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The Journal of Immunology, 2007, 179, 1711-1720
Copyright © 2007 by The American Association of Immunologists, Inc.
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Triggering of Dendritic Cell Responses after Exposure to Activated, but Not Resting, Apoptotic PBMCs1

Ulrika Johansson2, Lilian Walther-Jallow, Anna Smed-Sörensen and Anna-Lena Spetz

Department of Medicine, Center for Infectious Medicine, Karolinska Institutet, Karolinska University Hospital Huddinge, Stockholm, Sweden


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Dendritic cells (DCs) can be activated by signaling via pathogen receptors, by interaction with activated T cells or by exposure to inflammatory mediators. Clearance of apoptotic cells by DCs is generally considered a silent event that is not associated with an inflammatory response. Necrotic cell death, in contrast, leads to induction of inflammation. However, emerging data challenge the view of apoptotic cells as inherently nonimmunogenic. In this study, we report that the activation state of the apoptotic cell may determine whether the exposed DC becomes activated and rendered proficient in Ag presentation. We show that coculture with activated, but not resting, apoptotic PBMCs leads to up-regulation of surface expression of the costimulatory molecules CD80, CD83, and CD86 in human DCs as well as release of proinflammatory cytokines. Furthermore, we show that DCs exposed to allogeneic, activated apoptotic PBMCs induce proliferation and IFN-{gamma} production in autologous T cells. Together, these findings show that activated apoptotic PBMCs per se provide an activation/maturation signal to DCs, suggesting that activated apoptotic PBMCs possess endogenous adjuvant properties.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Dendritic cells (DCs)3 are potent APCs that have the capacity to stimulate naive Th cells and initiate primary T cell responses. DCs residing in peripheral tissues survey the microenvironment by engulfing both invading microbial material and dying cells of the host. The subsequent outcome of Ag-presentation by DCs depends upon their maturation status. Immature DCs require maturation signals to undergo phenotypic and functional changes to acquire a fully competent Ag-presenting capacity. Maturation of DCs involves several steps such as a transient increased capacity to take up Ag, migration to draining lymph-nodes, and simultaneous up-regulation of molecules including chemokine receptors and costimulatory molecules. Upon challenge with microbial or inflammatory stimuli, DCs mature and gain the capacity to stimulate naive Th cells by presenting the captured Ag on MHC and providing a costimulatory signal via molecules such as CD80 and CD86 to initiate primary T cell responses (1, 2, 3). In addition, priming of Th1 cell responses are dependent on IL-12 production by DCs, initiated via contact with pathogen and further induced via CD40-CD40 ligand (CD40L) interactions (4). Emerging data also support the involvement of an additional signal contributing to the polarization toward Th1 or Th2 responses (5, 6, 7).

DC maturation can be induced by a variety of signals. Among the most efficient are products of microbial origin termed pathogen-associated molecular patterns, which are displayed by pathogens but not normally found on host cells (e.g., LPS, peptidoglycans, nonmethylated CpG, and dsRNA) (8). These molecules are recognized by pattern-recognition receptors and include members of the TLR family (9, 10). Ligation of these receptors can lead to production of proinflammatory cytokines by DCs, such as type I IFNs, TNF{alpha}, and IL-1, which also have been shown to induce DC maturation (7, 11, 12, 13, 14, 15, 16). Some mature DC features may therefore be due to secondary effects mediated by their own cytokine production. However, one report suggests that the inflammatory mediators released after TLR signaling are insufficient to induce full DC activation (7).

A different approach to DC maturation is put forward in the danger hypothesis (17) where it is suggested that APCs are activated by danger/alarm signals from injured cells (17). Hence, injured cells of the host function as endogenous adjuvant that give rise to a danger signal leading to activation of APCs and subsequent stimulation of T cells (11, 18, 19, 20, 21). Apoptotic cells were reported to induce production of anti-inflammatory rather than proinflammatory cytokines in DCs (22, 23, 24, 25). In addition, in vivo experiments have demonstrated tolerance induction by apoptotic cells (26, 27). Yet other studies have conversely shown immunostimulatory effects mediated by apoptotic cells (28, 29, 30, 31, 32, 33). It is not clear why apoptotic cells exhibit such diverse effects in the different studies. However, a recent report by Obeid et al. (34) suggests that the exposure of calreticulin on the apoptotic cells is one factor that dictates their immunogenicity by providing an "eat me" signal. In addition, it was shown that different apoptotic pathways induce differential exposure of calreticulin.

We previously demonstrated that immunization with apoptotic HIV/murine leukemia virus-infected cells induced HIV-1-specific, both cellular and humoral, immune responses in vivo without using any adjuvant (35). In this system, the infected cells were activated before apoptosis induction and administration. The responses elicited in vivo led us to investigate whether the activation state of the apoptotic cells was of importance in achieving the adjuvant effect. In this study, we have set up an in vitro system comparing the potential of resting vs activated apoptotic PBMCs in providing human immature DCs with a maturation signal. We show that activated cells, induced to undergo apoptosis by gamma irradiation, but not resting apoptotic cells, induce expression of costimulatory molecules and release of proinflammatory cytokines in DCs. Furthermore, we show that uptake of allogeneic, activated, apoptotic cells rendered the DCs able to induce proliferation and IFN-{gamma} production in autologous T cells. These findings demonstrate that primary, activated apoptotic cells are able to promote maturation of DCs and may function as endogenous adjuvant for induction of specific T cell responses. These results have implications for the development of vaccines using apoptotic cells for induction of immune responses.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
In vitro differentiation of DCs

CD14+ monocytes were enriched from healthy blood donors by negative selection using RosetteSep Human Monocyte Enrichment (1 ml/10 ml of blood; Stem Cell Technologies) and separated using Lymphoprep density gradients (Nycomed). Cells were cultured for 6 days in complete medium (RPMI 1640 supplemented with 1% HEPES, 2 mM L-glutamine, 1% streptomycin and penicillin, and 10% endotoxin-free FBS; Invitrogen Life Technologies) and recombinant human cytokines IL-4 (6.5 ng/ml; R&D Systems) and GM-CSF (250 ng/ml; PeproTech) to obtain immature DCs (36).

