The fate of dendritic cells (DCs) after Ag presentation may be DC subset-specific and controlled by many factors. The role of activation-induced apoptosis in regulating DC function is not clear. We investigated the fate of cutaneous DCs (cDCs), specifically Langerhans cells (LCs), and observed that they undergo apoptosis after successful Ag presentation to CD4 T cells. Caspase-specific inhibitors revealed that LC lines use a type II apoptosis pathway in response to CD4 T cells. In support of this, BH3-interacting domain (Bid) protein was present at high levels and specifically cleaved in the presence of Ag-specific T cells. Significant resistance to apoptosis by OT-2 CD4 cells was also observed for Bid knockout (KO) LCs in vitro. To test whether Bid was required to regulate LC function in vivo, we measured contact sensitization and topical immunization responses in Bid KO mice and observed markedly enhanced ear swelling and proliferation responses compared with wild-type mice. Furthermore, when Ag-pulsed Bid KO migratory cDCs were inoculated into wild-type recipients, an increase in both the rate and percentage of expanded OT-2 T cells expressing IFN-γ was observed. Thus, enhanced Ag presentation function was intrinsic to Bid KO cDCs. Therefore, Bid is an important regulator of LC viability and Ag presentation function.

Langerhans cells (LCs)3 are a subset of dendritic cells (DCs), which reside in the epidermal layer of skin. They are potent APCs that serve as activators of skin-specific immune responses to contact sensitizers, viruses, and tumors. More recently they have been implicated in playing a major role for maintaining tolerance to damaged skin proteins (1, 2, 3, 4). As immature DCs, they can capture and process Ag at picomolar concentrations (5) but are weak stimulators of naive T cell responses. Once a breach in the skin integument is sensed, LCs migrate out of the epidermis to dermal lymphatics that lead to draining lymph nodes. As they migrate, LCs lose Ag capture function and gain “mature” DC function, which is completed by their interaction with T cells in the lymph node (6). Such activated mature DCs have two functions: 1) to activate naive T cells and 2) to direct T cell differentiation into Ag-specific effector and regulatory cells. Thus, LCs determine the magnitude and type of immunologic response made.

Tissue-resident LCs have been shown to possess self-renewing potential, remaining in the epidermis for >18 mo (7). However, the life span of LCs after they have been activated to migrate to lymph nodes for Ag presentation is not known. Dermal DCs (dDCs) are phenotypically similar to conventional DCs. In the case of conventional DCs, short half-lives have been reported. In spleen it is 1–3 days (8, 9), and in lymph node, Ag pulsed-DC survive for only 24 h in the presence of Ag-specific T cells (10). Multiple mechanisms may contribute to the brevity of DC life spans. First, pathogen stimuli sensed through Toll receptors on DCs have been shown to activate an internal “clock” that limits their remaining life span (11). Such stimuli induces high levels of the proapoptotic “Bcl-2 homology domain 3 (BH3)-only domain” protein Bim. Second, once in lymphoid tissue and after Ag presentation, depletion of DC cytokine secretion and/or costimulatory capacity may lead to apoptosis by exhaustion (12). Both of these rely on the intrinsic passive cell death pathway. A third possibility is that an active process of extrinsic death signals derived from successfully activated T cells hastens the demise of DCs (13). However, in vitro studies indicate that while immature DCs are sensitive to NK, T cell and Fas-mediated apoptosis, mature DCs are resistant (14). Therefore, it is not clear if, or at what stage, DCs undergo T cell mediated apoptosis in vivo, and it is likely that the mechanisms that regulate cell death may be unique for different DC subsets.

Extrinsic apoptotic signals through TNF family member death receptors use two types of caspase signaling cascades (15, 16, 17). Type I cells, such as T cells, enable “initiator” caspase-8 to directly activate “executioner” caspase-3. In type II cells, a specific member of the BH3-only proteins, called BH3-interacting domain (Bid) death agonist, has the unique task of linking the extrinsic and intrinsic pathways. Extrinsic signals in type II cells use Bid as the target substrate of caspase-8, generating a truncated active form of Bid that translocates to the mitochondrial membrane and directly activates proapoptotic Bcl-2 family members Bak and Bax. Thus, Bid activates the mitochondrial “amplification loop” leading to apoptosome assembly and caspase-9 activation, caspase-3 activation, and irreversible cell death (18). Although there is much evidence that mitochondrial pro- and antiapoptotic Bcl-2 family proteins have a profound effect on DC longevity, either in setting the intrinsic clock (11), in modulating Fas-mediated death in vitro (19), or when introduced epigenetically as transgenes (20, 21, 22), it is not clear what type of apoptosis pathway is normally used in response to extrinsic signals under physiologic conditions.

In this study, we followed the fate of LCs after Ag presentation to CD4 T cells and found that naive CD4 T cells were able to induce LC apoptosis in an Ag-specific manner. We then examined what type of apoptotic pathway is used in response to extrinsic signals and showed that LC lines are type II cells, expressing high levels of Bid protein that is activated upon Ag presentation to CD4 T cells. To prove that Bid plays a critical role in regulating LC Ag presentation, we then examined immune responses in topically immunized Bid knockout (KO) mice (23). We observed a marked increase in contact sensitization and topical immunization responses by T cells. Moreover, cutaneous DCs (cDCs) derived from Bid KO mice could, themselves, impart enhanced T cell responses in vivo and demonstrated resistance to CD4 T cell-mediated apoptosis in vitro. Thus, our studies indicate that Bid is an important regulator of LC viability and function.

Specific pathogen-free A/J mice, C57BL/6 mice, and TCR transgenic 3A9 (specific for hen egg lysozyme (HEL) in the context of I-Ak) mice were obtained from The Jackson Laboratory (stock no. 002597) (24). TCR transgenic CD45.1 OT-2 mice were provided by Dr. R. Pat Bucy (University of Alabama at Birmingham, Birmingham, AL). Bid-deficient (referred to here as Bid KO) mice on a C57BL/6 background were provided by Dr. S. L. Korsmeyer (Harvard Medical Center, Boston, MA) (23). In adoptive transfer experiments where both OT-2 and cDCs were injected, we used albino C57BL/6J-Tyrc-2J/J mice from The Jackson Laboratory to serve as wild-type (WT) recipients. Mice were bred and maintained under specific pathogen-free conditions at the University of Alabama at Birmingham and experiments performed with Institutional Animal Care and Use Committee-approved protocols.

The XS106 LC line was established from the epidermis of newborn A/J mice and maintained in vitro, as described previously (25) (obtained as a gift from Dr. A. Takashima, University of Toledo College of Medicine, Toledo, OH) and demonstrates potent LC function in vitro and in vivo (26, 27). XS106-GFP B1.1 clone was generated by limiting dilution of cells infected with reporter GFP encoding lentiviral vector obtained as a gift from Dr. X. Wu (University of Alabama at Birmingham) (28, 29). No functional difference was observed for this cell line when compared with parental XS106 cells. The 3A9 T cell hybridoma was obtained as a gift from Dr. P. M. Allen (Washington University, School of Medicine, St. Louis, MO) (30).

For all cell culture and assays, unless noted, we used RPMI 1640 supplemented with heat-inactivated FBS (10%), l-glutamine (200 mM), sodium pyruvate (100 mM), HEPES buffer (1 M), minimum essential amino acids (100 mM), and penicillin/streptomycin (10,000 IU/ml) all from Mediatech. For XS106 (LC) cell line cultivation, we supplemented further with 2-ME (5 mM) (Sigma-Aldrich), GM-CSF (0.5 ng/ml) (Sigma-Aldrich), and NS47 conditioned supernatant (5%), as described previously (25).

Mice were anesthetized then Ag applied to tape-stripped ears (10 times) by painting with 25 μg of OVA or HEL in 10 μl of PBS with or without inclusion of 10 ng/ml LPS per side or with PBS ± LPS alone, as indicated. After 30 min, mice were sacrificed, and ear tissue was harvested. Ears specimens were split into dorsal and ventral halves, floated dermal side down, and cultured for 2 days in 24-well plates (31). In some experiments, culture medium additionally contained 100 μg/ml of relevant or irrelevant Ag, as indicated. The cells that migrated from the skin specimens into the culture medium were harvested, passed through a screen to remove large skin debris, and examined for cell counts, viability by trypan blue exclusion, and phenotype. Migratory cells routinely contained >50% I-A and CD11c double-positive cells, as determined by flow cytometry (32). Additionally, the I-A-positive fraction was 70–90% double positive for the LC markers, CD205 (DEC-205 clone NLDC145 from Cedarlane Laboratories) and Langerin/CD207 (clones 205C1 and 929F3) (AbCys) (data not shown). Isotype controls were obtained from BD Pharmigen.

Naive splenic CD4 T cells were purified from either 3A9 or OT-2 transgenic mice using CD4-conjugated Dynabeads in conjunction with the Detach-a-bead kit (Dynal Biotech) or a MACS purification kit (Miltenyl Biotec). The purity of CD4 cells was confirmed by double staining for CD4 and TCR-specific Abs to the OT-2 TCR expressing Va5.1 (BD Biosciences Pharmingen) or the 3A9 TCR Vb8.2 (clone F23.2; provided by Dr. P. Marrack, National Jewish Medical and Research Center, Denver, CO; Ref. 33). Purified T cells were routinely >95% CD4 and TCR positive.

