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Cell Functional InteractionsDendritic−Activin A as a Mediator of NK

SozzaniBadolato, Angela Gismondi, Angela Santoni and Silvano Pascal Seeger, Daniela Bosisio, Silvia Parolini, Raffaele

http://www.jimmunol.org/content/192/3/1241doi: 10.4049/jimmunol.1301487January 2014;

2014; 192:1241-1248; Prepublished online 6J Immunol 

MaterialSupplementary

7.DCSupplementalhttp://www.jimmunol.org/content/suppl/2014/01/05/jimmunol.130148

Referenceshttp://www.jimmunol.org/content/192/3/1241.full#ref-list-1

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The Journal of Immunology

Activin A as a Mediator of NK–Dendritic Cell FunctionalInteractions

Pascal Seeger,* Daniela Bosisio,* Silvia Parolini,* Raffaele Badolato,†,‡ Angela Gismondi,x

Angela Santoni,x and Silvano Sozzani*,{

The interaction of NK cells with dendritic cells (DCs) results in reciprocal cell activation through the interaction of membrane proteins

and the release of soluble factors. In this article, we report that in NK–DC cocultures, among a set of 84 cytokines investigated, activin A

was the second highest induced gene, with CXCL8 being the most upregulated one. Activin A is a member of the TGF-b superfamily

and was previously shown to possess both proinflammatory and anti-inflammatory activities. In NK–DC cocultures, the induction of

activin A required cell contact and was dependent on the presence of proinflammatory cytokines (i.e., IFN-g, TNF-a, and GM-CSF),

as well as on NK cell–mediated DC killing. CD1+ DCs were the main activin A producer cells among myeloid blood DC subsets. In

NK–DC cocultures, inhibition of activin A by follistatin, a natural inhibitory protein, or by a specific blocking Ab, resulted in the

upregulation of proinflammatory cytokine release (i.e., IL-6, IL-8, TNF-a) by DCs and in the increase of DC maturation. In

conclusion, our study reports that activin A, produced during NK–DC interactions, represents a relevant negative feedback mecha-

nism that might function to prevent excessive immune activation by DCs. The Journal of Immunology, 2014, 192: 1241–1248.

Dendritic cells (DCs) and NK cells are essential players inearly immune defense against infections and cancer (1,2). In vitro and in vivo studies have provided evidence

that the activity of DCs and NK cells is regulated by reciprocalinteractions of these two cell types (3–7). Mature DCs induce NKcell proliferation and enhance their cytolytic activity (8, 9). Incontrast, activated NK cells promote the maturation of DCs andinduce immature DC death according to the NK/DC reciprocalcell ratio (3, 4, 10). Importantly, during this interaction, both celltypes release a number of cytokines able to potentiate immunefunctions (6, 11, 12). As a consequence, NK–DC interactions areimportant in the regulation of the early phase of the innate im-mune response and have a significant impact on adaptive immu-nity. Indeed, NK–DC interactions have been found to be pivotalfor antitumor response and in antiviral immunity, such as in CMVand influenza infections (6, 13, 14).The cross-talk between DCs and NK cells is regulated by different

mechanisms that include receptor–ligand interactions, as well asthe release of soluble factors. For instance, CD56brightCD162 NKcell expansion is induced by membrane-bound IL-15 on mature

DCs (8, 11, 15). NK cell activation was found to be mediated viathe release of TNF-a and IFN-g by DCs and activated NK cellsupon triggering of the NK cell receptor NKp30 (3, 16). DC-derived IL-12 and IL-18 were shown to induce IFN-g expres-sion by NK cells in the context of a positive feedback loop (17,18). Therefore, many evidences document the positive, proin-flammatory activation program activated by the NK–DC cross-talkin vitro and in vivo. Conversely, less information is available onthe existence of mechanisms that limit the output of NK–DCinteractions. For instance, it is known that NK cell–mediated DCmaturation and killing is negatively regulated by MHC-specificNK cell receptors, such as the killer Ig-like receptors and theCD94/NKG2A heterodimer, by elevating the threshold for DCactivation and killing (16). However, only little attention has beendevoted to the role of secreted molecules in limiting exaggeratedNK–DC immune activation.Activin A is a member of the TGF-b superfamily endowed with

pleiotropic functions, originally described as a regulator of follicle-stimulating hormone produced by the pituitary gland (19, 20).Similarly to other members of the TGF-b superfamily, activin Asignals via Smad- and MAPK-dependent pathways upon binding toserine/threonine kinase receptor proteins, namely, activin receptorstype I (Alk2, Alk4) and type II (ActRIIA, ActRIIB). During thepast few years, activin A became increasingly appreciated as amodulator of immune response, having both proinflammatory andanti-inflammatory properties. Activin A is expressed by many celltypes, including endothelial cells, mast cells, monocytes, macro-phages, and DCs (21–23). Recently, activin A has been suggested toattenuate the ability of DCs and NK cells to release certain cyto-kines and chemokines (24, 25). This study confirms and extendsthese observations showing that activin A is highly upregulatedduring NK–DC interactions and acts as a negative feedback mech-anism in the reciprocal regulation of these two cell types.

