

RESULTS

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RESULTS

ET induces maturation of MDDCs. During DC maturation, cell surface expression of proteins involvedin antigen presentation, DC migration, and T-cell stimulationincreases, while expression of markers of immature DCs decreases(6, 7, 50). To determine whether ET induces maturation of DCsin a manner similar to that of LPS, MDDCs were incubated witheither LPS or ET for 48 h and then assayed for cell surfacephenotype by flow cytometry. For ET challenge, cells and mediawere pretreated with the lipid A-neutralizing agent polymyxinB (43) prior to addition of toxin as previously determined toensure that any observed effects are specific to ET and notdue to potential LPS contamination (39). Treatment with eitherET or LPS led to a reduced expression of DC-SIGN, a marker ofimmature DCs (Fig. 1A). Furthermore, expression of the DC-specificmaturation marker CD83 and the costimulatory molecule CD86 wasincreased upon treatment (Fig. 1A). As predicted, treating withPA or EF alone did not alter expression of the surface markerstested (data not shown). Furthermore, treating with the cAMPanalogue dcAMP (Fig. 1A), CT, or forskolin (Fig. 1B) led toa similar maturation as induced by ET. These results suggestthat ET, like other cAMP-inducing agents, can promote maturationof DCs.

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FIG. 1. ET induces maturation of MDDCs. MDDCs were pretreated with 10 µg/ml polymyxin B for 45 min and were then treated with either 100 ng/ml LPS (no pretreatment with polymyxin B), 300 µM dcAMP, 100 ng/ml PA plus 100 ng/ml EF (ET), or no treatment (NT) (A), or 50 µM forskolin, 20 ng/ml of CT, or no treatment (B). Cells were harvested 48 h after treatment and incubated with anti-DC-SIGN-FITC, anti-CD83-APC, and anti-CD86-FITC (panel A only) and then subjected to flow cytometric analysis. The data depicted represent the results for one of six independent experiments conducted on the MDDCs generated from six different donors. The histograms depict data for the unstained sample (shaded histogram), data for the stained sample (solid black line), and the geometric mean fluorescence under each treatment condition.

ET induces migration toward MIP-3β. As a consequence of maturation induced upon encountering a pathogen,DCs migrate toward lymph nodes and present processed antigensto T cells in order to prime an adaptive immune response. Migrationto secondary lymphoid organs requires chemotaxis toward lymphnode chemokines and invasion through extracellular matrix (ECM).Movement through ECM requires the expression of adhesins andmetalloproteinases that facilitates migration (34, 58). We,therefore, next sought to determine whether ET-treated DCs chemotaxtoward the lymph node-homing chemokine, MIP-3β, and whetherthese ET-treated cells are capable of invading through ECM.MDDCs were treated with LPS, dcAMP, or ET and then assayed forchemotaxis toward MIP-3β in a transwell assay or assayedfor their ability to invade through ECM by employing a modifiedtranswell assay in which porous membranes were coated in Matrigel,a biologically active basement membrane matrix. All three treatmentsinduced robust chemotaxis toward MIP-3β and invasion throughMatrigel, compared to those induced in untreated DCs (Fig. 2A and B).The low level of migration of untreated DCs is consistent withthe lack of responsiveness of immature DCs to MIP-3β. Furthermore,whereas all three treatments induced migration toward MIP-3β,only the migration induced by ET treatment of MDDCs showed statisticalsignificance (P < 0.01) (Fig. 2A and B). Finally, robustmigration required the presence of an MIP-3β gradient,as all treatments led to a similar low level of migration asthat of untreated cells in the absence of this chemokine (datanot shown).

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FIG. 2. ET induces migration toward MIP-3β. MDDCs were pretreated with 10 µg/ml polymyxin B for 45 min and were then treated with either 100 ng/ml LPS (no pretreatment with polymyxin B), 300 µM dcAMP, 100 ng/ml PA plus 100 ng/ml EF (ET), or no treatment (NT). (A) Following 48 h of treatment, the migration of treated MDDCs toward MIP-3β was measured using a transwell assay employing uncoated inserts (5-µm pore). Data are shown as migration compared with that of immature (NT) MDDCs for each donor and are the average increase in migration for six donors ± standard deviation (SD). Statistical significance was calculated using one-way analysis of variance (ANOVA) with Bonferroni's multiple comparison test. *, P < 0.01, compared to NT. (B) Following 48 h of treatment, the migration of treated MDDCs for five donors was measured using a transwell assay employing Matrigel-coated inserts (8-µm pore). Data are shown as the increase in invasion compared with that of immature (NT) MDDCs for each donor and are the average values for five donors ± SD. Statistical significance was calculated using one-way ANOVA with Bonferroni's multiple comparison test. *, P < 0.01, compared to NT.