Pan caspase inhibitor Z-VAD-FMK, caspase-8 inhibitor Z-IETD-FMK, caspase-9 inhibitor Z-LEHD-FMK, caspase inhibitor control Z-FA-FMK (all from R&D Systems), and caspase-3 inhibitor Z-DEVD-FMK (Kamiya Biomedical) were used. The following mAb were used: mouse anti-Bid clone 40 (BD Transduction Laboratories), monoclonal anti-β-actin clone AC-15 (Sigma-Aldrich), polyclonal rabbit caspase-3 Ab (Cell Signaling Technology), polyclonal caspase-9 mouse specific (Cell Signaling Technology), polyclonal rabbit anti-caspase-8 (BD Pharmingen), anti-rabbit Ig HRP linked F(ab′)2 (Amersham Biosciences), anti-mouse Ig HRP linked whole Ab (Amersham Biosciences), Annexin VPE (BD Pharmingen), and FITC anti-mouse I-A/I-E (2G9) (BD Pharmingen). Annexin V binding buffer, 10× (BD Pharmingen), staurosporine (Sigma-Aldrich), HEL (Sigma-Aldrich), 7-aminoactynomicin D (7-AAD) (Sigma-Aldrich), and Pierce BCA Protein Assay (Pierce) were used.

Cell protein lysates were obtained from pellets of 107 cells, washed twice with PBS (Mediatech), and dissolved in 100 μl of lysis buffer containing 50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 10% Nonidet P-40, 1 mM EDTA, 0.5% PMSF (1 mM), 0.5% Na3VO4 (1 mM), 0.5% sodium fluoride, 0.1% leupeptin, and 0.1% apoprotinin. Protein concentrations were determined using the Pierce BCA Protein Assay (Pierce) and the Bio-Rad Model 550 microplate reader and the Microplate manager 4.01 software with an absorbance of 565 nm.

Proteins were subject to electrophoreses in Bio-Rad ready gel 12% Tris-HCl (Bio-Rad) and transferred to Bio-Rad nitrocellulose membranes (Bio-Rad). RNP800 m.w. marker was used (Amersham Biosciences) to determine protein sizes.

Membranes were blocked with 2% ECL Advance blocking agent (Amersham Biosciences), and hybridization was performed with the indicated primary Abs. Murine mAb to Bid was used at 1/1000 (clone 40; BD Transduction Laboratories) and murine mAb to β-actin (clone AC-15; Sigma-Aldrich) was used at 1:2 × 106. ECL Advance Western blotting detection kit from Amersham Biosciences was used. After incubation with appropriate secondary Abs, bands were visualized using the ECL advance Western blotting detection kit (Amersham Biosciences). Densitometry analyses were performed on a Macintosh computer using the public domain NIH Image program (developed at the U.S. National Institutes of Health and available on the Internet at 〈http://rsb.info.nih.gov/nih-image/〉).

cDCs were isolated from the supernatant of 2-day mouse ear skin organ cultures. DCs were further cultured overnight in 100 μg/ml Ag (HEL) or chicken OVA (Sigma-Aldrich) protein and/or Ag-specific T cells (3A9 T hybridoma or purified CD4 transgenic cells (HEL-I-Ak specific) or OT-2 CD4 T cells (OVA-I-Ab specific) at a ratio of 1:4 LC:TC. After overnight culture (18–24 h), cells were spun down at 300 × g for 10 min and incubated 30 min on ice with 100 μl of annexin binding buffer (1×) (BD Pharmingen) with 3 μl of Annexin VPE (BD Pharmingen) and FITC-anti-I-Ak (2G9) (BD Pharmingen) and/or allophycocyanin-CD11c (HL3) (BD Pharmingen) per sample. In some experiments, biotinylated-Langerin/CD207 (205C1) (Abcys Biologie) and PE-CD205/DEC205 (NLDC-145) (Cedarlane Laboratories) were used. All isotype controls were purchased from BD Pharmingen. After washing the samples with annexin binding buffer, pelleted cells were resuspended in 400 μl of annexin binding buffer containing 2.5 μg/ml 7-AAD (Sigma-Aldrich). Multiparameter flow cytometric analysis of Annexin VPE and 7-AAD staining was performed gated cells as indicated.

XS-GFP (clone B1.1) cells were pulsed with 100 μg/ml HEL (Sigma-Aldrich) overnight then mixed with either 3A9 T cells or unlabeled XS106 as a negative control. One hour before mixing cells, they were preincubated with the following caspase inhibitors: 100 μM Z-VAD-FMK, 200 μM Z-DEVD-FMK, 150 μM Z-LEHD-FMK, and 200 μM Ac-IETD-CHO and then continuously cultured in the presence of these inhibitors. In some experiments, the positive control for apoptosis were cells treated with staurosporine (500 nM) (Sigma-Aldrich) overnight. Apoptosis levels were assessed after 16–20 h, and XS-GFP were gated by green fluorescence and analyzed for apoptosis by Annexin VPE and 7-AAD staining (as described previously). Bar graphs of percent apoptosis values include the loss of GFP-positive cells that occurs in late stages of apoptosis and therefore unavailable for cytometric analysis and adjusted for nonspecific toxicity of peptides, as determined in control cultures and is calculated as follows (in this example, LCs are GFP positive, but in other experiments, LCs may be stained with other markers, such as Langerin instead) percentage of specific cell death = percentage of specific LC loss + specific apoptotic fraction of remaining percentage of LC, where percentage of specific loss = (control percentage of GFP cells − experimental percentage of GFP cells) and specific apoptotic fraction of remaining percentage of LC = ((experimental fraction of apoptotic cells (percentage of annexin V and/or 7-AAD positive/100) × percentage of experimental GFP cells) − (control fraction of apoptotic cells × percentage of control GFP cells)).

The density of epidermal LCs was ascertained from whole mount epidermal sheets stained for MHC class II molecules. Ear specimens were split and cartilage removed from the dermal side of the dorsal half. Ear epidermis was lifted from the dermis after floating the specimens’ dermal side down in 0.5 M ammonium thiocyanate (Sigma-Aldrich) in PBS for 20 min. Epidermal sheets were fixed with cold acetone and washed in PBS. Blocking Ab solution (clone 2.4G2 hybridoma (10 μg/sample) from American Type Culture Collection) was added and incubated 1 h at 4°C. LCs were labeled with FITC-anti-Ia Ab (clone 2G9 (1 μg/sample); BD Pharmingen) and incubated at 4°C overnight. Excess Ab was washed from sheets by three 30-min incubations with cold PBS then mounted in 0.2% n-propyl gallate-glycerol. Images at ×20 magnification were captured (MetaMorph Imaging System; Universal Imaging) and the number of positive cells counted by a third experimenter blinded to sample identity.

Mice were sensitized then challenged with the hapten 2–4-dinitrofluorobenzene (DNFB) to elicit a contact hypersensitivity response as per Xu et al. (34). For the induction phase, hapten painting treatment occurred on day 0 and day 1 as follows: five age-matched mice per group of WT or Bid KO were sensitized with either vehicle only (4:1 acetone and olive oil) as a negative control or 0.5% DNFB (Sigma-Aldrich) by application of 25 μl on the shaved abdomen and 5 μl on each footpad per mouse. On day 5 postsensitization, a baseline measurement of ear thickness was taken using an engineer’s micrometer (Mitutoyo Precision). Elicitation was induced by painting the ears of both vehicle control and sensitized mice with 10 μl of 0.2% DNFB per dorsal and ventral sides of each ear (referred to as day 0 with respect to day of challenge). Ear thickness measurements of each mouse was taken each day thereafter, as indicated. The averages of ear thickness over baseline measurements for each group (n = 5) are shown and expressed in micrometers. The magnitude of the swelling response is given as the mean and SEM of each group.

Purified OT-2 T cells (∼5 × 106/ml) were labeled with 1 μM CellTracker Green CMFDA (synonymous with CFSE) (Molecular Probes) for 8 min at room temperature, then neutralized with 20% FBS-PBS and washed three times. Cells were adjusted in PBS to deliver 1–2 × 106 cells by injecting 100–200 μl i.v. into the tail vein of WT or Bid KO mice. In some experiments, CFSE-OT-2 T cell injected WT mice also received 5 × 104 cutaneous migratory DC from WT or Bid KO skin cultures (as described above) in 20 μl of PBS into the right footpad. After 3 days, mice were sacrificed for harvesting spleen and draining popliteal lymph nodes, which were weighed and prepared for single cell suspension. Cell suspensions were stained with transgenic TCR-specific Abs and in some cases with the activation marker CD69 (as indicated), and the fluorescence intensity of CFSE was detected by flow cytometric analysis using a BD FACSCalibur system (BD Biosciences).