Activation of PBMCs

PBMCs were separated from blood from healthy blood donors using Lymphoprep density gradient (Nycomed). Total T cells were enriched by negative selection using RosetteSep Human CD3+ T Cell Enrichment (1 ml/10 ml of blood; Stem Cell Technologies). Cells were frozen in FBS and 10% DMSO or were directly cultured in complete medium containing 1% sodium pyruvate. PBMCs (106/ml) were activated with PHA (2.5 µg/ml; Sigma-Aldrich) overnight or for 4 days before they were frozen in FBS/DMSO. The monoclonal anti-human CD3 (2 µg/ml, clone OKT 3; Ortho Biotech) was adhered to plastic during 1 h in 4°C before addition of soluble monoclonal anti-human CD28 (2 µg/ml, L293; BD Biosciences) and PBMCs. Additional PBMCs (106/ml) were incubated overnight with soluble monoclonal anti-human CD2 (2 µg/ml, RPA-2.10; BD Biosciences). After overnight incubation, cells were frozen in FBS/DMSO.

Generation of apoptotic cells, necrotic cells, and supernatants

Frozen PBMCs were thawed and washed three times in complete medium. Cells were induced to undergo apoptosis by gamma irradiation (150 Gy). The gamma irradiation-induced apoptotic process has previously been demonstrated by morphological changes, flow cytometry, and DNA fragmentation on agarose gels (37, 38). In this study, apoptosis was confirmed by annexin V (Boehringer Mannheim) and propidium iodide (PI; 0.1 µg/sample, Sigma-Aldrich) stainings according to the manufacturer’s protocol. Necrotic cells were obtained by three rounds of freeze-thawing in culture medium and necrosis was confirmed by annexin V/PI staining. Supernatants were collected from irradiated and freeze-thawed cells after 4, 8, and 24 h and centrifuged at 16,000 x g (Eppendorf Centrifuge 5415D) for 30 min to remove cell debris.

DC/apoptotic PBMC cocultures

On day 6, immature DCs were counted and plated in 24-well plates, 5 x 105 cells in 0.5 ml of medium (complete medium supplemented with recombinant human IL-4 and GM-CSF). Newly irradiated PBMCs, irradiated PBMCs incubated for 24 h, or freeze-thawed PBMCs were added to DCs in a proportion 2:1 in a total volume of 1 ml. A control consisting of DCs cocultured with anti-CD2-treated apoptotic nonactivated PBMCs was also included. Supernatants (0.5 ml) from 106 irradiated PBMCs collected at 4, 8, and 24 h were also added to immature DCs. Supernatants were collected from cocultures at 4, 8, and 24 h. At 72 h, all samples were collected and DCs were phenotyped by flow cytometry. LPS (100 ng/ml; Sigma-Aldrich) was added as a positive control for maturation of DCs. For confocal microscopy analysis and flow cytometry analysis of phagocytosis, PBMCs and DCs were labeled before coculture with green fluorescent dye PKH67 (Sigma-Aldrich) and red fluorescent dye PKH26 (Sigma-Aldrich) or fluorescent dye FarRed (Molecular Probes), respectively. Labeling was performed according to the manufacturer’s protocol. Cytochalasin D (0.5 µg/ml; Sigma-Aldrich) was added to cocultures to inhibit phagocytosis and these cells were used as a negative control. The cells were analyzed by sequential spectrophotometric separation in a Leica SP102 laser scanning confocal microscope or analyzed by using a FACSCalibur flow cytometer (BD Biosciences).

Phenotypic characterization of DCs and PBMCs

DCs were washed and resuspended in PBS with 2% FBS. DCs were incubated for 30 min at 4°C with the following anti-human mAbs: CD1a (clone NA1/34; DakoCytomation), CD14 (clone TÜK4; DakoCytomation), CD19 (clone HD37; DakoCytomation), CD3 (clone SK7), CD80 (clone L307.4), CD83 (clone HB15e), CD86 (clone 2331/FUN-1), and HLA-DR (clone L243; all from BD Biosciences). PBMCs were washed and incubated with anti-human mAbs CD3 (clone SK7), CD4 (clone SK3), CD8 (clone G42-8), CD154 (clone TRAP-1), CD25 (clone 2A3), and CD69 (FN50; all from BD Biosciences). Cell surface expression was measured by flow cytometry and at least 105 cells/sample were collected. After 72 h of coculture, the cells were washed and stained as described above. For analysis of DCs, gates were set on CD4–/CD8– and CD1a+, or CD3–, CD1a+ cells.

Cytokine/chemokine production

Supernatants from cocultures or from irradiated or freeze-thawed PBMCs alone were analyzed for cytokine/chemokine content using a Bio-Plex assay (BioSource International). The assay was performed according to the manufacturer’s protocol and a Luminex reader was used to simultaneously quantify the concentrations of IL-6, IL-8, IL-2, IL-10, IL-12p70, TNF-{alpha}, IFN-{gamma}, and MIP-1beta in the supernatants.