Effects of LT on ET-mediated DC maturation and chemotaxis. In the context of a B. anthracis infection, DCs are simultaneouslyexposed to both ET and LT. Because LT inhibits MKK signalingimportant for DC maturation and migration (2, 49, 60, 65), wenext sought to determine whether LT interferes with ET-inducedmaturation of DCs. First, changes in surface expression markersof treated MDDCs were measured. As expected, LT treatment alonedid not induce MDDC maturation (Fig. 3A). Simultaneous treatmentwith LT did not interfere with the ET-induced decrease in DC-SIGNexpression (Fig. 3A). However, the presence of LF did dampenthe ET-induced increase in both CD83 and CD86 expression (Fig.3A). This response was dependent on LF activity since cotreatingwith a catalytically inactive LF, LF(H719C) (mLT) (32), ledto similar CD83 upregulation as that induced by treating withET alone (Fig. 3B).

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FIG. 3. Effects of LT on ET-mediated DC maturation and chemotaxis. (A and B) MDDCs were pretreated with 10 µg/ml polymyxin B for 45 min and were then treated either with 100 ng/ml PA plus 100 ng/ml EF (ET), 100 ng/ml PA plus 100 ng/ml LF (LT), 100 ng/ml PA plus 100 ng/ml EF plus 100 ng/ml LF (ET/LT), or no treatment (NT) (A) or with 100 ng/ml PA plus 100 ng/ml EF (ET), 100 ng/ml PA plus 100 ng/ml EF plus 100 ng/ml LF (ET/LT), 100 ng/ml PA plus 100 ng/ml EF plus 100 ng/ml LF(H719C) (ET/mLT), or no treatment (NT) (B). Cells were harvested 48 h after treatment and incubated with anti-DC-SIGN-FITC, anti-CD83-APC, and anti-CD86-FITC and then subjected to flow cytometric analysis. The data depicted represent the results for one of five independent experiments conducted on MDDCs generated from five different donors. The histograms depict data for the unstained sample (shaded histogram), data for the stained sample (solid black line), and the geometric mean fluorescence under each treatment condition. (C) MDDCs were pretreated with 10 µg/ml polymyxin B for 45 min and then treated with either 100 ng/ml PA plus 100 ng/ml EF (ET), 100 ng/ml PA plus 100 ng/ml LF (LT), 100 ng/ml PA plus 100 ng/ml EF plus 100 ng/ml LF (ET/LT), or 100 ng/ml PA plus 100 ng/ml EF plus 100 ng/ml LF(H719C) (ET/mLT). Following 48 h of treatment, the migration of MDDCs toward MIP-3β was measured using a transwell assay employing uncoated inserts (5-µm pore). Data are shown as the increase in migration compared with that of immature (NT) MDDCs for each donor and are the average values for five donors ± SD. Statistical significance was calculated using one-way ANOVA with Bonferroni's multiple comparison test. **, P < 0.05, compared to NT. (D) Cells were pretreated with 10 µg/ml polymyxin B for 45 min and then treated for 2 h with 500 ng/ml PA plus 500 ng/ml EF (ET), 500 ng/ml PA plus 500 ng/ml LF (LT), or 500 ng/ml PA plus 500 ng/ml EF plus 500 ng/ml LF (ET/LT) or were left untreated (NT). WCEs (20 µg/sample) were analyzed by Western blotting using antibodies specific for MEK2 (N terminus) or ${alpha}$ -tubulin.

We next sought to determine if MDDCs treated with ET and LTwould still migrate toward MIP-3β. LT inhibits Hsp27-dependentneutrophil migration as well as macrophage and T-cell chemotaxis(17, 18, 52). This inhibitory effect is attributed to the abilityof LT to inhibit MKK signaling (52, 65) and so could potentiallyhave an inhibitory role in ET-induced MDDC migration. Interestingly,LF did not significantly reduce ET-induced migration (Fig. 3C)even though MEK cleavage is observed (Fig. 3D). Treatment withLT alone did not significantly increase migration to a levelabove that observed in untreated controls (Fig. 3C), a resultthat was predicted since LT treatment does not lead to MDDCmaturation (Fig. 3A).

