γδ T cells participate in the innate immune response to a variety of infectious microorganisms. They also link to the adaptive immune response through their induction of maturation of dendritic cells (DC) during the early phase of an immune response when the frequency of Ag-specific T cells is very low. We observe that in the presence of Borrelia burgdorferi, synovial Vδ1 T cells from Lyme arthritis synovial fluid potently induce maturation of DC, including production of IL-12, and increased surface expression of CD40 and CD86. The activated DC are then able to stimulate the Vδ1 T cells to up-regulate CD25. Both of these processes are initiated primarily by Fas stimulation rather than CD40 activation of DC via high expression of Fas ligand by the Vδ1 T cells. DC are resistant to Fas-induced death due to expression of high levels of the Fas inhibitor c-FLIP. This effect serves to divert Fas-mediated signals from the caspase cascade to the ERK MAPK and NF-κB pathways. The findings affirm the importance of the interaction of certain T cell populations with DC during the early phases of the innate immune response. They also underscore the view that as levels of c-FLIP increase, Fas signaling can be diverted from induction of apoptosis to pathways leading to cell effector function.

The innate immune response is designed to respond rapidly to infection as a first line of defense. To accomplish this, it uses an array of nonpolymorphic receptors, such as TLR and CD1 molecules, which are designed to recognize conserved motifs of microorganisms (1, 2, 3, 4, 5). Among the cell types that express these receptors are dendritic cells (DC),3 which are critical for linking innate immunity to the second phase adaptive immune response (6, 7). Immature DC are programmed for uptake of foreign Ags for processing, whereas mature DC are highly efficient at presentation of Ags and costimulatory molecules that are necessary for the activation of naive T cells (1, 8, 9, 10). Maturation of DC results in up-regulation of surface MHC, costimulatory molecules CD80 and CD86, and secretion of IL-12 in response to IFN-γ (11). In vitro, this mature stage may be as brief as 12 h, after which mature DC are no longer able to secrete IL-12 (11, 12). Members of the TNF-α superfamily can promote DC maturation, including CD40L and TNF-α, which are derived from certain activated T cells in addition to other cell types (11). However, these maturation cues are not likely to be provided by naive Th cells, as both the frequency of Ag-specific naive T cells and their effector function are too limited at the initiation of an immune response to promote rapid DC maturation.

A link between the innate and adaptive immune responses can be provided by T lymphocytes that react to the CD1 family or directly to microbial products. This includes NKT cells that respond to CD1d and human γδ T cells of the Vδ1 subset that recognize CD1c (3, 4, 13, 14, 15). In addition, the Vδ2 subset can recognize small nonpeptidic phosphorylated Ags derived from Mycobacteria (16, 17), certain bisphosphonates (18), alkylamines (17), and the association of membrane F1-ATPase with apolipoprotein A-I (19) in the absence of classical MHC molecules. The contribution of γδ T cells to defense from infection has been examined in mice in a number of model systems including Listeria (20), Leishmania (21), Mycobacterium (22), Plasmodium (23), and Salmonella (24). All of these studies have shown a protective role for γδ T cells. However, in other infectious models subsets of γδ T cells can have opposing effects. Thus, in murine Coxsackievirus-induced myocarditis, the Vγ1 subset was protective, whereas the Vγ4 subset promoted disease (25).

γδ T cells also accumulate at inflammatory sites in autoimmune disorders such as rheumatoid arthritis (26), celiac disease (27), and sarcoidosis (28). The reason for this effect remains largely unknown. Some evidence suggests that γδ T cells may have a regulatory role in certain autoimmune models to suppress inflammation. Collagen-induced arthritis in mice (29), adjuvant arthritis in rats (30), and a murine model of orchitis (31) are all worse after depletion of γδ T cells. Similarly, MRL/lpr mice lacking γδ T cells develop a more aggressive lupus-like illness (32).