Autologous T cell proliferation and activation

Immature DCs and CD3+ T cells were isolated/differentiated from the same donor as described above. T cells were frozen in 10% DMSO. On day 6 of DC culture, DC/apoptotic cell cocultures were set up as described above and incubated for 48 h. After DC/apoptotic cell coincubation for 48 h, autologous T cells were thawed, washed three times in PBS, and labeled with CFSE as described previously (39). T cells (1.5 x 106) were added to the autologous DCs in a 5:1 ratio or to controls containing activated apoptotic or necrotic cells only in a total volume of 1.5 ml. In positive controls, staphylococcal enterotoxin B (SEB, 5 µg/ml; Sigma-Aldrich) was added. These cultures were incubated for 3, 4, 5, or 6 days. Brefeldin A (10 µg/ml; Sigma-Aldrich) was added to cultures 12 h before staining for surface markers CD1a and CD3 and intracellular IFN-{gamma} (clone 25723.11; BD Biosciences). Cells were first incubated with mAb directed against cell surface markers as described above. For intracellular staining, cells were subsequently fixed in 2% formaldehyde, washed in saponin buffer consisting of 2% FBS, 2% HEPES, and 0.1% saponin in PBS, and incubated with Abs directed against the intracellular Ags at 4°C for 30 min. Cells were finally washed in saponin buffer and analyzed by FACS for cell surface expression, proliferation, and IFN-{gamma} expression. Gates were set on CD1a–CD3+ cells to analyze T cells.

Statistical analysis

Statistical significance was assessed using the Mann-Whitney test and differences were considered significant at p ≤ 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Immature, monocyte-derived DCs ingest apoptotic PBMCs

Human monocytes were cultured for 6 days in the presence of IL-4 and GM-CSF to obtain immature DCs as defined by expression of CD1a, lack of CD14, and low expression of the costimulatory molecules CD80, CD83, and CD86 (36). We first determined whether the monocyte-derived, immature DCs had the capacity to ingest apoptotic cells. PKH26 (red)-labeled immature DCs were cocultured with nonactivated PKH67 (green)-labeled apoptotic PBMCs. Confocal microscopy analyses were performed after 1, 4, or 24 h of coculture. We could not detect uptake of apoptotic cells after 1 h (data not shown). However, after 4 h of coculture, colocalization of DCs and apoptotic cells was observed and apoptotic bodies could be visualized inside DCs (Fig. 1, a and b). After 24 h of incubation the intensity of the colocalization increased and the DCs were enlarged, showing an increased uptake (Fig. 1c). At this time point we could not observe intact apoptotic bodies inside the DCs. Cytochalasin D, which interferes with the phagocytic process by disruption of actin filaments (40, 41), was added to DCs along with the irradiated PBMCs and used as a negative control for uptake of apoptotic cells by the DCs. DCs were harvested after 24 h and very few double-positive cells were detected in the cultures containing cytochalasin D (Fig. 1d). This suggests that uptake of apoptotic PBMCs occurs via a phagocytic pathway since inhibition of actin filaments with cytochalasin D interferes with phagocytosis but leaves endocytic capacity intact (41). Uptake was quantified in cocultures with FarRed-labeled DCs and PKH67-labeled apoptotic PBMCs after 24 h using flow cytometry (Fig. 1e). FarRed/PKH67 double-positive cells can be either DCs that have taken up or bound apoptotic cells. Treatment with cytochalasin D reveals the frequency of DCs that bind apoptotic PBMCs without ingestion. The flow cytometry gates were set based on DC/apoptotic PBMC cocultures treated with both cytochalasin D and trypsin because this treatment abrogates uptake and removes bound apoptotic cells from the cell surface. The frequency of apoptotic cells taken up by DCs did not differ when comparing nonactivated apoptotic PBMCs (mean, 26.6%) and anti-CD3/CD28 activated apoptotic PBMCs (mean, 28.6%) (Fig. 1e). However, there was a higher frequency of DCs that bound activated apoptotic cells compared with nonactivated apoptotic cells. Flow cytometry data were confirmed by confocal microscopy (data not shown). These results show that human monocyte-derived, immature DCs have the ability to phagocytose gamma-irradiated nonactivated and activated PBMCs.


Figure 1
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  		FIGURE 1. Human monocyte-derived DCs ingest apoptotic PBMCs. Immature monocyte-derived DCs were labeled with PKH26 (red; a–d) or with FarRed (e). PBMCs were labeled with PKH67 (green; a–e) and thereafter induced to undergo apoptosis by gamma irradiation. The confocal images show DCs cocultured with nonactivated apoptotic PBMCs. After 4 h, DCs that had engulfed apoptotic cells (ac) appeared yellow, as exemplified by the cell highlighted with an arrow (a). Higher magnification of an apoptotic body within a DC after 4 h of coculture is shown in b. After 24 h of culture, a high frequency of the DCs had taken up ac (c). As a control, cytochalasin D (cyto.D) was added to the cocultures to block phagocytosis and uptake of ac. Inhibition of phagocytosis resulted in lack of colocalization of DCs and ac after 24 h (d). DC uptake of nonactivated and anti-CD3/CD28-activated ac was quantified after 24 h of coculture using flow cytometry (e). True uptake was considered as the difference between the total frequency of PKH67/FarRed double-positive DCs (dark bars) and the frequency of PKH67/FarRed double-positive DCs in samples where cytochalasin D was added to abrogate uptake (bright bars). The mean uptake of nonactivated ac was 26.6% and mean uptake of anti-CD3/CD28-activated ac was 28.6%. Confocal images show representative results from one experiment of four. Phagocytosis frequency is shown for three donors in one representative experiment of three.