Materials and MethodsCell preparation and cultures

CD14+ monocytes were isolated from buffy coats (Centro TrasfusionaleBrescia, Italy) by positive magnetic separation using CD14 immuno-magnetic beads (Miltenyi Biotec, Bergisch-Gladbach, Germany). To gen-

*Department of Molecular and Translational Medicine, University of Brescia, 25123Brescia, Italy; †Institute of Molecular Medicine “Angelo Nocivelli,” University ofBrescia, 25123 Brescia, Italy; ‡Department of Clinical and Experimental Sciences,University of Brescia, 25123 Brescia, Italy; xDepartment of Molecular Medicine, LaSapienza University, 00161 Rome, Italy; and {Humanitas Clinical and ResearchCenter, 20089 Rozzano, Italy

Received for publication June 6, 2013. Accepted for publication November 29, 2013.

This work was supported by the Associazione Italiana per la Ricerca sul Cancro,Istituto Superiore di Sanita, Ministero dell’Istruzione dell’Universita e della Ricerca,and Cariplo, Ministero della Salute, FP-7 EU Project “MOODINFLAME.”

The sequences presented in this article have been submitted to the Gene ExpressionOmnibus (http://www.ncbi.nlm.nih.gov/geo/) under accession number GSE52306.

Address correspondence and reprint requests to Prof. Silvano Sozzani, Departmentof Molecular and Translational Medicine, University of Brescia, Viale Europa 11,25123 Brescia, Italy. E-mail address: sozzani@med.unibs.it

The online version of this article contains supplemental material.

Abbreviations used in this article: BMP, bone morphogenetic protein; DC, dendriticcell; HPS-2, Hermansky–Pudlak syndrome type 2; MoDC, monocyte-derived DC;pDC, plasmacytoid DC; slanDC, 6-sulfo LacNAc DC.

Copyright� 2014 by The American Association of Immunologists, Inc. 0022-1767/14/$16.00

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erate monocyte-derived DCs (MoDCs), monocytes were cultured for 6 d intissue culture plates (Costar, Corning) in RPMI 1640 supplemented with10% heat-inactivated FCS (Lonza, Basel, Switzerland), 100 U/ml peni-cillin, 100 mg/ml streptomycin, 2 mM L-glutamine (Invitrogen, Carlsbad,CA); 50 ng/ml GM-CSF and 20 ng/ml IL-4 (Peprotech, London, U.K.)were added at day 0. MoDC maturation was induced by incubation with100 ng/ml LPS (Sigma-Aldrich, St. Louis, MO) for 24 h (26). Necrosis ofDCs and K562 cells was achieved via four cycles of freezing (in dry ice)and thawing (at 37˚C). CD141+ DCs were isolated using the CD141+

Microbead Kit (Miltenyi Biotec). CD1c+ DCs, 6-sulfo LacNAc DCs(slanDCs), and plasmacytoid DCs (pDCs) were isolated using the corre-sponding Cell Isolation Kit (Miltenyi Biotec). NK cells were isolated usingthe NK Cell Isolation Kit (Miltenyi Biotec). The purity of NK cells afterisolation was evaluated by allophycocyanin-conjugated anti-CD3 and PE-conjugated anti-CD56 mAb staining (Biolegend, San Diego, CA) by flowcytometry and was routinely .96% (CD3 was ,1%). NK cells were ac-tivated by incubation with 40 ng/ml IL-15 (Peprotech) for 24 h. Beforecoculture, NK cells and DCs were extensively washed. If not otherwisementioned, DCs (105 cells) were cocultured with allogeneic NK cells ata ratio of 1:1 in flat-bottom tissue culture plates for 24 h. NK–DCcocultures were performed in completed RPMI 1640 medium at the NK/DC ratio indicated. In some experiments, NK cells and DCs were separatedusing transwell inserts with 0.4-mm membranes (Costar). For someexperiments, cocultured NK cells and DCs were magnetically separatedusing anti-CD56-Microbeads (Miltenyi Biotec). NK cells purified fromPBMCs of Hermansky–Pudlak syndrome type 2 (HPS-2) patients and healthydonors were cultured on irradiated feeder cells in the presence of 100 U/mlIL-2 (Proleukin; Chiron, Emeryville, CA) and 1.5 ng/ml PHA (Invitrogen) toobtain polyclonal NK cell populations. The cells were then washed andcultured for an additional 24 h in the presence of 40 ng/ml IL-15.