WT mice were injected with 2 × 106 purified allotypically marked CD45.1-positive OT-2 T cells. Groups of mice (n = 3) were immunized either topically with OVA (as described above) as a positive control or cDC-immunized by injection of 2 × 105 migratory DC, from OVA painted WT or Bid KO ear skin cultures, into the right footpad. Mice were sacrificed after 4 days and single-cell suspensions from draining lymph nodes and spleen from each of the mice were prepared individually by collagenase D digestion (2 mg/ml; Roche). Lymph node cells were prepared immediately for cytokine secretion assay. Spleen cells were subjected to a restimulation culture period of 3 days in the presence or absence of 100 μg/ml OVA before intracellular cytokine assay. To activate cytokine synthesis, lymph node and spleen cells were cultured for 6 h with Golgi-Stop (BD Biosciences), 50 ng/ml PMA (Sigma-Aldrich), and 500 ng/ml calcium ionophore A23187 (Sigma-Aldrich) following BD Cytofix/Cytoperm kit instructions. Following treatment, cells were treated with FcR block and stained with FITC-anti-CD3, biotinylated anti-CD45.1, and PerCP-conjugated streptavidin. After fixation/permeabilization, intracellular cytokines were stained with PE-anti-IL-4 and Alexa 647-anti-IFN-γ Abs (all from BD Pharmingen). Flow cytometric analysis was performed on a FACSCalibur using CellQuest Pro software.

One-tailed Student’s t test was applied, and the p values are indicated in the text and figure legends.

To test whether LCs are down-regulated by apoptosis in a T cell-specific manner, we examined the susceptibility to apoptosis of both migratory murine cDCs and the epidermal-derived LC line, XS106, upon Ag presentation to specific 3A9 T cell hybridoma in 18–24 h cultures (Fig. 1, A and B). To more easily track XS106 cells in mixed cell assays, we developed a series of stable GFP transfectants of XS106. Using a XS-GFP clone, we tested the role of specific or nonspecific T cells in mediating apoptosis. Importantly, we extended these studies to test if normal, naive splenic T cells also induced apoptosis in LC lines (Fig. 1 C).

FIGURE 1.

T cell-mediated apoptosis of LCs or XS106 LC lines by Ag-specific T cells. A, LCs are susceptible to CD4 T cell-mediated apoptosis. LCs were isolated from the supernatant of 2-day mouse ear skin organ cultures. LCs were further cultured overnight in 100 ng/ml HEL protein and/or 3A9 T hybridoma at a ratio of 1:4 LC:TC. Cultured samples were stained with FITC-anti-I-Ak (2G9), Annexin VPE, and 7-AAD (5 μg/ml). The uptake of vitally excluded dye 7-AAD was measured by flow cytometry as an indicator of later stages of death and annexin V staining as an early indicator of apoptosis. Three-color flow cytometric analysis of Annexin VPE and 7-AAD staining was performed on FITC-I-Ak-positive gated cells. B, Ag-specific T cell-induced apoptosis in XS106 cells. GFP-positive clone XS106 B1.1 cells were incubated with or without HEL protein (1 mg/ml) overnight, then mixed with T hybridoma cells specific for HEL (3A9) or irrelevant OVA (DO-BW) Ags at a ratio of 1:4 LC:TC and cultured overnight. GFP-positive XS106 were gated and analyzed for Annexin VPE and 7-AAD staining by flow cytometry, and the results were displayed in a contour map. C, Naive CD4 splenocytes from 3A9 transgenic mice induce significant apoptosis in XS106 cells. XS106 cells were incubated with HEL protein (1 mg/ml) overnight, then mixed with purified CD4 3A9 splenocytes specific for HEL (3A9) at a ratio of 1:5 LC:TC and cultured overnight. CD4 T cell and LC populations were gated by large light scatter properties. CD4 cells alone, HEL-pulsed XS106 alone, or the mix of CD4 + HEL-pulsed XS106 were analyzed for Annexin VPE and 7-AAD staining by flow cytometry. Numbers indicate the percentage of gated cells within that quadrant. Lower left quadrant contains viable cells. D, Naive T cells respond to XS106 Ag presentation. Purified naive splenic CD4 T cells from 3A9 transgenic mice were mixed with LCs (XS106 cells) and cocultured for 20 h in the presence or absence of HEL (100 μg/ml). Small and blast T cells were easily distinguished from large XS106 and gated by light scatter properties for analysis of CD69 and TCR expression. The percentage of T cells in each quadrant is shown. E, IL-2 production from Ag-specific T cell hybridoma, 3A9. XS-GFP cells were prepulsed with or without HEL protein (1 mg/ml) overnight, then mixed with HEL-3A9-specific or irrelevant OVA-specific DO-BW T hybridoma cells at a ratio of 1:4 LC:TC and cultured for another 20 h. T hybridoma activation was tested by detecting IL-2 in the culture supernatants by ELISA.

FIGURE 1.

T cell-mediated apoptosis of LCs or XS106 LC lines by Ag-specific T cells. A, LCs are susceptible to CD4 T cell-mediated apoptosis. LCs were isolated from the supernatant of 2-day mouse ear skin organ cultures. LCs were further cultured overnight in 100 ng/ml HEL protein and/or 3A9 T hybridoma at a ratio of 1:4 LC:TC. Cultured samples were stained with FITC-anti-I-Ak (2G9), Annexin VPE, and 7-AAD (5 μg/ml). The uptake of vitally excluded dye 7-AAD was measured by flow cytometry as an indicator of later stages of death and annexin V staining as an early indicator of apoptosis. Three-color flow cytometric analysis of Annexin VPE and 7-AAD staining was performed on FITC-I-Ak-positive gated cells. B, Ag-specific T cell-induced apoptosis in XS106 cells. GFP-positive clone XS106 B1.1 cells were incubated with or without HEL protein (1 mg/ml) overnight, then mixed with T hybridoma cells specific for HEL (3A9) or irrelevant OVA (DO-BW) Ags at a ratio of 1:4 LC:TC and cultured overnight. GFP-positive XS106 were gated and analyzed for Annexin VPE and 7-AAD staining by flow cytometry, and the results were displayed in a contour map. C, Naive CD4 splenocytes from 3A9 transgenic mice induce significant apoptosis in XS106 cells. XS106 cells were incubated with HEL protein (1 mg/ml) overnight, then mixed with purified CD4 3A9 splenocytes specific for HEL (3A9) at a ratio of 1:5 LC:TC and cultured overnight. CD4 T cell and LC populations were gated by large light scatter properties. CD4 cells alone, HEL-pulsed XS106 alone, or the mix of CD4 + HEL-pulsed XS106 were analyzed for Annexin VPE and 7-AAD staining by flow cytometry. Numbers indicate the percentage of gated cells within that quadrant. Lower left quadrant contains viable cells. D, Naive T cells respond to XS106 Ag presentation. Purified naive splenic CD4 T cells from 3A9 transgenic mice were mixed with LCs (XS106 cells) and cocultured for 20 h in the presence or absence of HEL (100 μg/ml). Small and blast T cells were easily distinguished from large XS106 and gated by light scatter properties for analysis of CD69 and TCR expression. The percentage of T cells in each quadrant is shown. E, IL-2 production from Ag-specific T cell hybridoma, 3A9. XS-GFP cells were prepulsed with or without HEL protein (1 mg/ml) overnight, then mixed with HEL-3A9-specific or irrelevant OVA-specific DO-BW T hybridoma cells at a ratio of 1:4 LC:TC and cultured for another 20 h. T hybridoma activation was tested by detecting IL-2 in the culture supernatants by ELISA.

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The results show that both LCs (analysis gated on I-Ahigh-expressing cells representing LCs derived from skin explant cultures) and the XS-GFP line are highly susceptible to apoptosis mediated by Ag-specific T cell hybridomas in a T cell-specific and Ag-specific manner (Fig. 1, A and B). Moreover, naive CD4 transgenic T cells are also potent inducers of XS106 cell apoptosis (Fig. 1,C). The XS-GFP line has potent Ag-presenting function because successful T cell activation occurs in these cultures, as shown by CD69 expression and TCR down-regulation (Fig. 1,D), as well as Ag-specific IL-2 secretion (Fig. 1 E). In time course studies to determine the kinetics of LC death, we have observed little apoptosis within 8 h (data not shown). This late timing ensures that LC activation, effective Ag presentation, and activation of Ag-specific T cells are completed before LC apoptosis.

To investigate which caspase pathway (type I or type II) is activated in LCs during T cell interaction, we used selective caspase inhibitor peptides and tested them on the XS-GFP line. We reasoned that the caspase-9 inhibitor peptide (LEHD) would only have blocking activity if LCs were type II cells while all other caspase inhibitors should block regardless. After peptide preincubation of XS-GFP, 3A9 T hybridoma cells were added at a 1:4 ratio then incubated additionally overnight. Flow cytometric analysis on gated GFP-positive cells revealed that during interaction with T cells, DC apoptosis was markedly blocked with pan-caspase inhibitors, as well as the specific caspase inhibitor for caspase 3, and partially blocked with inhibitors with specificity for either caspase-8 or -9 (Fig. 2). Fig. 2,A shows a significant decrease in annexin V-positive cells for all caspase inhibitors tested; however, peptide-induced toxicity did occur in some samples, as indicated by an increase in single-positive 7-AAD staining. This peptide-induced toxicity pattern of staining was also present in parallel control cultures (without T cells; data not shown). Fig. 2,B shows the compilation of four independent experiments (mean ± SEM) adjusting for background toxicity and including the quantification of LC loss that occurs, in addition to detecting apoptosis (as shown in Fig. 2 A) of the remaining cells per sample (see Materials and Methods for details). The finding that inhibitors for caspase-9 have an impact on T cell-mediated LC apoptosis suggests that LCs use, at least partially, the type II caspase activation cascade.