 
Activated, but not resting, apoptotic PBMCs induce expression of costimulatory molecules in DCs

To investigate whether activated apoptotic primary PBMCs had the capacity to provide maturation signals to DCs, we polyclonally activated T cells present in the PBMCs using PHA or anti-CD3/CD28 mAbs. We determined the efficiency of T cell activation by analyzing induction of CD25 and CD69 expression on the T cells after stimulation (Fig. 2). Both PHA and anti-CD3/CD28 activation resulted in up-regulation of CD25 and CD69. Similar high frequencies of CD25- and CD69-positive T cells were obtained after PHA and anti-CD3/CD28 stimulation. T cells were also stained for CD40L expression as CD40-CD40L interactions can induce DC maturation. CD40L expression could be detected on purified, activated T cells, but not in the T cell population present in PBMCs (data not shown). This could be due to B cell-mediated endocytosis of CD40L on activated T cells (42, 43). We used gamma irradiation to induce apoptosis as previously described (35, 37, 38). Nonactivated and activated PBMC preparations were irradiated or freeze-thawed and apoptosis and necrosis induction was determined by annexin V and PI staining followed by flow cytometry analysis (Fig. 3). Both nonactivated and activated PBMCs contained cells in early apoptosis and secondary necrosis after gamma irradiation as measured by the annexin V/PI staining. Twenty-four hours after gamma irradiation and after three rounds of freeze-thawing, the majority of cells were double positive for annexin V and PI, indicating that the gamma irradiation effectively induced apoptotic cell death in both resting and activated PBMCs with similar kinetics and that necrosis induction by freeze-thawing was effective.


Figure 2
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  		FIGURE 2. Characterization of activated PBMCs. Human PBMCs were activated with PHA overnight (a and d) or for 4 days (b and e) or were treated with anti-CD3 and anti-CD28 Abs overnight (c and f). Nonactivated and activated PBMCs were stained for T cell activation markers CD25 and CD69. Samples were analyzed by flow cytometry and gates were set on lymphocytes based on forward scatter and side scatter profiles. The bars and numbers show the frequency of positive cells. The stainings show up-regulation of CD25 and CD69 in Ab- and PHA-stimulated PBMCs (black line) when compared with nonactivated cells (gray line). Representative data from 1 experiment of at least 10 performed.

 

Figure 3
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  		FIGURE 3. Apoptosis induction in resting and activated PBMCs. Nonactivated (a–d) and anti-CD3/anti-CD28-activated (e–h) PBMCs were stained with annexin V and PI before gamma irradiation (a and e) and 6 h (b and f) or 24 h (c and g) after irradiation or after three rounds of freeze-thawing (d and h) to determine the frequency of apoptotic and necrotic cells (nc) in the populations. Samples were analyzed by flow cytometry and the total PBMC population was included in the analysis. Both in resting and in activated cells, the frequency of apoptotic (annexin V+PI–) and necrotic (annexin V+PI+) cells increased similarly after gamma irradiation. The vast majority of resting and activated cells subjected to freeze-thawing were necrotic (annexin V+ PI+). Representative data from 1 experiment of at least 10 performed is shown.

 
We added newly irradiated apoptotic PBMCs to immature DCs and cultured them for 72 h before analyses of phenotypic changes associated with DC maturation. To exclude the possibility that activation of DCs occurred via FcR-mediated Ab signaling as a result of residual anti-CD3 and anti-CD28 mAbs in the apoptotic cell preparations, anti-CD3 and anti-CD28 Abs were added in control DC cultures. In addition, FcR-mediated DC signaling events were taken into account by adding anti-CD2-opsonized apoptotic PBMCs. Cells were collected and stained for CD1a, CD80, CD83, CD86, and HLA-DR and analyzed by flow cytometry. Mature DCs were defined as CD1a+ cells with distinct, high expression of CD86. Quadrants were set based on negative controls (medium) and positive controls (LPS) (Fig. 4a). There was a significant increase in the frequency of CD86-expressing DCs in cocultures containing activated PBMCs when compared with the medium control. Resting apoptotic cells, anti-CD3/CD28 Abs alone, or anti-CD2-treated apoptotic cells did not induce up-regulation of CD86 expression in DCs (Fig. 4b). A tendency toward a stronger induction of CD86 using anti-CD3/anti-CD28-activated apoptotic cells was observed compared with PHA-activated apoptotic cells. For this reason, we used anti-CD3/CD28 mAbs to activate PBMCs in the majority of subsequent experiments. Mean fluorescence intensity values for CD80, CD83, CD86, and HLA-DR expression on DCs cocultured with nonactivated or anti-CD3/CD28-activated apoptotic cells were compared with the medium control (Fig. 4c). The expression of CD80, CD83, and CD86 molecules was up-regulated in DCs cocultured with activated apoptotic cells, while HLA-DR expression did not differ significantly from the medium control. Purified, anti-CD3/anti-CD28-activated, apoptotic CD4+ T cells were also able to induce expression of costimulatory molecules in DCs (data not shown). These results show that activated, but not resting, apoptotic PBMCs are potent inducers of DC maturation as defined by up-regulation of costimulatory molecules.