Cytokines and reagents

The following Abs were used: 5 mg/ml anti–TNF-a (clone 1825; R&DSystems, Minneapolis, MN), 10 mg/ml anti–GM-CSF (polyclonal; R&DSystems), 1 mg/ml anti–IL-1b (polyclonal; R&D Systems), isotype controlAb (polyclonal; R&D Systems). The anti–IFN-g (clone B133) was kindlyprovided by Prof. Cassatella (University of Verona, Italy). For receptorcross-linking, the wells were coated with 10 mg/ml anti-NKp30 (clone P30-15; Biolegend) or anti-CD16 (clone 3G8; Biolegend). Activin A activitywas blocked by the addition of 200 ng/ml follistatin (Peprotech), 10 mg/mlanti-activin A (polyclonal; R&D Systems), or 10 mM SB431542 (Sigma-Aldrich). The activity of bone morphogenetic protein (BMP) signaling wasblocked by the addition of 5 mM dorsomorphin (Sigma-Aldrich). NK cellswere stimulated with 200 U/ml IL-2 (Proleukin), 100 ng/ml IL-18 (MBL,Nagoya, Japan), 2 ng/ml IL-12, 40 ng/ml IL-15 (Peprotech), or 20 ng/mlPMA (Sigma-Aldrich) and 5 mM ionomycin (Sigma-Aldrich).

Real-time PCR

RNA was extracted using TRIzol reagent (Invitrogen, Carlsbad, CA)according to the manufacturer’s instructions. After RNA purification,samples were treated with DNase to remove contaminating genomic DNA(DNaseI Amplification grade; Invitrogen). Reverse transcription was per-formed using random hexamers and Superscript II Reverse transcriptase(Invitrogen). Gene-specific primers were purchased from Fambiotech(Brescia, Italy). The iQ SYBR Green Supermix (Bio-Rad, Hercules, CA)was used to run relative quantitative real-time PCR of the samplesaccording to the manufacturer’s instructions. Cytokine expression in NK–DC cocultures was evaluated by a PCR array–based screening including 84common cytokines (RT2 Profiler PCR Array; Qiagen). Reactions were runon an iCycler (Bio-Rad), and generated products were analyzed withiCycler iQ Optical System Software (Version 3.0a; Bio-Rad). Gene ex-pression was routinely normalized based on HPRT content. The primersused were 59-CCA GTC AAC AGG GGA CAT AAA-39 (hHPRT-f), 59-CAC AAT CAA GAC ATT CTT TCC AGT-39 (hHPRT-r), 59-GCA GAAATG AAT GAA CTT ATG GA-39 (hINHBA-f), 59-GTC TTC CTG GCTGTT CCT GAC T-39 (hINHBA-r), 59-GCG TGC TAATGG TGG AAA-39(TGF-b1-f), 59-CGG TGA CAT CAA AGA TAA CCA C-39 (TGF-b1-r),59-GCA GGT CAT TCA GAT GTA GCG G-39 (IFN-g-f), 59-CCA CACTCT TTT GGA TGC TCT GG-39 (IFN-g-r), 59-CAA GCC TGT AGCCCA TGT TGT AG-39 (TNF-a-f), 59-CCT GGG AGT AGA TGA GGTACA GG-39 (TNF-a-r).

Cytokine evaluation by ELISA

The protein concentration of human Activin A, TNF-a, IL-6, IL-8, IL-1b,GM-CSF, TGF-b1, TGF-b2, BMP-6, and IFN-g was evaluated using col-orimetric ELISA DuoSet (R&D Systems), according to the manufacturer’sinstructions.

Flow cytometric analysis

Surface phenotype analysis was performed using PE-conjugated anti-CD86(clone IT2.2; Biolegend) and anti-CD83 (clone HB15a; Santa Cruz Bio-technology, Santa Cruz, CA) Abs. The expression of NK cell receptorswas analyzed using anti-CD16 (IgG1; clone C127), anti-NKp44 (IgG1;clone Z231), anti-NKp46 (IgG1; clone BAB281), anti-NKG2D (IgG1;clone ON72), and anti-NKG2A (IgG1; Z270), which were kindly providedby Prof. A. Moretta, and anti-NKp30 (IgG1; clone AZ20) was generated inlaboratory. TNF-a and IFN-g were stained by intracellular staining usinganti–TNF-a-PE and anti–IFN-g-PE Abs (Miltenyi Biotec). Before intra-cellular staining, NK cells and DCs were cocultured for 4 h followed by6 h in the presence of 2 mg/ml brefeldin A (Sigma-Aldrich). Cells wereanalyzed with a Cyflow Space cytometer (Partec, Munster, Germany). InNK–DC cocultures, forward and side parameters were used to distinguishviable DCs from NK cells or dead DCs. DC survival was calculated as thepercentage of viable DCs cocultured with NK cells, relative to DCs cul-tured alone. CD56bright and CD56dim NK cell subsets were distinguishedusing PC5-conjugated anti-CD56 Abs (Immunotech, Marseille, France).