FIGURE 2.

Caspase inhibition of T cell-induced apoptosis of the LC line XS106. A, GFP-positive XS106 B1.1 cells analyzed for apoptosis by Annexin VPE and 7-AAD staining. XS106 B1.1 cells were pulsed with or without HEL Ag overnight and mixed with either 3A9 T cells or unlabeled XS106 as a control. One hour before mixing cells, they were preincubated with the following caspase inhibitors: 100 μM Z-VAD-FMK, 200 μM Z-DEVD-FMK, 150 μM Z-IETD-FMK, and 300 μM Ac-LEHD-CHO and then continuously cultured in the presence of these inhibitors. Apoptosis levels were assessed after 16–20 h by flow cytometry. The listed percent control LC values indicate the background-adjusted percentage of remaining XS106 B1.1cells. B, Percent inhibition summary. The average ± SEM from four independent experiments (see Materials and Methods for calculations).

FIGURE 2.

Caspase inhibition of T cell-induced apoptosis of the LC line XS106. A, GFP-positive XS106 B1.1 cells analyzed for apoptosis by Annexin VPE and 7-AAD staining. XS106 B1.1 cells were pulsed with or without HEL Ag overnight and mixed with either 3A9 T cells or unlabeled XS106 as a control. One hour before mixing cells, they were preincubated with the following caspase inhibitors: 100 μM Z-VAD-FMK, 200 μM Z-DEVD-FMK, 150 μM Z-IETD-FMK, and 300 μM Ac-LEHD-CHO and then continuously cultured in the presence of these inhibitors. Apoptosis levels were assessed after 16–20 h by flow cytometry. The listed percent control LC values indicate the background-adjusted percentage of remaining XS106 B1.1cells. B, Percent inhibition summary. The average ± SEM from four independent experiments (see Materials and Methods for calculations).

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In type II cells, caspase-8 does not directly cleave procaspase-3, rather it cleaves the protein Bid, generating the active form called truncated Bid. Truncated Bid translocates to the mitochondria where it activates Bak and Bax proapoptotic BCL-2 family proteins, culminating in the activation of caspase-9 and resulting in apoptosis (18). To verify the type II character of the apoptosis pathway in LCs, we looked for the presence of Bid in protein lysates of our LC lines, as detected with a full-length Bid-specific mAb in Western blots (Fig. 3). As expected, Bid was detected in lysates from WT spleen but not from the splenocytes of Bid KO mice. Bid protein was not detected in T cells, such as 3A9 T hybridoma cells or naive CD4 T cells, known to be type I cells. However, high levels of Bid protein were observed in our LC line. LC Bid protein could be activated (cleaved) by staurosporine treatment (Fig. 3,A) and, significantly, after Ag presentation to T cells (Fig. 3, B and C), reducing the Bid-specific signal by 99 and 43% in the presence of 3A9 T hybridoma and naive CD4 T cells, respectively (Fig. 3 D). (Activation is detected by disappearance of full-length Bid due to activation-induced cleavage of the Bid-specific mAb epitope.) The specific expression of Bid in LCs and the demonstration that Bid is cleaved in response to apoptotic stimuli provides further evidence that LCs use the type II caspase cascade.

FIGURE 3.

Bid activation in DC upon Ag-specific interaction with T cells. A, Bid activation in DC. To test LC Bid cleavage, we used staurosporine-induced release of cytochrome c, which induces caspase-3-mediated Bid cleavage. Cells were cultured overnight with or without staurosporine (500 nM). Cell lysate was obtained, and 100 μg of protein was loaded per sample and analyzed by Western blot with the indicated Ab. Spleen cells from Bid KO mice are loaded as a control. B, Ag-specific T cell-mediated activation of Bid cleavage by 3A9 hybridoma T cells. T cells were mixed 4:1 with DC pulsed or unpulsed with HEL Ag. Protein lysates from half of the indicated culture were loaded for each sample (∼100 μg) and analyzed by Western blot. C, Splenic naive CD4 T cells induce Bid cleavage in DC. CD4 T cells purified from 3A9 transgenic mice were mixed 4:1 with DC pulsed with or without HEL Ag. Half of the total cell culture input was loaded (∼50 μg). D, Densitometry of Bid as a percentage of actin band intensities quantifies the level of Bid activation. Values indicate the percent loss of Bid intensity (and therefore activation) as compared with paired controls.

FIGURE 3.

Bid activation in DC upon Ag-specific interaction with T cells. A, Bid activation in DC. To test LC Bid cleavage, we used staurosporine-induced release of cytochrome c, which induces caspase-3-mediated Bid cleavage. Cells were cultured overnight with or without staurosporine (500 nM). Cell lysate was obtained, and 100 μg of protein was loaded per sample and analyzed by Western blot with the indicated Ab. Spleen cells from Bid KO mice are loaded as a control. B, Ag-specific T cell-mediated activation of Bid cleavage by 3A9 hybridoma T cells. T cells were mixed 4:1 with DC pulsed or unpulsed with HEL Ag. Protein lysates from half of the indicated culture were loaded for each sample (∼100 μg) and analyzed by Western blot. C, Splenic naive CD4 T cells induce Bid cleavage in DC. CD4 T cells purified from 3A9 transgenic mice were mixed 4:1 with DC pulsed with or without HEL Ag. Half of the total cell culture input was loaded (∼50 μg). D, Densitometry of Bid as a percentage of actin band intensities quantifies the level of Bid activation. Values indicate the percent loss of Bid intensity (and therefore activation) as compared with paired controls.

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A stringent proof for the role of Bid in regulating LC apoptosis was investigated using Bid KO mice. We examined whether a loss-of-function mutation in the Bid gene could affect the immune response to topical Ags using a number of model systems.

C57BL/6 (B6) Bid KO mice and WT control B6 mice were sensitized on the abdomen and challenged 5 days later on the ears with the hapten DNFB. Ear-swelling responses were accelerated in Bid KO mice and enhanced >2-fold over WT mouse responses (Fig. 4,A). After 5 days postchallenge, cellular infiltrates could still be seen in epidermal layers of Bid KO, whereas cellular infiltrates were mainly seen only in the dermal layer of WT specimens (Fig. 4,B). The marked augmentation of the contact sensitivity response seen by Bid KO mice could not be explained by a difference in LC density in this mutant strain (Fig. 4,C). Examination of untreated epidermal sheets from age-matched Bid KO and WT mice revealed no significant difference in LC densities, and numbers were consistent with previously published values for B6 mice (WT, 675/mm2 ± SE 143; Bid KO, 668/mm2 ± SE 178) (35). Neither could it be explained by a difference in LC migration rate or LC culture viability. We examined the numbers of LCs, identified as Langerin-positive cells, that migrated from WT and Bid KO skin cultures and their viability over time and found no significant differences (Fig. 4, D and E).

FIGURE 4.

Enhanced contact sensitization in Bid null mice. A, Kinetics of the swelling response. Four age-matched mice per group of WT or Bid KO were sensitized with vehicle only or 0.5% DNFB on the stomach and paws of each mouse. On day 5, baseline measurement of ear thickness was taken for each ear. Mice were subsequently challenged with 0.2% DNFB applied on dorsal and ventral sides of the ears (day 0 is day of challenge). Ear thickness measurements were taken thereafter on the following days, as shown. Ear thickness shown are that over baseline measurements for each ear. All significant differences are noted in the following manner: ∗, p < 0.05; ∗∗, p < 0.01; and ∗∗∗, p < 0.001. This is representative of three similar experiments. B, Histological comparison of WT and Bid epidermal sections. Paraffin-embedded sections of ear specimens obtained on day 0 and day 5 posthapten challenge were stained by H&E. Magnifications of ×10 upper panel, ×20 middle panels, and ×40 in lower panels are shown for WT and Bid contact-sensitized ears. The bar length indicates 100 μM. C, LC densities are equivalent in Bid null and WT mice. Whole mounts of epidermal sheets from age-matched Bid KO and WT mice were stained with FITC-anti MHC class II or isotype control (data not shown) and examined by fluorescence microscopy. Digital images at ×20 magnification and the number of MHC class II-positive cells were determined. D and E, LC migration and viability are similar for WT and Bid KO cells during ear skin culture. Ear skin samples were prepared for culture after tape stripping six mice per group. Ear skin halves were transferred to fresh wells daily for 4 days. Pooled samples per day of culture were counted and stained with DC specific Abs (FITC-CD11c, PE-DEC205, APC-biotin-Langerin, and isotype controls) and 7-AAD. Flow cytometry was performed to assess phenotype and viability. D, Migration kinetics. Total CD11c positive (open symbols) and Langerin positive (closed symbols) cells per ear culture of WT (squares) and Bid KO (circles) mice were calculated based on total cell counts and percent positive cells. B, Migratory cell viability. The percentage of 7-AAD-negative, CD11c-positive cells that migrated from ear skin samples each day are shown. The total viable LC yields from WT (1.27 × 105) and Bid KO (1.20 × 105) ears were similar.