Lyme arthritis represents an inflammatory synovitis caused by infection with Borrelia burgdorferi, but can also result in a persistent antibiotic-resistant arthritis after B. burgdorferi has been eradicated (33). This can manifest features of an autoimmune arthritis resembling rheumatoid arthritis (34). In addition, although γδ T cells are found at low levels in peripheral blood, the Vδ1 subset is increased to considerable levels in Lyme arthritis synovial fluid (35). We have previously observed that these synovial Vδ1 cells are intensely cytolytic due to expression of high and sustained levels of Fas ligand (FasL) (35). However, we now observe that monocyte-derived DC are resistant to lysis by Vδ1 clones. In exploring the mechanism for this Fas resistance we find that DC express high levels of the Fas inhibitor, c-FLIP, whereas the monocytes manifest very low c-FLIP expression.

c-FLIP is a homologue of caspase-8 but bears a mutation in the caspase domain that renders it enzymatically inactive (36, 37). As such, c-FLIP acts as a competitive inhibitor for recruitment of caspase-8 to Fas-associated death domain (FADD) protein following Fas ligation (38). c-FLIP has additional functions given its ability to associate with TNFR-associated factor (TRAF)1, TRAF2 and RIP1, which activate the NF-κB pathway, as well as with Raf-1, which activates the ERK MAPK pathway (39). Increased expression of c-FLIP can thus not only inhibit Fas-induced caspase-8 activation, it can also divert Fas signals toward the NF-κB and ERK pathways (39). We observe that synovial Vδ1 cells strongly activate DC in the presence of B. burgdorferi to produce IL-12, and to up-regulate surface CD40 and CD86. This effect is mediated largely by the high levels of FasL, as Vδ1 cells express very low to negligible levels of surface CD40L. Furthermore, FasL stimulation of DC induces rapid activation of the NF-κB and ERK pathways. Thus, the interaction of synovial Vδ1 cells with DC represents another example in a growing list of instances in which Fas activation can lead to outcomes of cell growth, differentiation, or effector function (40, 41, 42, 43, 44).

Lyme arthritis patients were followed at the Lyme Disease Clinic at the University of Medicine and Dentistry of New Jersey (Robert Wood Johnson Medical School, New Brunswick, NJ). All patients had histories, examinations, and serologies consistent with Lyme arthritis. Each had Abs to B. burgdorferi in both synovial fluid and serum detected by ELISA and confirmed by immunoblot.

Lymphocytes were purified from synovial fluid by Ficoll-Hypaque centrifugation (Sigma-Aldrich), and cultured in AIM-V medium (Invitrogen Life Technologies) containing 5% FBS (HyClone) and 50 U/ml human rIL-2 (Cetus; complete medium). Cells were stimulated with 10 μg/ml sonicate of B. burgdorferi strain N40, grown in BSK II medium (Barbour-Stoenner-Kelly II medium; Sigma-Aldrich) as previously described (35, 45). From these bulk cultures, responding cells were cloned at 0.3 cells/well in complete medium in the presence of irradiated PBL (3 × 105/well) and 10 μg/ml B. burgdorferi sonicate. After 14–21 days, cells from positive wells were phenotyped and those containing γδ+ T cells were expanded by restimulation with either B. burgdorferi or PHA (1 μg/ml; Murex Biotec), at ∼14-day intervals. All synovial γδ clones were Vδ1 by Ab screening and DNA sequencing and proliferated in response to B. burgdorferi stimulation (46).

Monocyte-derived immature DC were derived from bead-purified CD14+ monocytes (Miltenyi Biotec) obtained from peripheral blood of healthy volunteers and cultured with 800 U/ml GM-CSF (BioSource International), and 200 U/ml IL-4 (BioSource International).

Abs were to the determinants CD4 (S3.5; Caltag Laboratories), γδ TCR (5A6.E9; Caltag Laboratories), human Fas (DX2; BD Pharmingen), CD25 (CD25-3G10; Caltag Laboratories), CD80 (MEM-233; Caltag Laboratories), CD86 (BU63; Caltag Laboratories), CD40 (14G7; Caltag Laboratories), HLA-DR (TDR31.1; Ancell), and human FasL (monoclonal ALF2.1a; Ancell, or monoclonal NOK-1; BD Pharmingen). Surface FasL was analyzed using the catalyzed reporter deposition system of enzymatic amplification staining (EAS kit; Flow-Amp Systems) (47). Cells were stained as previously described (44). Samples were analyzed on either a Coulter Elite (Coulter), or BD LSR II flow cytometer (BD Biosciences). At least 2 × 104 events were accumulated for analysis.