During testing of different donors for the ability of MDDCsto migrate toward MIP-3β, we did not observe a decreasein viability following LT treatment, in contrast to previousreports (3, 49). Interestingly, while ET plus LT (ET+LT) treatmentdid not decrease viability in the majority of donors tested,one donor (donor 3) reproducibly demonstrated a reduced viabilityafter ET+LT treatment. Upon further inspection, we observedthat ET+LT treatment strongly induced caspase-3 activation indonor 3 similar to that induced by the caspase-3-activatingcompound camptothecin (Fig. 4). Strikingly, the appearance ofthe 17-kDa and 19-kDa caspase-3 fragments, indicative of activation,varied between donors (Fig. 4). Because decreased MDDC viabilitywould be seen as decreased migration in the assay presentedin Fig. 3C, the slight inhibitory effect of LT on ET-mediatedmigration may be due to either a direct inhibitory effect ofLF on migration in all donors or reduced cell viability in asubset of donors. Indeed, when data in Fig. 3C were analyzedso as to exclude donor 3, no difference in migration was seenbetween ET and ET+LT treatments (data not shown), indicatingthat LF does not directly influence ET-mediated migration.

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FIG. 4. ET and LT differentially activate caspase-3 in MDDCs. MDDCs were generated from three different healthy donors (donors 2, 3, and 6) and either treated with 100 ng/ml LPS or pretreated with 10 µg/ml polymyxin B for 45 min and then treated with 10 µM camptothecin (Camp) to induce caspase-3-mediated apoptosis, 100 ng/ml PA plus 100 ng/ml EF (ET), 100 ng/ml PA plus 100 ng/ml LF (LT), or 100 ng/ml PA plus 100 ng/ml EF plus 100 ng/ml LF (ET/LT), or no treatment (NT). Following 24 h of treatment, WCEs were generated and analyzed (5 µg/sample) by Western blotting using antibodies specific for procaspase-3 or cleaved caspase-3.

DISCUSSION

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DISCUSSION

DCs are potent APCs with an important role in the developmentof pathogen-specific immune responses. Phenotypic and functionalchanges mark the DC maturation response to pathogen and inflammatorystimuli. Here, we demonstrate that in an in vitro system, ETinduces maturation of human DCs and migration toward the lymphnode-homing chemokine MIP-3β.

Toxins that increase cAMP levels induce expression of DC maturationsurface markers, such as CD83, CD86, and HLA-DR (4, 5, 23, 56),a response strictly dependent on cAMP (4, 5). However, thismaturation is aberrant, and DCs activated by such toxins arehampered in their ability to produce cytokines, such as IL-12and tumor necrosis factor alpha. Also, maturation of DCs inducedby the combination of LPS and CT or by Bordetella bronchisepticainfection leads to secretion of high levels of the anti-inflammatorycytokine IL-10 (23, 56). In a similar fashion, anthrax ET altersDC cytokine profiles during a B. anthracis infection, with adecrease in tumor necrosis factor alpha and IL-12 secretionand an increase in IL-10 secretion (11, 60).

In this study, we tested the hypothesis that ET promotes DCmaturation and determined whether LT interferes with this response.We found that ET induced maturation, as determined by changesin expression of three DC surface markers. Interestingly, whileLT did not affect the ET-induced decrease in DC-SIGN expression,it did dampen the increase in both CD83 and CD86 expression.The observed ET-induced DC maturation is accompanied by an increasedability to migrate toward the lymph node-homing chemokine, MIP-3β.MKK signaling plays an important role in chemotaxis (65), andLT inhibits chemotaxis of macrophages, T cells, and neutrophilstoward stromal cell-derived factor 1 ${alpha}$ , MIP-1 ${alpha}$ , and N-formyl-methionyl-leucyl-phenylalanine,respectively (18, 52). Strikingly, LT did not block ET-inducedmigration toward MIP-3β even though MEK cleavage is observed.This suggests that ET-induced migration is, in large part, independentof MKK signaling. Further, this differential effect of LT onET-induced responses suggests that anthrax toxins have evolvedto work cooperatively in order to modulate specific immune cellfunctions.

Although we observed no LT-mediated cell death in MDDCs fromthe donors tested, we did note that MDDCs from a single donorshowed decreased viability following treatment with both ETand LT. Further characterization of cells from this donor demonstratedthat treatment with ET activated caspase-3, a key mediator ofapoptosis. In addition, caspase-3 activation increased withthe addition of LF, suggesting that LT and ET act synergisticallyto induce DC death. Donor variation in capase-3 activation andcell death implies that additional factors, including host geneticbackground, influence host response to anthrax toxins. Indeed,genetic background differences have been suggested to influencethe manner in which LT mediates DC killing in mice (3).