FIGURE 4.

Enhanced contact sensitization in Bid null mice. A, Kinetics of the swelling response. Four age-matched mice per group of WT or Bid KO were sensitized with vehicle only or 0.5% DNFB on the stomach and paws of each mouse. On day 5, baseline measurement of ear thickness was taken for each ear. Mice were subsequently challenged with 0.2% DNFB applied on dorsal and ventral sides of the ears (day 0 is day of challenge). Ear thickness measurements were taken thereafter on the following days, as shown. Ear thickness shown are that over baseline measurements for each ear. All significant differences are noted in the following manner: ∗, p < 0.05; ∗∗, p < 0.01; and ∗∗∗, p < 0.001. This is representative of three similar experiments. B, Histological comparison of WT and Bid epidermal sections. Paraffin-embedded sections of ear specimens obtained on day 0 and day 5 posthapten challenge were stained by H&E. Magnifications of ×10 upper panel, ×20 middle panels, and ×40 in lower panels are shown for WT and Bid contact-sensitized ears. The bar length indicates 100 μM. C, LC densities are equivalent in Bid null and WT mice. Whole mounts of epidermal sheets from age-matched Bid KO and WT mice were stained with FITC-anti MHC class II or isotype control (data not shown) and examined by fluorescence microscopy. Digital images at ×20 magnification and the number of MHC class II-positive cells were determined. D and E, LC migration and viability are similar for WT and Bid KO cells during ear skin culture. Ear skin samples were prepared for culture after tape stripping six mice per group. Ear skin halves were transferred to fresh wells daily for 4 days. Pooled samples per day of culture were counted and stained with DC specific Abs (FITC-CD11c, PE-DEC205, APC-biotin-Langerin, and isotype controls) and 7-AAD. Flow cytometry was performed to assess phenotype and viability. D, Migration kinetics. Total CD11c positive (open symbols) and Langerin positive (closed symbols) cells per ear culture of WT (squares) and Bid KO (circles) mice were calculated based on total cell counts and percent positive cells. B, Migratory cell viability. The percentage of 7-AAD-negative, CD11c-positive cells that migrated from ear skin samples each day are shown. The total viable LC yields from WT (1.27 × 105) and Bid KO (1.20 × 105) ears were similar.

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Because T cells have been shown to use the type I pathway of apoptosis by extrinsic signals, the increased immune response seen in Bid KO mice was not likely to be due to aberrant Bid KO T cell proliferation. However, the dynamic expression patterns of Bid in cells and tissues have not been fully characterized. Therefore, to rule out a role for aberrant T cell responses in Bid mice, we directly tested the response of naive WT T cells to topical immunization in Bid KO and WT mice in adoptive transfer experiments. The protein Ag OVA or PBS was painted on tape-stripped ears, and thereafter, ∼1 million CFSE-labeled OT-2-transgenic CD4 T cells were injected i.v. via tail vein. Proliferation (reduction in CFSE staining) of OT-2 T cells was examined on day 3 from draining auricular lymph nodes (Fig. 5,A). No significant differences in weight or cell numbers were seen in the lymph nodes or spleens from PBS control-treated WT and Bid KO mice, but modest increases were noted for immunized Bid KO spleen (Table I). While both strains supported an efficient response by topical immunization, OT-2 T cell expansion was 1.46-fold greater in Bid KO mice (WT 2.6 ± 0.3% vs Bid KO 3.8 ± 0.2%; p < 0.003) (Fig. 5,A), and a significant increase in the number of T cells that divided seven times was detected in draining lymph nodes (WT 15 ± 3.1% vs Bid KO 26 ± 2.2%; p < 0.05) (Fig. 5,B). Furthermore, a significant increase in the number of OT-2 T cells that had divided and exited to the periphery was also detected in spleen from Bid KO mice (WT 33 ± 1.8% vs Bid KO 59 ± 3.3%, p < 0.001) (Fig. 5 C). The accumulation of dividing cells in lymph node and spleen suggest that Bid KO mice do not induce abortive activation of OT-2 T cells (36). Furthermore, these results suggest that enhanced T cell responsiveness from Bid KO mice may not be due to aberrant T cell proliferation but due to mechanisms that activate normal T cells.

FIGURE 5.

Enhanced OT-2 T cell responses in Bid null mice. WT and Bid KO mice were topically immunized by application of 25 μg of OVA onto tape-stripped ear surfaces. CD4-purified CFSE-labeled OT-2 cells (1.5 × 106) were injected into topically immunized mice via tail vein. Draining lymph node cells and spleen cells were harvested from mice on day 3 and analyzed for dilution of CFSE within the TCR Vb5-positive population. A, Increased rate of proliferation by OT-2 T cell in Bid mice. CFSE log fluorescence from gated OT-2 cells transferred to control Bid KO (or WT, data not shown) mice (left panels), immunized WT mice (middle panels), or immunized Bid KO mice (right panels) are shown. OT-2 gate was set as shown and represented 0.05, 2.6, and 3.8% of total lymph node cells for immunized Bid KO, WT-OVA, and Bid KO-OVA mice, respectively. Values of p < 0.003 comparing WT-OVA and Bid KO-OVA responses, n = 3. B, Enhanced proliferation and emigration to spleen by OT-2 T cells in Bid KO mice. Lymph nodes (left panel) and spleen (right panel) were harvested from triplicate mice per group. OT-2 cells were analyzed as shown in A, and the percentage of cells present in each division region, based on fluorescence intensity, was determined. The average value and SEM are shown for each group (▦ for WT, ▪ for Bid KO). Values of p are represented by asterisks as follows: ∗, p < 0.05; ∗∗, p < 0.01; and ∗∗∗, p < 0.001.

FIGURE 5.

Enhanced OT-2 T cell responses in Bid null mice. WT and Bid KO mice were topically immunized by application of 25 μg of OVA onto tape-stripped ear surfaces. CD4-purified CFSE-labeled OT-2 cells (1.5 × 106) were injected into topically immunized mice via tail vein. Draining lymph node cells and spleen cells were harvested from mice on day 3 and analyzed for dilution of CFSE within the TCR Vb5-positive population. A, Increased rate of proliferation by OT-2 T cell in Bid mice. CFSE log fluorescence from gated OT-2 cells transferred to control Bid KO (or WT, data not shown) mice (left panels), immunized WT mice (middle panels), or immunized Bid KO mice (right panels) are shown. OT-2 gate was set as shown and represented 0.05, 2.6, and 3.8% of total lymph node cells for immunized Bid KO, WT-OVA, and Bid KO-OVA mice, respectively. Values of p < 0.003 comparing WT-OVA and Bid KO-OVA responses, n = 3. B, Enhanced proliferation and emigration to spleen by OT-2 T cells in Bid KO mice. Lymph nodes (left panel) and spleen (right panel) were harvested from triplicate mice per group. OT-2 cells were analyzed as shown in A, and the percentage of cells present in each division region, based on fluorescence intensity, was determined. The average value and SEM are shown for each group (▦ for WT, ▪ for Bid KO). Values of p are represented by asterisks as follows: ∗, p < 0.05; ∗∗, p < 0.01; and ∗∗∗, p < 0.001.

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Table I.

Lymphoid organ weights and cell yields from WT and Bid KO mice

OrganaImmunizebWeight (mg)Cell Yield (×106)
WTBid KOWTBid KO
LN PBS 6.3 ± 1.2 5.6 ± 0.9 8.4 ± 1.8 9.4 ± 3.8 
 OVA 8.8 ± 1.0 10.1 ± 3.8 17.4 ± 5.6 17.3 ± 4.1 
pc  NS  NS  
Spleen PBS 93.2 ± 4.0 95.4 ± 26.9 72.8 ± 9.9 87.0 ± 21.7 
 OVA 93.9 ± 5.9 116.0 ± 7.9 70.3 ± 22.0 87.9 ± 20.9 
p≤  0.05  0.02  
OrganaImmunizebWeight (mg)Cell Yield (×106)
WTBid KOWTBid KO
LN PBS 6.3 ± 1.2 5.6 ± 0.9 8.4 ± 1.8 9.4 ± 3.8 
 OVA 8.8 ± 1.0 10.1 ± 3.8 17.4 ± 5.6 17.3 ± 4.1 
pc  NS  NS  
Spleen PBS 93.2 ± 4.0 95.4 ± 26.9 72.8 ± 9.9 87.0 ± 21.7 
 OVA 93.9 ± 5.9 116.0 ± 7.9 70.3 ± 22.0 87.9 ± 20.9 
p≤  0.05  0.02  
a

LN, Auditory lymph node. Organs were harvested on day 3 after immunization.

b

Groups of mice (n = 3) were immunized by tape stripping and painted with PBS (control) or 25 μg of OVA per car side (see Materials and Methods).

c

Values of p were determined using a one tailed, paired Student’s t-test comparing OVA immunized WT and Bid KO values. No significant differences were found in PBS-treated groups.