Cell conjugate formation assay

NK–DC conjugates were detected by flow cytometry using NK cells la-beled with 10 mM CFSE (Sigma-Aldrich) and MoDCs labeled with 4 mMPKH26 (Sigma-Aldrich). After cell labeling, the cells were centrifuged at200 3 g for 2 min at room temperature and incubated at the NK/DC ratiosof 1:1 at 37˚C for 30 min. The percentage of conjugates formed wascalculated by determining the percentage of the double-positive populationusing a Cyflow Space cytometer (Partec).

Statistical analysis

Statistical significance between the experimental groups was calculatedwith Prism 4 (GraphPad) using paired Student t test.

ResultsActivin A is upregulated in NK–DC cocultures

A PCR-based array screening was performed to investigate thecytokine expression profile of MoDCs cocultured with allogeneicNK cells preactivated with IL-15 (1:1 NK/DC ratio). The genes of 22cytokines of 84 were upregulated.4-fold during NK–DC coculture.Most of the cytokines were proinflammatory, such as IL-8, IL-1a,IL-1b, and TNF-a, and were rapidly upregulated at 2 h coculture.Interestingly, also a number of cytokines (e.g., IL-3, IL-7, IL-15,GM-CSF, and M-CSF), known to be involved in the regulation ofhematopoiesis and cell proliferation, were rapidly induced. Strik-ingly, the second most upregulated gene (40-fold) was inhba, a genethat encodes for the bA subunit of activin A, activin AB, and inhibinA (Fig. 1A, Supplemental Fig. 1; accession GSE52306, http://www.ncbi.nlm.nih.gov/geo). Quantitative RT-PCR confirmed the induc-tion of the activin bA subunit mRNA starting at 1 h and reachinga peak after 6 h of coculture with a kinetics that was delayed withrespect of those of proinflammatory cytokines, such as TNF-a andIFN-g (Fig. 1B). Neither the a-subunit of inhibin A nor the bB-subunit of Activin AB was upregulated (data not shown), suggestingan exclusive induction of activin A.TGF-b family members are known to be implicated in the

regulation of immune responses (27, 28). In our experimentalconditions, the mRNA of other TGF-b family members weremodestly upregulated, namely, TGF-b2 (8-fold; 6 h) and BMP-6(4-fold; 2 h; Supplemental Fig. 1). TGF-b1 mRNA was consti-tutively expressed, although no transcriptional regulation duringNK–DC interactions occurred (Fig. 1B). Protein quantificationrevealed negligible levels of TGF-b1 and TGF-b2 (138 6 111 and53 6 80 pg/ml; n = 3), whereas BMP-6 protein expression did notreach the ELISA detection limit, possibly reflecting the low levelsof mRNA transcription (data not shown). In contrast, high levelsof activin A (2.28 6 1.24, range 0.77–4.61 ng/ml; n = 23) weredetected after 24 h of coculture of DCs with activated NK cells,whereas lower levels of activin A (0.67 6 0.17 ng/ml) were foundwhen resting NK cells were used in cocultures (Fig. 1C). Similar

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levels of activin A production were reached using autologous andallogeneic culture conditions, although in the autologous setting,activin A protein and mRNA production linearly increased withNK/DC ratios (up to 10:1), whereas in allogeneic cultures, activinA production peaked at the 1:1 NK/DC ratio and then declined(Fig. 1C and data not shown). Autologous cocultures acquired theexpression pattern of allogeneic cocultures when HLA class Imolecules were blocked by mAbs, suggesting a negative regulationof activin A production by HLA-I molecules (data not shown).

DCs cultured alone produced low basal levels of activin A (0.3960.19 ng/ml; n = 25). In contrast, no activin A was detected inpurified NK cells at resting or activated conditions (i.e., after IL-2,IL-12, IL-15, IL-18, anti-CD16, anti-NKp30, or PMA/ionomycinstimulation; Supplemental Fig. 2). In addition, no mRNA expres-sion of activin bA subunit was found in NK cells that had beenmagnetically purified after coculture for 6 h with DCs, whereasactivin bA expression was strongly detected in DCs (Fig. 1D). Theseresults demonstrate that during NK–DC cocultures, activin A is

FIGURE 1. Activin A is strongly upregulated in NK–DC cocultures. (A) Immature MoDCs were cocultured with allogeneic IL-15–activated NK cells at

a NK/DC ratio of 1:1. The RNA (pooled from three independent experiments) was quantified by an RT-PCR array including 84 common cytokine genes.