Figure 4
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  		FIGURE 4. Activated, apoptotic PBMC induce maturation in human monocyte-derived DCs. DCs were cocultured with ac derived from nonactivated PBMCs (nonact. ac), PHA-activated PBMCs stimulated overnight (PHA o.n. ac) or for 4 days (PHA 4d ac), or finally with apoptotic anti-CD3/CD28-activated PBMCs (anti-CD3/anti-CD28 ac). Control samples included DCs cultured in medium alone, with anti-CD3/CD28 mAbs (ab control), or with ac pretreated with anti-CD2 mAb. LPS was used as a positive control for induction of DC maturation. DCs were cocultured with ac for 72 h and subsequently analyzed by flow cytometry. a, CD86 expression in DC cultures from one representative donor. Gates were set on CD3– and CD1a+ DCs or CD4–CD8– and CD1a+ DCs. b, The frequency of CD86-positive and (c) the mean fluorescence intensity of CD80, CD83, CD86, and HLA-DR expression on DCs were determined in multiple donors after the indicated treatment: n = 22 for medium and LPS (b), n = 18 for nonact ac, n = 16 PHA 4d ac, and anti-CD3/anti-CD28 ac, n = 4 for PHA o.n. ac, and n = 6 for ab control and anti-CD2 ac. c, n = 6 for all samples. Significant up-regulation of costimulatory molecules compared with medium control is indicated as **, p ≤ 0.01 or ***, p ≤ 0.001.

 
Resting, necrotic PBMCs do not induce DC maturation

To examine the possibility that necrotic cells present in the samples after gamma irradiation (Fig. 3) caused maturation of DCs, we compared the state of maturation in DCs cocultured with resting or anti-CD3/CD28-activated, newly gamma-irradiated PBMCs as well as resting or activated necrotic PBMCs. To exclude the possibility that potentially live cells were the cause of DC maturation, we also incubated gamma-irradiated PBMCs for 24 h before addition to DCs. We did not detect significant up-regulation of CD86 expression in DCs cocultured with resting, necrotic, or apoptotic cells, whereas both the newly irradiated activated apoptotic, the 24-h incubated, activated apoptotic, and activated necrotic cells induced significant CD86 expression compared with the medium control (Fig. 5a). Newly irradiated and 24-h incubated, activated apoptotic cells were however more potent inducers of DC maturation as compared to the activated necrotic PBMCs. There was no significant difference between the newly irradiated and the 24-h incubated, activated apoptotic cells in DC maturation potency. As additional controls, we incubated DCs with freshly isolated irradiated PBMCs and freshly isolated or frozen-live PBMCs (Fig. 5b). Nonactivated PBMCs were still unable to induce DC maturation. The freshly isolated, activated irradiated PBMCs induced similar expression of CD86 as frozen cells. Live, activated PBMCs, freshly isolated or frozen, induced expression of CD86 relative to the medium control but were less effective compared with irradiated activated PBMCs. These results demonstrate that necrotic, primary PBMCs do not per se induce DC maturation. In addition, these data show that newly irradiated PBMCs in early apoptosis and PBMCs incubated for 24 h postirradiation in late-stage apoptosis are equally potent in inducing DC maturation. We also conclude that the DMSO-freezing process of PBMCs does not affect their ability to induce DC maturation. Finally, the occurrence of live cells among irradiated PBMCs cannot explain the induced DC maturation because live, activated PBMCs were less efficient compared with apoptotic PBMCs in inducing DC maturation. In summary, the propensity of PBMCs to acquire an immunostimulatory potential requires activation before induction of cell death and the acquired capacity to induce DC maturation remains also in later stages of apoptosis.


Figure 5
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  		FIGURE 5. Resting, necrotic PBMCs are not able to induce DC maturation. a, DCs were cocultured with ac derived from nonactivated PBMCs that were added directly after irradiation (nonact. ac n = 8) or 24 h after irradiation (nonact. ac 24 h, n = 6), anti-CD3/CD28-activated PBMCs added directly after irradiation (anti-CD3/anti-CD28 ac n = 8), or 24 h after irradiation (anti-CD3/anti-CD28 ac 24 h, n = 6), nonactivated necrotic PBMCs (nonact nc, n = 9) or anti-CD3/CD28-activated necrotic PBMCs (anti-CD3/anti-CD28 nc, n = 9). Control samples included DCs cultured in medium alone (n = 9). LPS was used as a positive control for induction of DC maturation (n = 9). b, The capacity of freshly isolated, frozen, live, and apoptotic PBMCs to induce DC maturation was measured. Cocultures consisted of DCs and freshly isolated, newly irradiated cells (nonact. ac fresh and anti-CD3/anti-CD28 ac fresh), frozen newly irradiated cells (nonact. ac frozen and anti-CD3/anti-CD28 ac frozen), freshly isolated live cells (nonact. live fresh and anti-CD3/anti-CD28 live fresh), or frozen live cells (nonact. live frozen and anti-CD3/anti-CD28 live frozen). DCs were cocultured with ac for 72 h before flow cytometric analyses were performed. Gates were set on large, CD1a+ DCs. a, Significant differences are indicated as ***, p ≤ 0.001 compared with medium control. Significant differences between samples are indicated as *, p ≤ 0.05 and **, p ≤ 0,01. b, The graph shows data from three different donors.

 
Supernatants from activated, apoptotic PBMCs do not induce DC maturation

We next investigated whether the up-regulation of costimulatory molecules on DCs after coculture with activated, irradiated PBMCs was due to factors released by the apoptotic cells. Supernatants from anti-CD3/anti-CD28-activated, irradiated PBMCs were therefore collected 4, 8, and 24 h after induction of apoptosis. DCs cocultured with activated PBMCs significantly up-regulated CD86 expression, whereas supernatants from the same apoptotic cell preparations were not effective in inducing CD86 expression (Fig. 6).