Anthrax toxins alter immune cell functions, including alterationof the chemotactic response. LT inactivation of MKK signalingleads to inhibition of neutrophil, macrophage, and T-cell chemotaxis(17, 18, 52), while ET inhibits endothelial cell, macrophage,and T-cell migration (29, 52). Interestingly, a recent reportdemonstrated that ET increases motility of infected macrophagesin the presence of a serum gradient (31). The contrasting migratorycapacities of ET-treated macrophages observed by the two studiesmay be due to differences in assay setup, differences in macrophagegeneration, and/or the chemotactic signal used (31, 52). Indeed,distinct functional subsets of macrophages have been described,and their phenotype may change in response to alterations intheir microenvironment (59). It is conceivable that ET may havedifferential effects on macrophages depending on their phenotypicprogramming. Our findings are in line with those of Kim et al.(31) and support a role for ET-mediated maturation and inductionof migration in monocyte-lineage cell types.

Although cAMP contributes to DC chemotaxis (38, 53), prostaglandinE₂, which increases cAMP levels, is not able to significantlypromote migration toward lymph node-homing chemokines in theabsence of an additional maturation signal (38, 53). In contrast,the ET-induced increase in macrophage motility is dependentonly on the ability of EF to raise cAMP levels (31). In thepresent work, we show that cAMP signaling induced by ET is sufficientto induce DC chemotaxis and invasion through Matrigel. The abilityof ET to promote migration toward MIP-3β may be due inpart to the potency of the catalytic component of ET (EF) orthe manner in which EF induces cAMP signaling (15, 28, 37).Why does ET promote MDDC migration? Given the positioning ofDCs at sites of infection, the ability of ET to activate andpromote exit of these cells from the periphery may be a mechanismfor enhanced dissemination of B. anthracis into sterile tissuesor for immune evasion. Recently, it was demonstrated that ETproduced by germinating spores promotes migration of macrophagesin the presence of a serum gradient (31). It was suggested thatET-induced migration may serve to accelerate the transport ofintracellular germinating bacilli to regional lymph nodes, therebyestablishing a systemic infection (31). Alveolar macrophageshave been implicated in promoting spore dissemination leadingto establishment of the systemic infection, although recentstudies support a role for lung DCs in this process (10, 12,26). Thus, in a manner analogous to that proposed by Kim andcolleagues (31), spores germinating within DCs may use ET topromote rapid migration from the lung, allowing bacilli to enterthe circulatory system. However, it is not clear whether ETis produced before DCs or macrophages exit the lung, and thetemporal relationship between germination, toxin production,and phagocyte migration must be tested before conclusions canbe drawn regarding the relevance of this model. Alternatively,ET produced during a localized infection, such as is the casefor cutaneous anthrax, may serve an immune evasion role by promotingefflux of DCs away from the site of infection.

Regardless of its role in virulence, the ability of ET to promoteDC migration toward MIP-3β is important for the potentialuse of ET as an adjuvant in a similar fashion as that proposedfor other cAMP-elevating agents (19, 21). Several bacterialproducts, including the enterotoxins produced by Vibrio choleraeCT and E. coli heat labile toxin (hLT), function as potent mucosaladjuvants (21, 27). In particular, CT has been shown to promotemigration of antigen-loaded DCs to T- and B-cell areas of thePeyer's patches (55). Yet, while CT and hLT are useful adjuvants,their ability to induce watery diarrhea upon oral administrationor the link of nasal administration of hLT to effects on thecentral nervous system may limit their use in humans (35, 44).ET also functions as an adjuvant for mucosal immune responses(19), but unlike hLT, it does not target central-nervous-systemtissues, making ET an alternative to the enterotoxin adjuvants(14, 19, 44). Finally, the fact that ET promotes DC migrationtoward MIP-3β may represent a means to boost the efficacyof vaccines consisting of inactive pathogens, inert particles,or soluble antigens that may not trigger DC migration or maturation(20, 55).

ACKNOWLEDGMENTS

This work was funded by National Institutes of Health (NIH)award AI057870 to K.A.B. F.J.M.-A. is supported by the NIH award5F31AI061837-03. We also acknowledge support of the UCLA JonssonComprehensive Cancer Center (JCCC) and the UCLA AIDS Instituteand the flow cytometry, virology, mucosal immunology cores,which are supported by the NIH awards CA-16042 and AI-28697(UCLA-CFAR grant), the UCLA AIDS Institute, and the David GeffenSchool of Medicine at UCLA.

FOOTNOTES

* Corresponding author. Mailing address: Department of Microbiology, Immunology, & Molecular Genetics, University of California at Los Angeles, 609 Charles E. Young Dr. East, Los Angeles, CA 90095. Phone: (310) 206-7465. Fax: (310) 206-5231. E-mail: kbradley{at}microbio.ucla.edu

Published ahead of print on 9 March 2009.

Editor: J. B. Bliska

REFERENCES

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