The scatter plot shows gene expression at 6 h of coculture, compared with the gene expression at time 0. The genes with a fold change of .4 or ,24 were

considered as upregulated or downregulated, respectively. PCR array reproducibility was validated using RT2 Profiler PCR Analysis software (Qiagen). The

calculated SD for obtained Ct values was 61.3%. (B) Changes of mRNA of the activin bA subunit, TNF-a, IFN-g, and TGF-b1 were quantified during

cocultures of immature MoDCs and allogeneic activated NK cells at a NK/DC ratio of 1:1 by RT-PCR. *p # 0.05 (vs. control 0 h); n = 3. (C) Activin A

expression was quantified by ELISA after coculture of allogeneic or autologous immature MoDCs with resting or activated NK cells at the indicated ratios

for 24 h. *p# 0.05 (versus DC alone); n$ 4. (D) Activated NK cells were cocultured with MoDCs at a NK/DC ratio of 1:1 for 2 and 6 h. NK cells (CD56+)

and DCs were then separated by magnetic bead separation. The RNA of the activin bA subunit was quantified by RT-PCR. *p # 0.05 (versus control 0 h);

n = 3. (E) CD1c+ DCs, slanDCs, CD141+ DCs, and pDCs were cocultured with allogeneic activated NK cells at the indicated NK/DC ratio for 24 h. Activin

A expression was quantified by ELISA. *p # 0.05 (versus DC alone); n = 3.

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exclusively produced by DCs, confirming and extending previ-ous findings obtained with cytokine-activated NK cells (25).DC subsets, directly isolated from blood, differed in their ca-

pacity to produce activin A upon the encounter with NK cells (Fig.1E). Although pDCs were not able to produce activin A, all my-eloid DC subsets showed significant upregulation of activin Awhen cocultured with activated NK cells, although with differentexpression levels. Of all myeloid DC subsets tested, CD1c+ DCsreleased the highest amounts of activin A (4.83 6 0.99 ng/ml; n =3), whereas slanDCs and CD141+ DCs were only poor producersof activin A (0.60 6 0.39 and 0.32 6 0.06 ng/ml; n = 3). Inter-estingly, LPS (100 ng/ml), although a powerful inducer of activinA expression in MoDCs (24, 29), was a weaker inducer of activinA expression in myeloid CD1c+ DCs (1.496 0.22 ng/ml; n = 3) ascompared with NK–DC interaction.

Role of cytokine secretion in activin A production in NK–DCcocultures

To elucidate the molecular basis responsible for activin A pro-duction in NK–DC cocultures, we initially focused on the role ofinflammatory cytokines, such as IL-1b, TNF-a, IFN-g, and GM-CSF, cytokines known to be upregulated in our coculture con-ditions (Fig. 1A, Supplemental Fig. 1) and known to be able toinduce activin A in DCs (24, 29). Fig. 2 shows that the addition ofblocking mAbs for each of the mentioned cytokines, used at activeblocking concentrations (data not shown), was able to decreaseactivin A production, with anti–TNF-a being the most active one.The simultaneous neutralization of the four cytokines resulted inthe nearly complete abrogation of activin A production, suggest-ing a role for multiple cytokines. In contrast, cytokine blockingmAbs did not have any effect on basal production of activin Aby DCs (data not shown). Separation of NK cells and DCs bytranswell inserts abrogated activin A expression; this result is inagreement with the known contact-dependent production of thesecytokines in NK–DC cocultures (3, 12).

Role of NK cell–mediated killing of DCs in activin Aproduction

It was previously described that NK cells are able to kill immatureDCs in vitro at high NK/DC cell ratios (3). As shown in Fig. 3A,activin A production was directly associated with NK cell–me-diated killing of DCs, at least up to a 5:1 NK/DC cell ratio (inautologous cocultures); thereafter, activin A production did notfurther increase possibly because of the low number of survivingDCs. The correlation (R2 = 0.63; allogeneic cocultures; NK/DCratio = 1:1) between cell death and activin A production promptedus to evaluate the possible relationship between the two events. To

FIGURE 2. Role of cytokine secretion in activin A expression in NK–

DC cocultures. Immature MoDCs and activated NK cells were cocultured

in the presence of neutralizing concentrations of Abs specific for TNF-a,

GM-CSF, and IFN-g and IL-1b for 24 h. Activin A expression was

quantified by ELISA. The results were normalized to the amount of activin

A induced in cocultures with each isotype-specific control Ab (100%). The

group labeled “DC” represents the basal production of activin A by DCs

cultured alone. In some experiments, NK cells and DCs were cocultured

separated by transwell inserts. ns, not significant. *p # 0.05; n $ 3.

FIGURE 3. Induction of activin A by dead DCs and K562 cells. (A)

Viability of MoDCs was quantified after coculture with autologous NK

cells at the indicated NK/DC ratios after 24 h of coculture. Activin A

concentration was evaluated in the supernatants by ELISA. Data represent

means 6 SD of triplicates of one representative experiment of three per-

formed. *p # 0.05 (versus DC alone). (B) MoDCs (105 cells) were incubated

together with dead DCs at the indicated ratios. Data represent means 6 SD

of triplicates of one experiment representative of three. (C) To demonstrate

the synergistic effect of dead DCs, we added low amounts of dead DCs

(alive/dead DC ratio = 10:1) to DCs alone (105 cells) or NK–DC cocultures

using the suboptimal 1:5 NK/DC ratio. Data represent means 6 SD of

triplicates of one representative experiment of three performed (*p # 0.05).