Figure 6
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  		FIGURE 6. Supernatants from apoptotic PBMCs do not have the capacity to induce DC maturation. Supernatants from anti-CD3/CD28-activated, irradiated PBMCs (act. ac sup) were collected 4, 8, and 24 h after induction of apoptosis. Supernatants were subsequently added to immature DCs. Simultaneously, nonactivated or anti-CD3/CD28-activated, apoptotic PBMCs from the corresponding donors were added. Cocultures were incubated for 72 h. Cells were then stained for CD86 and analyzed by flow cytometry. In the graph presented, n = 4 for medium control, LPS control, DC/nonactivated ac and DC/supernatant 24 h, n = 5 for DC/anti-CD3/CD28-activated ac and n = 6 for DC/supernatants at 4 and 8 h. Significant differences compared with medium control are indicated as **, p ≤ 0.01 or ***, p ≤ 0.001.

 
Activated, apoptotic PBMCs induce proinflammatory cytokine release in DCs

To further analyze DC activation after addition of activated dying cells, we analyzed the cytokine and chemokine secretion from the DCs after coculture with resting or activated apoptotic or necrotic PBMCs. Immature DCs were cocultured with nonactivated apoptotic PBMCs, apoptotic PBMCs activated with PHA overnight or for 4 days, apoptotic PBMCs activated with anti-CD3/anti-CD28 Abs overnight, nonactivated necrotic PBMCs, or anti-CD3/CD28-activated necrotic PBMCs. Supernatants from the DC/dying cell cocultures were collected after 4, 8, and 24 h. The supernatants were frozen and subsequently analyzed for their content of IL-6, IL-8, IL-2, IL-10, IL-12p70, TNF-{alpha}, IFN-{gamma}, and MIP-1beta. In general, we detected release of IL-6, TNF-{alpha},and MIP-1beta in the cocultures containing DCs and activated apoptotic or necrotic cells (Fig. 7). However, the cytokine levels detected in cocultures containing DCs and activated necrotic cells were lower than concentrations in DC/activated apoptotic cell cocultures. Significantly lower levels of these cytokines were detected in supernatants collected from apoptotic and necrotic cells alone, suggesting production and release from the DCs and not from the apoptotic/necrotic cells per se. In cocultures containing DCs and apoptotic cells activated with PHA for 4 days, we could not detect release of TNF-{alpha}, whereas significant release was detected in cocultures with apoptotic cells activated with PHA or anti-CD3/CD28 overnight. Cocultures containing DCs and activated necrotic cells contained low levels of TNF-{alpha}. Activated necrotic cells released significant levels of MIP-1beta in the supernatant compared to the medium control.


Figure 7
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  		FIGURE 7. Activated apoptotic PBMCs induce proinflammatory cytokine release in DCs. Immature DCs were cocultured with nonactivated ac (nonact. ac), ac activated with PHA overnight (PHA o.n. ac) or for 4 days (PHA 4d), anti-CD3/CD28-activated ac (anti-CD3/anti-CD28 ac), nonactivated nc (nonact. nc), or anti-CD3/CD28-activated nc (anti-CD3/anti-CD28 nc). Supernatants from the cocultures or from anti-CD3/anti-CD28 ac or anti-CD3/anti-CD28 nc alone were collected after 4, 8, and 24 h and analyzed for their cytokine content by Luminex. For IL-6, TNF-{alpha}, and MIP-1beta supernatants, n ≥6 except for DC/PHA o.n., where n = 4 and anti-CD3/anti-CD28 nc alone, where n = 3. For statistical comparison of samples, the Mann-Whitney test was used and significant differences are indicated as *, p ≤ 0.05; **, p ≤ 0.01; or ***, p ≤ 0.001.

 
In most donors analyzed, anti-CD3/CD28-activated apoptotic cells induced the highest levels of IL-6, TNF-{alpha}, and MIP-1beta and also the most rapid release from DCs. Supernatants from DCs cocultured with nonactivated apoptotic cells or nonactivated necrotic cells did not contain significant amounts of cytokines compared with the medium control. IL-2, IL-8, and IFN-{gamma} were detected in supernatants both from cocultures containing activated apoptotic cells and from the actively apoptotic cells per se. Therefore, it was not possible to attribute the production of these cytokines to the DCs (data not shown). IL-10 and IL-12 p70 release was detected from DCs activated with LPS but not in any of the DC/apoptotic cell cocultures examined (data not shown). The results show that DCs produce proinflammatory cytokines IL-6 and TNF-{alpha} and the chemokine MIP-1beta after interaction with activated apoptotic PBMCs and to some extent with activated necrotic PBMCs. However, these DCs failed to produce IL-10 and IL-12p70.

DCs stimulate proliferation and IFN-{gamma} production in autologous T cells after uptake of allogeneic, activated apoptotic PBMCs