To directly test whether Bid KO LCs are apoptosis resistant, migratory cDCs were cultured from the ear skin of both strains in the presence of OVA and subsequently cocultured with OT-2 or irrelevant 3A9 CD4 T cells overnight. Apoptosis staining was examined by flow cytometry and gated on double-positive cells for CD11c and DEC-205, and the LC population was identified as Langerin positive. (The LC-specific Ab used reacts with Langerin molecules on the cell surface, and stained 95% of the gated population (Fig. 6,A)). We found that the viability of LC after 2-day culture for all samples was 75%, indicating the inevitable fate of death based on intrinsic signals (11). However, the hastening of LC death by extrinsic T cell-derived signals was demonstrated in cells from WT mice demonstrating both a loss of Langerin positive cells (from 23 to 16%) (Fig. 6,A) and an increase in apoptotic LCs (increasing from 26 to 42% 7-AAD-positive cells) detected in cultures with OT-2 T cells. (Fig. 6,B). In contrast, OT-2 T cells did not affect Bid KO LC viability since both the percentage of Langerin-positive cells and viability of LCs was similar to cultures with 3A9 T cells (Fig. 6,B). Despite the difference in susceptibility to apoptosis, LCs from both strains supported maximal activation of OT-2 T cells (Fig. 6,C). A slight increase in the percentage of large (by forward angle side scatter) CD69-positive T cells was observed (59 vs 65% for OT-2 T cells activated by WT vs Bid KO LCs, respectively). No activation of irrelevant 3A9 CD4 T cells was detected. Resistance to T cell-mediated, Ag-specific apoptosis was consistently observed in Bid KO cDCs (Table II). These results suggest that part of the mechanism for enhanced T cell responsiveness in vivo may due to an increased duration of Ag-presenting function by apoptosis-resistant Bid KO cDCs.

FIGURE 6.

Bid KO LCs are resistant to Ag-specific T cell-mediated apoptosis. cDCs migrated from ear skin in the presence of indicated Ag over 2 days, harvested, and mixed with purified OT-2 cells at a 1:4 ratio. After 20 h, cultures were subjected to flow cytometric analysis after staining with fluorochrome-conjugated Abs and dyes as follows: FITC-CD11c, PE-CD205, 7-AAD, and APC-biotin-Langerin. A, The percentage of Langerin-positive cells is shown, indicating the remaining LCs in each mixed culture. B, Double-positive CD11c,CD205 cells were gated and analyzed for Langerin and 7-AAD uptake. Numbers displayed in the right quadrant indicate the percentage of 7-AAD-positive cells. C, Cells falling in the small lymphocyte gate based on light scatter properties were examined for early activation marker CD69, with percentage of positive cells displayed; one of three similar experiments FALS, Forward angle light scatter.

FIGURE 6.

Bid KO LCs are resistant to Ag-specific T cell-mediated apoptosis. cDCs migrated from ear skin in the presence of indicated Ag over 2 days, harvested, and mixed with purified OT-2 cells at a 1:4 ratio. After 20 h, cultures were subjected to flow cytometric analysis after staining with fluorochrome-conjugated Abs and dyes as follows: FITC-CD11c, PE-CD205, 7-AAD, and APC-biotin-Langerin. A, The percentage of Langerin-positive cells is shown, indicating the remaining LCs in each mixed culture. B, Double-positive CD11c,CD205 cells were gated and analyzed for Langerin and 7-AAD uptake. Numbers displayed in the right quadrant indicate the percentage of 7-AAD-positive cells. C, Cells falling in the small lymphocyte gate based on light scatter properties were examined for early activation marker CD69, with percentage of positive cells displayed; one of three similar experiments FALS, Forward angle light scatter.

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Table II.

Summary of in vitro apoptosis detected in LCs from WT and Bid KO mice

Expt.aPercent-Specific LC LossbPercent-Specific ApoptosiscPercent-Specific LC Death
WTdBid KOWTBd KOWTBd KO
46.7 0.5 21.0 2.0 67.7 2.5 
47.4 5.0 12.7 3.0 60.1 8.0 
30.4 7.4 16.0 0.0 46.4 7.4 
Averagef 41.5 4.3 16.6 1.7 58.1 6.0 
SD 9.6 3.5 4.2 1.5 10.8 3.0 
p<e  0.02  0.02  0.01 
Expt.aPercent-Specific LC LossbPercent-Specific ApoptosiscPercent-Specific LC Death
WTdBid KOWTBd KOWTBd KO
46.7 0.5 21.0 2.0 67.7 2.5 
47.4 5.0 12.7 3.0 60.1 8.0 
30.4 7.4 16.0 0.0 46.4 7.4 
Averagef 41.5 4.3 16.6 1.7 58.1 6.0 
SD 9.6 3.5 4.2 1.5 10.8 3.0 
p<e  0.02  0.02  0.01 
a

Results from three independent experiments are shown. cDCs were gated for each experiment as follows: Expt. no. I, I-A; 2, CD11c; 3, Langerin (CD207). WT or Bid KO LCs comprises the majority of cDCs isolated from ear skin cultures, as described in Materials and Methods.

b

Percent-specific LC loss is calculated as the difference in percent of LCs remaining after overnight culture in the presence of OVA Ag and relevant OT-2 vs irrelevant 3A9 naïve purified CD4 T cells.

c

Percent-specific Apoptosis is calculated as the difference in apoptosis detected by annexin V and/or 7-AAD staining of remaining DCs after overnight culture in the presence of OT-2 or 3A9 T cells.

d

Percent-specific LC death = percent-specific LC loss + percent-specific Apoptosis.

e

Values of p were determined using a one-tailed, paired Student’s t test.

f

The average values for all three experiments are shown in bold.

To determine the contribution of the augmented immune response by cDCs, as opposed to contributing factors from other tissues and cells in Bid KO mice, we isolated migratory cDCs (comprising LCs and dDCs) from WT and Bid KO skin explant cultures and tested their stimulatory capacity in adoptive transfer experiments. WT mice containing transferred CFSE-labeled OT-2 T cells were inoculated with equivalent numbers of OVA-pulsed Bid KO or WT migratory cells into WT recipient footpads. All I-A-positive cells expressed both CD11c and CD205 (data not shown). Draining popliteal lymph nodes from OVA-pulsed Bid KO cDCs were 1.8-fold larger than those obtained from OVA-pulsed WT cDCs (233 ± 44 vs 129 ± 14 μg/g mouse weight, respectively; p < 0.05). Flow cytometric analysis revealed that CD4 T cell proliferation responses were Ag specific, observed only in CFSE stained cells that were TCR Vb5 positive (Fig. 7,A). OT-2 cells responded to OVA-pulsed Bid KO cDCs with both an increase in the rate, as indicated by the faction of cells that divided six or more times (28 ± 2.7% for Bid KO vs 13 ± 0.3% for WT; p < 0.05) and the total number of T cells that responded, as indicated by the percentage of CD69-positive OT-2 cells in nondividing fraction plus the percentage of proliferating cells (93 ± 0.3% for Bid KO vs 86 ± 0.1% for WT; p < 0.001) (Fig. 7,C). In addition, a greater percentage of mature cells (CSFElow CD69neg) was generated by Bid KO cDC (Fig. 7, C and E). Surprisingly, in this particular experiment, fewer I-A-positive cDCs comprised the population of migratory cells from the Bid KO cultures as compared with WT cultures (58 vs 80%, respectively with equivalent mean fluorescence channels; data not shown), indicating that Bid KO cDCs were highly potent APCs. (The yield of migratory cDCs (I-A, CD11c, and Langerin positive) obtained from each mouse strain differed within some experiments, but these differences revealed no statistically significant trend when all experiments were analyzed.) To correlate increased proliferation with effector function, we examined expanded lymph node and splenic OT-2 T cells for Th1 and Th2 cytokine secretion. Again, draining lymph nodes from Bid KO cDC-injected mice were significantly larger and demonstrated greater expansion of CD45.1 OT-2 T cells than control mice (Fig. 8, A and C). The percentage of IFN-γ-secreting OT-2 cells was modestly increased in lymph node cells (Fig. 8, A and B) but dramatically increased in restimulation cultures of spleen cells from Bid KO cDC-injected mice (Fig. 8, B and E) as compared with control mice. IL-4 secretion was not detectable for any of the immunization regimes. These results show that increased numbers of mature Th1 effector cells are induced by Bid KO cDCs within WT recipients. Thus, enhanced T cell responses did not require the environment of the Bid KO mouse but was intrinsic to the function of Bid KO cDCs.

FIGURE 7.