(D) MoDCs were cocultured with NK cells isolated from a healthy donor or

from NK cells of two patients with HPS-2. After 24 h, the concentration of

activin A in the supernatant was quantified by ELISA. The results obtained

with two different DC donors are shown. (E) MoDCs (105 cells) were in-

cubated with dead K562 cells at the indicated ratios. Data represent means6SD of triplicates of one experiment representative of three (*p # 0.05). (F)

Alive K562 cells (104 cells) were cultured together with NK cells (105 cells),

MoDCs (105 cells), or autologous NK–DC cocultures (105 cells each). After

24 h, the concentration of activin A in the supernatants was quantified by

ELISA. Data represent means 6 SD of triplicates of one experiment repre-

sentative of three (*p # 0.05).

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this goal, we cultured DCs in the presence of dead DCs and an-alyzed them for released activin A. Activin A expression sig-nificantly increased in the presence of dead DCs, obtained byrepeated freezing and thawing (but not by UV irradiation), al-though only at relatively high ratios (Fig. 3B and data not shown).However, the addition of low concentrations of dead DCs to NK–DCs cocultured at the suboptimal NK/DC ratio of 1:5 (charac-terized by no NK cell–induced killing and negligible expression ofactivin A) caused a significant induction of activin A (Fig. 3C).To further support the role of NK cell–mediated killing of DCs inNK–DC cross-talk, we performed experiments using cultures ofNK cells originally obtained from peripheral blood of HPS-2patients; these patients have a genetic defect associated with areduced cytotoxic activity (30). NK cell lines obtained from twoHPS-2 patients showed significantly lower cytotoxicity againstDCs during NK–DC cocultures (data not shown) and a reducedproduction of activin A (Fig. 3D). Notably, TNF-a production alsowas strongly reduced in these experimental conditions (data notshown), confirming the importance of TNF-a in activin A in-duction. Importantly, conjugate formation assays revealed that NKcells from HPS-2 patients could bind and form conjugates withDCs with an efficiency equal to that of NK cells isolated fromhealthy donors (Supplemental Fig. 3). Similarly, lysates and viableK562 tumor cells, classical NK cell targets, were able to induceactivin A expression in DC cultures (Fig. 3E) and in autologousNK–DC cocultures (Fig. 3F), respectively.

Activin A inhibits cytokine expression and NK cell–mediatedDC maturation

Previous studies have shown that cytokines produced during NK–DC interaction affect DC maturation and NK cell functions (4,6, 11, 16). Recent studies described a negative role for activin Aon cytokine production by activated NK cells and DCs stimulatedwith CD40L, but not with LPS (24, 25). To investigate the pos-sible role of endogenously produced activin A during NK–DCinteractions, follistatin, a natural occurring high-affinity ligandthat binds in a stoichiometric manner to activin A (31), was usedto neutralize endogenous activin A activity. The addition of fol-listatin to NK–DC cocultures significantly upregulated the ex-pression of TNF-a, IL-6, and IL-8, but did not change the levelsof IL-1b, and caused only a nonstatistically significant increaseof the NK cell–derived cytokines IFN-g and GM-CSF (Fig. 4A).Addition of follistatin had no effect on basal DC and NK cellcytokine expression. Similar results were obtained with SB431542,an inhibitor of the activin receptor (Alk4) signaling, and with ananti-activin A blocking polyclonal Ab. Conversely, dorsomor-phin, an inhibitor of BMP receptor signaling, did not affect cy-tokine expression in NK–DC cocultures (Fig. 4B and data notshown). To evaluate the possible effect of endogenous activin Aon DC viability, DC survival was determined at the end of 24 hincubation with follistatin alone or in the presence of NK cells. Inboth conditions, no difference in DC viability was observed,suggesting that endogenously produced activin A did not influ-ence DC survival and NK cell cytotoxic activity (SupplementalFig. 4A).Finally, we analyzed the influence of activin A on NK cell–

mediated DC maturation. Fig. 5 shows a significantly higher up-regulation of CD86 and CD83 on DCs cocultured with NK cellsin the presence of follistatin. Conversely, follistatin treatment did notproduce any effect on DCs cultured alone (Fig. 5), or on the ex-pression of NK cell receptors (NKp30, NKp40, NKp46, NKG2D,CD16, and NKG2A) in NK–DC cocultures (Supplemental Fig.4B). In conclusion, these results provide evidence that the en-dogenous production of activin A in NK–DC cocultures acts as

a negative feedback mechanism of cytokine production and ofDC maturation.