Cultures of DC with Vδ1 clone cells were done at a ratio of 1:1 (each 1 × 106/ml) in complete medium. Supernatants were removed after 24 h for cytokine analysis by ELISA, and cells were stained for expression of TCR γδ, CD25, CD80, CD86, CD40, and HLA-DR. Inhibition studies were done using blocking Abs to FasL (5G51; Alexis), CD40L (24-31; a gift of Dr. R. Noelle, Dartmouth Medical School, Hanover, NH), IFN-γ (MAB25718; R&D Systems), IL-12 (clone 8.6; Endogen), and TNF-α (MAB1; BD Pharmingen).

Chemical fixation of DC was performed by incubating the cells in the presence of B. burgdorferi at 10 μg/ml at 37°C in 7% CO2 overnight, then washed in 5% FBS/RPMI 1640 (Mediatech), and fixed by the addition of ice-cold EDCI (1-ethyl-3(3′-dimethyl-aminopropyl)-carbodiimide (Pierce) at 75 mM in PBS for 60 min on ice. Following fixation the cells were extensively washed with 5% FBS/RPMI 1640.

Quantitation of specific cytokines (IL-12p70, IFN-γ, and IL-1β) in cell culture supernatants was performed using sandwich ELISA kits (Biosource International) according to the manufacturer’s protocol. Briefly, specific supernatants, along with appropriate standards and controls, were added to microtiter wells previously coated with a mAb specific for each cytokine. A second cytokine-specific biotinylated mAb was added at the same time. The mixture was then allowed to incubate for a specified time for each cytokine ELISA according to the manufacturer’s protocol. Excess biotinylated Ab was removed by extensive washing, followed by the addition of a solution containing streptavidin-HRP. Excess unbound enzyme was removed by extensive washing, followed by the addition of a chromogenic enzyme substrate solution. Plates were read on a Bio-Tek plate reader model ELX 800 (Bio-Tek Instruments) at 450 nm.

Target DC were labeled by incubation with 51Cr for 1 h, washed three times, and then mixed in 200 μl of complete medium at a 1:1 ratio with HEK 293 cells that were either mock-transfected (293) or transfected with human FasL (293hFasL) bearing a mutation at the membrane proximal proteolytic cleavage site, which was a gift of Dr. P. Schneider (University of Lausanne, Lausanne, Switzerland). Target cells were also treated with soluble FLAG-tagged FasL (Apotech) that was cross-linked with anti-FLAG Ab M2 (Sigma-Aldrich), or with anti-FLAG alone. After 4 h at 37°C, 100 μl supernatant was removed and counted for gamma emission. Spontaneous release was determined from labeled targets in the absence of effector cells. Maximal release was determined by lysing targets with 1.0 N HCl. The percentage of maximal 51Cr release was calculated as (experimental cpm − spontaneous cpm)/(maximal cpm − spontaneous cpm).

Primers for human FasL were designed to amplify an 84-bp fragment. The primers were: forward 5′-TGGCCCATTTAACAGGCAA-3′ and reverse 5′-CCAGAAAGCAGGACAATTCCA-3′. The amplified fragment contained the sequence bound by the fluorochrome-labeled primer: 5′-6-FAM TCCAACTCAAGGTCCATGCCTCTGG TAMRA-3′ (Biosearch Technologies). Control amplification was assessed using endogenous control 18S ribosomal RNA (PE Biosystems) labeled with a VIC reporter dye. RNA was extracted from cells using Ultraspec (Biotecx Laboratories), or RNeasy kit (Qiagen) treated with RNase-free DNase (Ambion), and cDNA was made using Superscript reverse transcriptase (Invitrogen Life Technologies). Quantitative PCR was performed using a 7700 Sequence Detection System (Applied Biosystems) at the Vermont Cancer Center DNA Analysis Facility (Burlington, VT). Fluorescence signal was expressed as normalized reporter (Rn) signal, which represents the reporter signal (FAM or VIC) divided by the fluorescence signal of a passive reference dye (Rox). Validation control experiments were performed to measure efficiency of the target (FasL) and reference (18S) gene amplifications over a range of 3 logs of sample dilution. All assays were run in duplicate and corrected to a reference-pooled sample (calibrator), which was included in each separate run. Threshold cycle (CT) values were expressed as 2−ΔΔCT (CT FasL−CT 18 S).