We next asked whether DCs matured by coculture with activated apoptotic PBMCs were able to induce proliferation and activation of naive autologous T cells using alloantigen as a model Ag (Fig. 8). Autologous T cells were added to DCs that had ingested either nonactivated or activated, allogeneic apoptotic PBMCs. Hence, the DCs and the T cells were autologous whereas the apoptotic cells used to feed the DCs were of allogeneic origin. The DC/apoptotic cell cocultures were incubated for 48 h before addition of autologous CFSE-labeled T cells. CFSE-labeled T cells alone or T cells added to activated apoptotic cells were used as negative controls for induction of T cell proliferation. As a positive control, the superantigen SEB was added to DCs along with autologous T cells. Cocultures were incubated for 3, 4, 5, or 6 days to determine the peak of T cell proliferation. At these time points, cells were collected and stained for CD1a and CD3 as well as intracellular IFN-{gamma} production. In the SEB-stimulated cultures T cell proliferation (CD1a–CD3+) peaked at day 4, which coincided with the highest frequency of IFN-{gamma}-positive T cells. DCs cocultured with activated, apoptotic allogeneic cells were capable of inducing both proliferation and IFN-{gamma} production in autologous T cells. As in the SEB control, both proliferation and IFN-{gamma} production peaked at day 4 (Fig. 8, b, f, and g). In the cultures containing only DCs and autologous T cells, T cells only, or in samples where DCs were fed resting apoptotic allogeneic cells, neither T cell proliferation nor IFN-{gamma} production was detected at any of the time points analyzed (Fig. 8, a, c, e, and g). In samples containing T cells and activated, allogeneic apoptotic cells but no DCs, there was in some donors a small frequency of proliferating T cells producing IFN-{gamma} (Fig. 8g). This fraction was however not significantly different compared with background in samples containing DCs and resting apoptotic allogeneic cells. DCs cocultured with activated, allogeneic necrotic cells induced both proliferation and IFN-{gamma} production in T cells, whereas DCs fed with nonactivated allogeneic necrotic cells were unable to induce these responses (Fig. 8g). We also wanted to control for possible FcR-mediated effects on DC maturation and induction of efficient Ag-presenting capacity. It has earlier been shown that opsonization of apoptotic cells with autoantibodies leads to enhanced Fc{gamma}R- mediated uptake and Ag-specific T cell proliferation (44, 45). We therefore incubated PBMCs with an anti-CD2 mAb before exposure to gamma irradiation and addition to DCs. Anti-CD2 did not activate the allogeneic T cells as measured by CD25 and CD69 expression and did not induce DC activation and subsequent autologous T cell proliferation using monocyte-derived DCs (Fig. 8g). Therefore, we conclude that FcR-mediated signaling events alone may not be sufficient for induction of DC maturation and subsequent presentation of alloantigens to specific T cells. These results show that activated, but not resting, allogeneic apoptotic and necrotic PBMCs are able to induce DC maturation that leads to efficient presentation of alloantigens to autologous T cells.


Figure 8
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  		FIGURE 8. Alloantigenpresentation and T cell activation by DCs after uptake of activated, apoptotic PBMCs. Immature DCs were cocultured with nonactivated or activated allogenic ac. In control wells, only medium (a) or activated ac without DCs (d) were added. After 48 h, CFSE-labeled autologous T cells were added to all wells in a ratio of 5:1 (T cells:DCs). SEB was added as a positive control (b). Three, 4, 5, or 6 days after addition of T cells, the cultures were stained for cell surface markers and intracellular IFN-{gamma} and were analyzed by flow cytometry. Gates were set on CD3+CD1a– T cells. In medium- and T cell-only controls (a and c) and in samples where DCs were fed resting ac (e), no T cell proliferation or IFN-{gamma} production was detected at any of the time points analyzed. Low levels of proliferation and IFN-{gamma} production compared with medium control was detected for some donors in autologous T cells that encountered activated ac only (d). T cell division and IFN-{gamma} production was detected at day 3 and peaked at day 4 in positive controls (b) and in samples where DCs were cocultured with activated ac (f). a–f, Data from one representative donor of six at day 4 of the experiment are shown and numbers indicate percentages of IFN-{gamma}+ T cells. g, The percentage of IFN-{gamma}+ T cells collected from six different donors at day 4 of the experiment is shown. In control samples where DCs were fed anti-CD2-treated ac, data originate from three different donors. Similar to DCs cocultured with anti-CD3/CD28-activated ac cells (APC: DC, ag: act ac), DCs cocultured with anti-CD3/CD28-activated nc (APC: DC, ag: act nc) induced autologous T cell proliferation and IFN-{gamma} production. DCs cocultured with anti-CD2-opsonized ac (APC: DC, ag: anti-CD2 ac), with resting nc (APC: DC, ag: nonact nc) or activated nc alone (APC: –, ag: act nc) were unable to induce this response in T cells. Significant up-regulation of IFN-{gamma} production in proliferating T cells compared with DC medium control is indicated as **, p ≤ 0.01 or ***, p ≤ 0.001.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
In this study, we demonstrated that activated, but not resting, apoptotic PBMCs are able to induce activation of DCs in terms of up-regulation of costimulatory molecules, induction of proinflammatory cytokine release, and presentation of alloantigens that leads to autologous T cell proliferation and IFN-{gamma} production. Necrotic PBMCs also induced DC maturation as previously shown (11, 20). This occurred if the necrotic PBMCs were initially activated, but failed to do so in the absence of preceding activation. We observed that PBMCs activated with PHA for 4 days were less efficient in maturing DCs. A possible explanation could be that these cells have entered a later stage of activation where the factor or factors inducing DC maturation are not as frequently exposed as in newly activated PBMCs. These observations suggest that one factor influencing the immunogenicity of a dying cell is determined by its activation state before death. The frequency of DCs that up-regulated CD86 did not correspond to the frequency of DCs phagocytosing activated apoptotic cells. Most likely this is due to a bystander effect where surrounding DCs are affected by cytokines released by the phagocytosing DCs.