Endogenous activin A exclusively inhibits DC cytokineexpression

To discriminate the effect of activin A on DCs and on NK cells,TNF-a and IFN-g production was analyzed by intracellular stainingin coculture experiments performed in the presence of follistatin.Although both NK cell subsets produced similar levels of TNF-a,IFN-g was higher expressed by CD56bright NK cells. However,neither the CD56bright nor the CD56dim subsets showed any sig-nificant increase in the expression of activin A in the presence offollistatin. On the contrary, DCs showed significantly increased

FIGURE 4. Inhibitory role of activin A on cytokine production in NK–

DC cocultures. (A) Activated NK cells and MoDCs were cocultured at the

1:1 NK/DC ratio in the presence of 200 ng/ml follistatin for 24 h. Cytokine

expression was quantified by ELISA. *p , 0.05; n $ 3. (B) To confirm the

results obtained with follistatin, we cocultured cells in the presence of the

Alk4 inhibitor SB431542 (10 mM), a polyclonal anti–activin A Ab (10 mg/

ml), or Dorsomorphin (5 mM), an inhibitor of BMP receptor signaling.

TNF-a and IL-6 expression were quantified after 24 h. The correspondent

controls (DMSO and irrelevant Ab) did not alter TNF-a and IL-6 ex-

pression. ND, not detected. *p # 0.05; n = 3.

FIGURE 5. Role of activin A production on DC maturation. Allogeneic

immature MoDCs and activated NK cells were cocultured at the 1:1

NK/DC ratio in the presence of 200 ng/ml follistatin for 24 h. The expression

of CD86 and CD83 on DCs was analyzed by flow cytometry. Statistical

analysis of the percentage of positive cells cultured in the absence (white

bars) or presence (black bars) of follistatin are shown. *p # 0.05; n $ 3.

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expression of TNF-a, but not of IFN-g, upon inhibition of en-dogenous activin A (Fig. 6A). The inhibitory effect of activin Awas further analyzed as mRNA cytokine levels in NK cells andDCs, separated after coculture. Once again, NK cells did not up-regulate any of the cytokines tested, whereas DCs showed signifi-cantly higher levels of TNF-a, IL-6, and IL-8 (Fig. 6B).

DiscussionFine-tuning of the immune response represents the key mechanismto ensure proper activation of protective immunity. The excessiveactivation of immune effector cells may result in host detrimentalconditions, such as in chronic inflammation and autoimmunity. Toensure the proper regulation of effector mechanisms, the pro-duction of proinflammatory molecules is always associated withthe upregulation of secreted and membrane-bound inhibitorypathways (32).In the past 15 years, it has become evident that NK cells

cooperate with DCs in the regulation of innate immunity and inshaping adaptive immune response. In this interplay, NK cellspromote DC maturation and “edit” those cells that have notreached complete activation. At the same time, DCs promoteNK cell activation and function through the secretion of activatingcytokines. This complex cell–cell interaction relies on the pro-duction of soluble and membrane-bound cytokines (e.g., TNF, IL-12, IL-15, and IL-18) and on the interaction of specific NK cellreceptors (e.g., NKp30 and NKG2A) with their cognate ligandsexpressed by DCs (13, 14). Recently, activin A, a member of theTGF-b cytokine family, has been suggested to negatively regulateNK and DC functions (24, 25). This study further extends thishypothesis, providing for the first time, to our knowledge, evi-dence that activin A represents one of the main cytokines inducedduring NK–DC cocultures and represents a main negative feed-back mechanism of this interaction.Activin A was identified by low-density macroarray analysis as

the second highest induced cytokine in NK–DC cocultures. Theexpression of activin A was rapid, but delayed with respect toother immune-activating cytokines, such as TNF-a and IFN-g. Ofthe two cell types, only DC was responsible for activin A pro-duction, with no contribution from NK cells. This is consistentwith previous finding reporting that DCs are potent producers ofactivin A when activated with TLR ligands or upon CD40 trig-gering (24, 29), and that NK cells do not produce activin A upon

stimulation (25). Importantly, besides MoDCs, also CD1c+, CD141+,and slanDC subpopulations expressed activin A upon NK cell en-counter, although at dissimilar expression levels. During NK–DCinteraction, the production of activin A was dependent on the pres-ence of cytokines, namely, TNF-a, IFN-g, and GM-CSF. Dataavailable in literature, as well as our own observations, suggest thatduring their interaction, both NK cells and DCs produce TNF-a,whereas only NK cells are responsible for the IFN-g and GM-CSFrelease (3, 16).Our study also reports that activin A expression is potentiated by