Inoculated Bid KO cDCs promote augmented T cell activation in vivo. Bid KO and WT mouse ears were tape-stripped and painted with 25 μg of OVA or HEL in PBS. After an hour, mice were sacrificed, and ear specimens were harvested for explant culture. Migratory cDCs were cultured from specimens over 48 h in the presence of OVA (100 μg/ml) or HEL (100 μg/ml). One million CFSE-labeled CD4 OT-2 T cells were injected via tail vein to WT mice (in this experiment, 50% of the CFSE-labeled CD4 cells were OT-2 TCR negative, as indicated by absence of TCR Vb5 staining—and served as an internal specificity control), and cDCs (5 × 104) were inoculated into the footpad the same day. Popliteal lymph nodes were harvested from individual mice (three per group) and weighed before processing for TCR and CD69 staining. Flow cytometric analysis plots of the gate used for CFSE-TCR Vb5 cells (A), the level of CFSE dilution (indicating cell divisions) (B), and the level of T cell activation/maturation-indicated by transient CD69 up then down-regulation (C) are shown. B, Mean channel fluorescence values are shown in parentheses. The percentage of cells that divided five times or more is displayed over the lower bar. D, Significant differences in the rate of proliferation and E, mature cell phenotype between WT and Bid KO. Percentage of OT-2 cells found in each division from three mice per group is shown (□ WT, ▪ Bid KO). Students’ t test p values are depicted as follows: ∗, p < 0.05; ∗∗, p < 0.001. CFSE T cells showed no proliferation or activation in mice receiving HEL-pulsed LCs and represented 0.01% of total lymph node cells (data not shown). OT-2 T cells expanded from 0.55 to 0.73% of total lymph node cells in mice receiving OVA-pulsed cDCs from WT vs Bid KO mice, respectively (shown in A); p < 0.05, n = 3.

FIGURE 7.

Inoculated Bid KO cDCs promote augmented T cell activation in vivo. Bid KO and WT mouse ears were tape-stripped and painted with 25 μg of OVA or HEL in PBS. After an hour, mice were sacrificed, and ear specimens were harvested for explant culture. Migratory cDCs were cultured from specimens over 48 h in the presence of OVA (100 μg/ml) or HEL (100 μg/ml). One million CFSE-labeled CD4 OT-2 T cells were injected via tail vein to WT mice (in this experiment, 50% of the CFSE-labeled CD4 cells were OT-2 TCR negative, as indicated by absence of TCR Vb5 staining—and served as an internal specificity control), and cDCs (5 × 104) were inoculated into the footpad the same day. Popliteal lymph nodes were harvested from individual mice (three per group) and weighed before processing for TCR and CD69 staining. Flow cytometric analysis plots of the gate used for CFSE-TCR Vb5 cells (A), the level of CFSE dilution (indicating cell divisions) (B), and the level of T cell activation/maturation-indicated by transient CD69 up then down-regulation (C) are shown. B, Mean channel fluorescence values are shown in parentheses. The percentage of cells that divided five times or more is displayed over the lower bar. D, Significant differences in the rate of proliferation and E, mature cell phenotype between WT and Bid KO. Percentage of OT-2 cells found in each division from three mice per group is shown (□ WT, ▪ Bid KO). Students’ t test p values are depicted as follows: ∗, p < 0.05; ∗∗, p < 0.001. CFSE T cells showed no proliferation or activation in mice receiving HEL-pulsed LCs and represented 0.01% of total lymph node cells (data not shown). OT-2 T cells expanded from 0.55 to 0.73% of total lymph node cells in mice receiving OVA-pulsed cDCs from WT vs Bid KO mice, respectively (shown in A); p < 0.05, n = 3.

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FIGURE 8.

Bid KO cDCs promote increased OT-2 cell expansion and function in lymph node and spleen. cDCs from topically immunized ear skin explants were injected (3 × 105) into the footpad of WT recipient mice (three per group) containing CD45.1 allotypic OT-2 T cells (2 × 106). Control mice were topically immunized on ears (see Materials and Methods). A, Lymph node cells were harvested after 4 days and stained for CD45.1 to reveal OT-2 T cells (upper panel) that were gated for analysis of cytokine staining (lower panel). B, Spleen cells harvested after 4 days were restimulated with 100 μg/ml OVA in vitro for 3 days before CD45.1 and cytokine staining. C, Lymph node weights are increased in mice receiving Bid KO cDCs. Lymph nodes from mice receiving PBS (□) or immunization by topical OVA or by cDC transfer (▪) were weighed and the mean ± SD per group calculated. ∗, p < 0.05. D and E, Bid KO cDCs promote increased numbers of IFN-γ-producing cells in lymph node (D) and spleen (E). The number of IL-4 (□)- or IFN-γ (▪)-producing OT-2 cells per million CD3 T cells was calculated for each individual mouse and mean ± SD determined per group. ∗, p < 0.05. Representative density plots and statistical analyses are shown for one of three independent experiments.

FIGURE 8.

Bid KO cDCs promote increased OT-2 cell expansion and function in lymph node and spleen. cDCs from topically immunized ear skin explants were injected (3 × 105) into the footpad of WT recipient mice (three per group) containing CD45.1 allotypic OT-2 T cells (2 × 106). Control mice were topically immunized on ears (see Materials and Methods). A, Lymph node cells were harvested after 4 days and stained for CD45.1 to reveal OT-2 T cells (upper panel) that were gated for analysis of cytokine staining (lower panel). B, Spleen cells harvested after 4 days were restimulated with 100 μg/ml OVA in vitro for 3 days before CD45.1 and cytokine staining. C, Lymph node weights are increased in mice receiving Bid KO cDCs. Lymph nodes from mice receiving PBS (□) or immunization by topical OVA or by cDC transfer (▪) were weighed and the mean ± SD per group calculated. ∗, p < 0.05. D and E, Bid KO cDCs promote increased numbers of IFN-γ-producing cells in lymph node (D) and spleen (E). The number of IL-4 (□)- or IFN-γ (▪)-producing OT-2 cells per million CD3 T cells was calculated for each individual mouse and mean ± SD determined per group. ∗, p < 0.05. Representative density plots and statistical analyses are shown for one of three independent experiments.

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The results from this study support the hypothesis that an additional layer of immune regulation occurs after successful Ag presentation by LCs to Ag-specific CD4 T cells. During the successful interaction with LCs, T cells provide signals for further activation of LCs (Ref. 6 and our unpublished results) and subsequent induction of LC apoptosis. The death of incoming LCs may be part of a mechanism for lymph node cells to sense the peripheral load of Ag and may serve to disperse LC-incorporated Ag to endogenous lymph node DC subsets for cross-presentation. This CD4 T cell-mediated apoptosis of LCs occurs via type II apoptotic signals, requiring the BH3-only domain protein Bid, for linking extrinsic death receptor signals to mitochondrial mechanisms of apoptosis. The physiological significance of this observation is demonstrated by the markedly enhanced capacity to augment Ag-specific T cell function in vivo. This was observed for multiple protocols of epicutaneous immunization in Bid KO mice or by inoculation of cDCs from Ag-painted Bid KO mice into WT recipients. Thus, the data suggest that the longevity of activated, Ag-pulsed LCs in vivo directly correlates with the magnitude of the immune response that can be generated.

We found that peptide inhibitors for caspase-8 or -9 could only partially inhibit T cell-mediated LC apoptosis, whereas pan-caspase or “effector” caspase 3 inhibitors almost completely blocked LC apoptosis. The inability to completely inhibit “initiator” caspases 8 and 9 may due to a number of reasons: 1) the amount of caspase protein may be too high to allow saturation by the inhibitor peptide at the concentrations used, 2) the caspases may be molecularly sequestered from inhibitory effects of the peptide, or 3) for caspase-9, the type I or II apoptosis pathway used may not be absolute. In the later case, perhaps both pathways may be used simultaneously in the same cell or used independently in different LC subsets depending on their interaction with the T cell. Therefore, the classification of LC as a type II cell by this criterion may not be strictly applied. Nevertheless, the type II cascade is in place within LC, supported by our finding that high levels of the Bid protein is expressed in our LC line and specifically activated after Ag-specific T cell interaction. Furthermore, the role of Bid in primary LC was studied in Bid KO mice, and the data from those studies indicate that if a compensatory type I pathway is used by LC, it is not complete. Thus, taken together, these data support our hypothesis that there is a preference for the use of the type II cascade in CD4 T cell-mediated LC apoptosis.

Our results are consistent with a number of studies that have identified a critical role for the Bcl-2 family genes, bcl-xL and bcl-2, in maintaining DC longevity. However, studies on the impact of Bcl-2 family gene transfection do not indicate whether intrinsic, extrinsic, or both apoptotic pathways are affected. DCs from CD11c-promoter-driven bcl-2 transgenic mice display increased longevity in vitro and in vivo and induce augmented immune responses when transferred into WT recipients (21). Activated T cells secrete TNF-related activation-induced cytokine (TRANCE) and express CD40L, molecules that have been shown to improve survival of cultured bone-derived or splenic DCs, and correlate with an increase in Bcl-xL and Bcl-2 expression, respectively (37, 38). Whether cDCs or LCs are similar to conventional DCs in response to TRANCE or CD40L needs to be addressed. While our studies indicate LCs undergo cell death after prolonged interaction with Ag-specific T cells, these observations do not exclude a role for TRANCE- and CD40L-induced survival signals early during T cell interaction—promoting prolonged cell-cell contact that is required for successful CD4 T cell activation and differentiation (39, 40). Alternatively, different DC subsets may respond differently to activated T cell signals and cytokines. Other studies using gene gun-mediated DNA-based vaccines suggest that cDCs do rely on bcl-xL and bcl-2 gene expression for generating or enhancing such cutaneous vaccines (41, 42). Our studies support and extend those studies, showing that the mitochondrial pathway is essential for mediating extrinsic activation-induced apoptosis signals from Ag-specific T cells and that it is Bid dependent.