The present report supports earlier studies where apoptotic cells were found to mature DCs and lead to induction of T cell activation in vitro (29, 30, 32, 46, 47, 48). In addition, our data support previous findings that certain apoptotic cells can be better than necrotic cells to induce immune responses (49, 50). The apoptotic cells inducing DC activation in the former studies all contained different forms of tumor or viral Ags. The danger signaling features of the apoptotic cells in the previous studies are still not fully characterized, but we speculate that the capacity of apoptotic cells to mediate DC maturation could at least in part be associated with a "nonresting" state. It should be noted that no exogenous TLR ligand, tumor, or viral source of Ag was present in the experiments described in the present study. The PBMCs used in the present study were obtained from healthy blood donors and the experiments were always set up using activated and nonactivated PBMCs from the same donors in parallel. We suggest that the state of activation, before entering apoptosis or necrosis, determined whether the cells were able to activate DCs. Use of resting apoptotic cells in the experimental setup could partly explain why some studies have found apoptotic cells to be unable to mature DCs in vitro (11, 20), to possess anti-inflammatory properties (23, 24, 25, 51), or to induce tolerance instead of immune activation (26, 27, 52). Recent data suggest that different apoptotic pathways may also determine the relative immunogenicity of dying cells (34). Opsonization of apoptotic cells has been shown to trigger FcR signaling in DCs, leading to DC maturation/activation (53, 54, 55, 56). The opsonization of primary, nonactivated, apoptotic, PBMCs using anti-CD2 Ab in the present report did not lead to human DC maturation or increased Ag presentation. These data do not exclude that FcR interactions can contribute to signaling events leading to DC maturation in our experiments when using the anti-CD3/anti-CD28 mAbs, but it shows that these FcR interactions alone are not sufficient to trigger DC maturation. We suggest that primary dying cells, subjected to activation before death, express molecules that are lacking in primary resting dying cells and that these molecules trigger the maturation process in DCs, leading to efficient Ag presentation. Other cells, such as apoptotic tumor cells, may share these immunogenic features to different extents. Opsonization of these cells may enhance their ability to mature DCs by providing additional FcR- mediated signals. Characterization of the molecules expressed by dying cells that determines their immunogenic potential would facilitate the development of treatment strategies using dead cells as an Ag delivery system for induction of immune responses.

Several endogenous factors originating from dying cells have been identified as possible mediators of endogenous adjuvant effects for induction of DC activation. Heat shock proteins and uric acid have previously been shown to induce DC maturation (18, 57, 58, 59, 60, 61, 62, 63) and exert adjuvant activity (64, 65, 66, 67, 68). These molecules are normally intracellular and released upon loss of membrane integrity. Heat shock proteins, uric acid, or other endogenous adjuvant molecules released upon loss of membrane integrity could possibly have some effect in the in vitro system used here, where some of the irradiated PBMCs enter secondary necrosis before uptake of DCs. Yet this is not a fully satisfying explanation of our results for two reasons. First, supernatants collected from apoptotic cells contained factors released from cells in secondary necrosis. These supernatants lacked the ability to induce DC maturation. Second, comparing apoptotic and necrotic cells, the latter were less efficient in up-regulating costimulatory molecules on DCs.

The finding that DCs cocultured with activated PBMCs up-regulated CD86 expression, while supernatants from the same apoptotic cell preparations were not effective in inducing CD86 expression indicates that interaction between DCs and activated apoptotic cells was required for induction of DC maturation. However, it does not exclude the possibility that extracellular factors released from apoptotic cells in a close proximity to or within the DC can mediate up-regulation of costimulatory molecules.

The present report shows that exposure to activated apoptotic cells induces production of the proinflammatory cytokines IL-6 and TNF-{alpha} as well as the chemokine MIP-1beta in DCs. In an inflammatory event, caused by pathogens or injury of host cells, immune cells are recruited to the site of inflammation. Under circumstances where T cells are recruited and activated upon Ag recognition, they will eventually die by apoptosis. We speculate that apoptotic cell death of activated T cells could function as a positive feedback mechanism for both the innate and adaptive responses in an inflammatory event. We detected similar induction of DC maturation in the present report irrespective of whether the activated apoptotic cells were autologous or allogeneic (data not shown). This could implicate that uptake of activated apoptotic lymphocytes by immature DCs, residing at the site of infection or injury, leads to proinflammatory cytokine and chemokine release from the DCs. The release of proinflammatory cytokines and chemokines would increase the recruitment of immune cells to the site of danger.

The endogenous adjuvant effect attributed to activated apoptotic cells reported here could also be of relevance for design of vaccines. Certain vectors currently under development, such as canarypox or Semliki forest viruses, induce apoptosis in their target cell, which may subsequently lead to cross-presentation of Ags (32, 69). In addition, apoptotic cells may be used to load DCs in vitro with Ag or could be used directly for immunization against, for example, cancer or infectious agents (70). The present report suggests that the choice of apoptotic cells to be used for vaccination can per se be of relevance to obtain induction of an effective immune response.

Taken together, the findings of this study show that dying cells have different abilities to elicit immune responses depending on their state of activation. An increased understanding of how to potentially benefit from apoptosis of host cells, DC maturation, and subsequent induction of immune responses could be used for development of vaccines that use cross-presentation of apoptotic cells.


   Acknowledgments
 
We thank Anette Hofmann for confocal microscopy and Steven Applequist for critically reading this manuscript.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
Lilian Walther-Jallow is a full-time employee of Avaris, Anna-Lena Spetz is a part-time employee of Avaris. EU support to Avaris AB is patent pending. Anna-Lena Spetz is owner of stock in Avaris.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This research was supported by the Swedish Research Council, Swedish Cancer Society, Swedish International Development Cooperation Agency/Department for Research Cooperation, Swedish Physicians Against AIDS Research Foundation, European Commission STREP-018953, and Swedish Foundation for Strategic Research. Back

2 Address correspondence and reprint requests to Dr. Ulrika Johansson, Department of Medicine, Center for Infectious Medicine, F59, Karolinska Institutet, Karolinska University Hospital Huddinge, Stockholm, Sweden. E-mail address: ulrika.johansson{at}ki.se Back

3 Abbreviations used in this paper: DC, dendritic cell; PI, propidium iodide; SEB, staphylococcal enterotoxin B; CD40L, CD40 ligand; ac, apoptotic cell; nc, necrotic cell; o.n., overnight. Back

Received for publication August 16, 2006. Accepted for publication May 29, 2007.


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 Introduction
 Materials and Methods
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