NK cell–mediated killing of DCs. This conclusion is supported bythe observation that the highest degree of activin A induction wasdetected at an elevated NK/DC ratio, a condition known to beassociated to NK cell–mediated DC death (3). Further support forour hypothesis came from experiments performed using cells fromHPS-2 patients with a genetic defect known to be associated witha reduced cytotoxic activity of NK cells (30). Killing of immatureDCs by NK cells was suggested to represent an “editing” step forthe quality control of DCs undergoing maturation (13), althoughthe in vivo relevance of this process still needs to be fully clarified.Multiple mediators are known to be released by dead cells, such asheat shock proteins, HMGB1, and other alarmins (18, 33, 34). Inpreliminary results, we have observed that HMGB1 is able toinduce TNF-a and activin A in DCs, but the exact mechanism bywhich dead DCs induce activin A is still under investigation. Thebiological relevance of the recognition and uptake of dead cellsby DCs is well documented. Hence necrotic cells are known toupregulate TNF-a and CD86 in DCs (4, 35, 36), and in vivoexperiments have shown that the uptake of dead tumor cellspromotes DC maturation and tumor Ag cross-presentation (37).Our data show that the recognition of NK cell–killed tumor cellsby DCs also promotes activin A production. Because activin Areleased within the tumor microenvironment was described todirectly promote tumor growth, it is tempting to speculate thatactivin A may be involved in the establishment of tumor-mediatedimmunosuppression (38). Indeed, inhibition of endogenous activinA resulted in the upregulation of NK cell–induced expression ofDC maturation markers, such as CD86 and CD83, proposing anegative role for activin A on DC maturation. NK cell–inducedDC maturation was shown to be dependent on the release ofproinflammatory cytokines, such as TNF-a (16). In keeping withthese observations, a significant upregulation of TNF-a, IL-6, and

FIGURE 6. Distinct effect of activin A on DCs and

NK cells. (A) MoDCs were cocultured with allogeneic

activated NK cells at a NK/DC ratio of 1:1 in the

presence of follistatin for 4 h. After addition of brefeldin

A (2 mg/ml), cells were cocultured for an additional 6 h.

CD56bright and CD56dim NK cell subsets and DCs were

gated based on CD56 positivity and side scatter. Intra-

cellular TNF-a and IFN-g were quantified for each cell

type as mean fluorescence intensity (MFI). The results

were normalized to the MFI of DCs in control cocul-

tures. *p # 0.05; n = 3. (B) Activated NK cells were

cocultured with MoDCs at a NK/DC ratio of 1:1 in the

presence of follistatin (200 ng/ml) for 6 h. NK cells

(CD56+) and DCs were then separated by magnetic

bead separation. The RNA of the relative mRNA was

quantified by RT-PCR. The graph shows the media of

percentage changes (follistatin-treated versus untreated

cocultures) of three independent experiments. *p #

0.05 (versus untreated control); n = 3.

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IL-8 release was detected in the presence of follistatin, a naturallyactivin A inhibitor. However, activin A selectively inhibited cyto-kines production by DCs, but not by NK cells. This is apparentlyin contradiction with previously published data describing theinhibitory effect of activin A on IFN-g production by NK cellscocultured with DCs (25). The different results are likely to be dueto the different experimental settings, because Robson et al. (25)performed their experiments in the presence of polyI:C and IL-12to stimulate activin A and IFN-g production. On the contrary, inthis study, we focused on the interactions of DCs and activatedNK cells without the addition of cytokines during the cocultures.Interestingly, it had been described that the ability of activin A to

downregulate cytokine expression was selectively restricted to theCD40-CD40L activation pathway, with no effect in LPS-stimulatedDCs (24). However, during the NK–DC cross-talk, CD40–CD40Linteraction seems to have a negligible role and neither activin Anor TNF-a expression was affected in the presence of a CD40L-blocking mAb (data not shown).It is likely that activin A produced during NK–DC interaction

may also act in a paracrine manner. In lymph nodes, NK cells andDCs reside in the outer paracortex region, where they colocalizeand influence the differentiation of T lymphocytes (6, 11). ActivinAwas reported to favor the differentiation of T cells versus a type 2or regulatory T cell phenotype (22, 39, 40). In addition, the inhi-bition of NK cell–mediated DC maturation by activin A mayfurther contribute to orient adaptive immune response toward atype 2 or Treg phenotype (41, 42). Of note, NK–DC interaction isknown to orient T cell response toward a Th1 phenotype throughthe production of IFN-g (43). The release of activin A by DCs maythus represent a strategy to counterbalance Th1-biased T cell po-larization. Finally, activin A released during NK–DC interactionsmay contribute to the increased local and systemic levels of thesecytokines detected in many acute and chronic inflammatory dis-eases, such as lichen planus or rheumatoid arthritis, characterizedby the concomitant recruitment of these two cell types (20, 44–47).In conclusion, our results provide direct evidence that activin A,

produced during NK–DC interactions, acts as a negative feedbackregulatory pathway to limit excessive DC effector functions andcontribute to the so far limited characterization of the inhibitorymechanisms that regulate NK–DC interactions. In addition, ourresults support a complementary role of inhibitory proteins duringNK–DC cross-talk, with inhibitory NK cell receptors (e.g., killerIg-like receptors and CD94/NKG2A) hindering the initiation ofNK–DC interactions and activin A production acting with a moredelayed kinetics aimed at controlling ongoing interactions andlimiting excessive immune activation.

AcknowledgmentsWe thank Claudia Carlino and Giovanna Tabellini for technical support and

TizianaMusso, Claudia Cantoni, andMassimoVitale for discussion of the data.

DisclosuresThe authors have no financial conflicts of interest.

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