Primers for human CD40L were designed to amplify a 74-bp fragment. The primers were: forward 5′-ACCCACAGTTCCGCCAAAC-3′ and reverse 5′-GCACCTGGTTGCAATTCAAATA-3′. The amplified fragment contained the sequence bound by the fluorochrome-labeled probe: 5′-6-FAM TGCGGGCAACAATCCATTCACTTG BHQ-1–3′ (Biosearch Technologies). Control amplification was assessed using endogenous human 18S ribosomal cDNA as well as cDNA from CD40L-transfected Chinese hamster ovary cells. cDNA from human FasL (293hFasL) cells was included as a negative amplification control for CD40L. Assays were run in duplicate, and values were expressed as 2−ΔΔCT (CT CD40L−CT 18 S).

Cells were washed twice in ice-cold PBS and solubilized in lysis buffer (1% Nonidet P-40, 50 mM Tris-HCl, pH 7.6, 150 mM NaCl, 2 mM DTT, protease inhibitor mixture) (Complete; Boehringer Mannheim). Postnuclear lysates were collected after centrifugation (15,000 × g) and protein samples (40 μg of each) were separated by 10% SDS-PAGE. Proteins were transferred to polyvinylidene difluoride membranes (ImmunoBlot; Bio-Rad) and blots were blocked and probed with mAbs to phospho-IκBα, phospho-ERK, and ERK2 (5A5 and E10; Cell Signaling Technology, and C-14; Santa Cruz Biotechnology) as well as a polyclonal Ab to IκBα (Cell Signaling Technology) or monoclonal anti-c-FLIP (Dave-2; Apotech) in 4% nonfat milk in PBS/Tween 20 (0.1%). Immunoreactive proteins were visualized using HRP-labeled conjugates (The Jackson Laboratory) and ECL blotting substrate (Amersham).

Paired and unpaired t tests were used to assess the significance of differences in production of IFN-γ and IL-12 by, respectively, Vδ1 clones or DC, as well as CD25 expression for Vδ1 clones over the course of several experiments as noted.

Immature myeloid DC are highly efficient at activation of Lyme arthritis synovial γδ T cells of the Vδ1 subset in the presence of B. burgdorferi and exogenous IL-2 (46). Although the determinant recognized by these Vδ1 cells is not known, a strong induction of CD25 expression on the Vδ1 cells was apparent after 24 h of stimulation by DC that were pulsed for 16 h with a sonicate of B. burgdorferi (Fig. 1,A). Fixation of DC with EDCI following an overnight incubation with B. burgdorferi reduced the ability of DC to activate Vδ1 cells to either up-regulate CD25 (Fig. 1,A) or to produce IFN-γ (data not shown). This suggested that full activation of Vδ1 cells required metabolically active DC. In addition to CD25 expression, the production of IFN-γ by the Vδ1 cells, and production of bioactive IL-12 (p70) by DC were both considerably augmented when the Vδ1 cells and DC were cocultured in the presence of B. burgdorferi (Fig. 1, B and C). Culture supernatants from B. burgdorferi-pulsed DC were insufficient to activate Vδ1 cells, and vice versa, demonstrating that cell contact was necessary (data not shown).

FIGURE 1.

Mutual activation of Vδ1 cells and DC in the presence of B. burgdorferi. Synovial Vδ1 clone Bb03 and immature myeloid DC were either cultured separately or cocultured at a 1:1 ratio (each 1 × 106 cells/ml) overnight in the absence or presence of 10 μg/ml sonicate of B. burgdorferi as indicated (+). Cells were then stained for expression (A) of CD25 on the Vδ1-gated subset, or supernatants examined for production of IFN-γ (B) by Vδ1 clones or IL-12 (C) by DC. In some cultures in A, DC were first cultured overnight with B. burgdorferi sonicate alone, then washed and left either unfixed (un) or fixed (fx) with EDCI. Shown are the findings from one of five similar experiments. Statistically significant increase (∗) in CD25 (p = 0.0006), IFN-γ (p = 0.0132), and IL-12 (p = 0.0025) for cultures of DC plus Bb03 in the presence of B. burgdorferi compared with its absence.

FIGURE 1.

Mutual activation of Vδ1 cells and DC in the presence of B. burgdorferi. Synovial Vδ1 clone Bb03 and immature myeloid DC were either cultured separately or cocultured at a 1:1 ratio (each 1 × 106 cells/ml) overnight in the absence or presence of 10 μg/ml sonicate of B. burgdorferi as indicated (+). Cells were then stained for expression (A) of CD25 on the Vδ1-gated subset, or supernatants examined for production of IFN-γ (B) by Vδ1 clones or IL-12 (C) by DC. In some cultures in A, DC were first cultured overnight with B. burgdorferi sonicate alone, then washed and left either unfixed (un) or fixed (fx) with EDCI. Shown are the findings from one of five similar experiments. Statistically significant increase (∗) in CD25 (p = 0.0006), IFN-γ (p = 0.0132), and IL-12 (p = 0.0025) for cultures of DC plus Bb03 in the presence of B. burgdorferi compared with its absence.