The nature of the extrinsic signal(s) provided by T cells is currently under investigation. While it is known that Ag bearing LCs from lpr-mutant (Fas null) mice demonstrate increased longevity (43) and resist apoptosis in vivo after hapten (FITC) painting or in a graft-vs-host disease model (7), we have been unsuccessful in blocking in vitro T cell-mediated apoptosis of LCs using neutralizing agents for either Fas ligand or TRAIL (data not shown; D. T. Warren, L. Dandridge, J. Genebriera, S. Pradhan, C. A. Elmets, and L. Timares, manuscript in preparation). This may indicate that either the LC-T cell interface may be impenetrable to such soluble neutralizing agents or that other mediators of apoptosis are involved and remain to be identified.

Very little is known regarding Bid expression and use in normal tissues. A limited immunohistochemical survey was done using anti-Bid sera (44). The strongest staining was detected in neuronal cells, stratified squamous epithelium, and in short-lived leukocytes (germinal center cells, granulocytes, macrophages), whereas monocytes, immature bone marrow cells, and cortical thymocytes were reported as negative. This is consistent with our inability to detect significant Bid protein levels in splenic T cells or T hybridoma lines. Definitive studies on the expression of Bid in DC subsets have not been performed. Thus, this is the first demonstration that a DC subset, specifically LCs, expresses and activates Bid in response to extrinsic apoptotic stimuli. Investigations of the expression and use of Bid by different DC subsets in response to various stimuli are currently underway.

Because Bid has also been detected in differentiated squamous keratinocytes (44), we considered that there may be aberrations in keratinocyte biology that might contribute to the enhanced ear swelling observed in Bid KO mice. Therefore, we performed histochemical analysis of resting and contact sensitized ear specimens. Extensive pathological examination comparing Bid and WT epidermis revealed normal epidermal morphology in resting tissues. However, while both WT and Bid KO specimens demonstrated a similar influx infiltrating cells affecting the dermal and epidermal layers during the swelling response peak at day 3, the extended swelling response, seen on day 5 for Bid KO, when the swelling response diminished for WT, correlated with an increased incidence of infiltrating cells within the epidermis. These findings suggest that prolonged active immunity continued in Bid KO as compared with WT. Alternatively, Bid KO inflammatory cells, reportedly shown to express Bid (i.e., granulocytes and macrophages) may also play a role in prolonging the local inflammatory and/or lymphocyte recruitment response due to extended life spans or unresponsive down regulatory mechanisms. However, control vehicle sensitized Bid mice treated with a challenge dose of hapten did not develop swelling responses. This observation indicates that differences in inflammatory mechanisms alone do not contribute to the enhanced contact sensitization response in Bid KO mice. It is likely that multiple components play a role, and additional experiments will be needed to address these possibilities.

The complexity of the Bid KO response to topical immunization required that we examine the behavior of cDCs in absence of the Bid KO host environment. Therefore, we harvested migratory cDCs from topically immunized skin explant cultures and examined their capacity to activate normal T cells in a WT host environment. We observed that upon adoptive transfer into WT recipient mice containing CFSE-labeled CD4 T cells, draining lymph nodes from mice inoculated with Ag-pulsed Bid KO cDCs were ∼2-fold larger as compared nodes of mice receiving Ag-pulsed WT cDCs. The cell proliferation that occurred within these enlarged lymph nodes was shown to be Ag specific, since TCR Vb5-negative CFSE-labeled CD4 T cells, cotransferred into in the same mouse showed no proliferation response. The increased rate and magnitude of the OT-2 T cell proliferation observed correlated with an increase in Th1 IFN-γ-producing cells found in both draining lymph nodes and spleen. Thus, Bid KO cDCs, in normal host recipient mice, promote increased effector T cell differentiation, not abortive proliferation.

We recognize that migratory cells from split ear specimens contain CD11c, I-A-positive LCs, and dDCs derived from both epidermal and dermal layers. Therefore, the effects seen in vivo by cDCs may be contributed by either or both DC populations. We have found that 70–90% of I-A+ or CD11c+ cells are also positive for the LC marker Langerin (CD207) (45). To enrich for activated LCs in our in vitro analysis we examined I-Ahigh and/or Langerin-expressing cells, both shown to correlate with LC phenotype (45 , 46). Therefore, migratory Bid KO-derived cDCs likely comprise mostly LCs (CD11c, CD205, and Langerin-positive cells). The Bid KO LC population demonstrates resistance to CD4 T cell-induced apoptosis in vitro and supports the hypothesis they may have a longer life span in the lymph node. The life span of such apoptosis-resistant LCs within the lymph node during the generation of Ag-specific responses will need to be examined to verify this interpretation.

Recent evidence points to the requirement of prolonged CD4-DC interactions and prolonged presence of Ag to sustain CD4 activation, proliferation (39, 47), and differentiation (40). Thus, an increased life span within the LC population may directly impact the duration of productive Ag presentation and sustained CD4 proliferation within the lymph node, allowing a larger number of CD4 T cells to modulate the effector arm of the immune response. Our finding that Bid KO cDCs promote increased numbers of IFN-γ-positive OT-2 cells supports this hypothesis. The impact of apoptosis-resistant LCs on development of memory and on supporting the generation of CD8 T cell effector and memory cells requires further investigation.

While Bid KO mice are reported to be phenotypically normal (23), detailed analysis of immune system function has not been reported previously. Interestingly, while Bid protein expression is restricted to neurons and to terminally differentiated cells fated to possesses short life spans (44), no increase in the rate of cancer development (including skin) has been documented, with the exception of a profound increase (53% in Bid KO vs 3% in WT) in the incidence of chronic myelomonocytic leukemia (48). This phenotype suggests that in young mice Bid-independent compensatory mechanisms exist to regulate tissue and cellular homeostasis, but with age, myeloid lineage cells have greater dependence on the type II apoptosis pathway for regulating their life span. Alternatively, Bid’s other function, controlling cell-cycle checkpoints during replicative stress, may be critical in aged myeloid precursors (49, 50). These observations are consistent with our findings that Bid has a critical role in the biology of myeloid lineage-derived LCs (51) and suggests that other myeloid DC subsets may also use the type II pathway.

The finding that Bid is used in a physiologically relevant manner to regulate LC function makes it an attractive molecular target for manipulating immune responses. There are a number of advantages in targeting Bid for such purposes: 1) selective cell targeting; DCs and cells with reduced proliferative potential (i.e., terminally differentiated) primarily express Bid. And so far, only LCs (as reported here) and hepatocytes (23) have been shown to use the type II pathway in normal hemopoietic tissues. Although, we expect that with more detailed investigation of the Bid KO strain, other cells may demonstrate some dependency on Bid for, perhaps, subtle impacts on normal mechanisms of homeostasis. 2) Reduced dysregulation; Bid KO cells can still respond to intrinsic and extrinsic type I signals of cell death, hence a normal phenotype is observed in Bid KO mice. 3) T cells unaffected; Bid-restricted expression spares T cells if systemic inhibitors were used. 4) Inhibition time frame is short; only short-term inhibition during Ag presentation is required to be effective in enhancing an immune response. Therefore, we propose that Bid-specific inhibitors may be used therapeutically as potent augmenters of “engineered” immune responses.

In appreciation of Dr. Stanley J. Korsmeyer, whose generosity and scientific excellence will continue to inspire an indebted generation of scientists. We thank Dr. Hui Xu for his guidance in performing contact sensitization experiments, Dr. Trenton R. Schoeb for pathology assessments, Kai Shi for assistance in performing tail vein injections in B6 WT and Bid KO mice, Christopher J. Thrash, James W. Evans, Holly Harris, Taylor C. Preston, and Eun Young Kho for excellent technical assistance.

The authors have no financial conflict of interest.

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 work has been supported by grants from the American Cancer Society, Dermatology Foundation, Charlotte Geyer Foundation, and National Institutes of Health Grants R01-AI50150, R01-CA86172, and P30-AR050948, and Department of Defense Grant W81XWH-0510296.

3

Abbreviations used in this paper: LC, Langerhans cell; 7-AAD, 7-aminoactinomycin D; AR-LC, apoptosis-resistant LC; BH3, Bcl-2 homology domain 3; Bid, BH3-interacting death domain; cDC, cutaneous DC; DC, dendritic cell; dDC, dermal DC; DNFB, 2–4-dinitrofluorobenzene; HEL, hen egg lysozyme; KO, knockout; TRANCE, TNF-related activation-induced cytokine; WT, wild type.

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