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CD40L is a principal molecule by which CD4+ αβ T cells activate DC (48). Whereas B. burgdorferi-reactive CD4+ αβ T cell clones expressed high levels of surface CD40L upon activation, the synovial-derived Vδ1 cell clones expressed very low to negligible levels of CD40L. As shown in Fig. 2, A and B, following B. burgdorferi stimulation, the CD4+ αβ T cell clones expressed high levels of surface CD40L for at least 3 days before declining to baseline levels. By contrast, following B. burgdorferi stimulation, Vδ1 cells expressed nearly undetectable levels of surface CD40L over the same period of time (Fig. 2, A and B). This finding was also true of fresh γδ T cells in Lyme synovial fluid (Fig. 2, A and B). These findings were similar at the message level over an 18-day period using quantitative PCR (Fig. 2 C).

FIGURE 2.

Vδ1 cells express negligible levels of CD40L. B. burgdorferi-specific TCR αβ CD4+ T cell clones 114B and 2-7, or Vδ1 clones Bb01 and Bb03 were stimulated with syngeneic DC in the presence of B. burgdorferi and IL-2 (40 U/ml). In parallel, fresh Lyme synovial fluid (SF) lymphocytes were also stimulated with B. burgdorferi and IL-2 in the absence of additional DC. A, FACS profiles of surface CD40L expression on day 3 after stimulation. B, Kinetics of CD40L expression from a second experiment over a 10-day period following activation. C, Kinetics of CD40L mRNA expression over 10 days by quantitative PCR. A positive control was provided by a stable Chinese hamster ovary (CHO) transfectant cell line expressing human CD40L. Negative controls were provided by untransfected Chinese hamster ovary cells and 293 cells stably expressing 293hFasL. The findings were consistent in a total of three experiments.

FIGURE 2.

Vδ1 cells express negligible levels of CD40L. B. burgdorferi-specific TCR αβ CD4+ T cell clones 114B and 2-7, or Vδ1 clones Bb01 and Bb03 were stimulated with syngeneic DC in the presence of B. burgdorferi and IL-2 (40 U/ml). In parallel, fresh Lyme synovial fluid (SF) lymphocytes were also stimulated with B. burgdorferi and IL-2 in the absence of additional DC. A, FACS profiles of surface CD40L expression on day 3 after stimulation. B, Kinetics of CD40L expression from a second experiment over a 10-day period following activation. C, Kinetics of CD40L mRNA expression over 10 days by quantitative PCR. A positive control was provided by a stable Chinese hamster ovary (CHO) transfectant cell line expressing human CD40L. Negative controls were provided by untransfected Chinese hamster ovary cells and 293 cells stably expressing 293hFasL. The findings were consistent in a total of three experiments.

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The opposite pattern of expression was observed for FasL (Fig. 3). Although both αβ and γδ B. burgdorferi-reactive T cells quickly increased surface FasL following stimulation, the αβ T cells rapidly down-regulated surface FasL, whereas the Vδ1 cells maintained FasL expression for at least 16 days (Fig. 3,B), and in further studies for as long as 30 days (data not shown). This was also apparent at the mRNA level (Fig. 3 C). The findings are also consistent with our previous observations of intense cytolytic activity by synovial Vδ1 cells (49).

FIGURE 3.

Vδ1 cells express high and sustained FasL. The same clones and synovial fluid lymphocytes as in Fig. 2 were analyzed in a similar manner for expression of FasL following stimulation with B. burgdorferi. A, FACS profiles of one of five similar experiments for surface FasL on day 10 after activation. B, Temporal expression of surface FasL over 15 days. C, Representative quantitative PCR for FasL on day 13 after activation.

FIGURE 3.

Vδ1 cells express high and sustained FasL. The same clones and synovial fluid lymphocytes as in Fig. 2 were analyzed in a similar manner for expression of FasL following stimulation with B. burgdorferi. A, FACS profiles of one of five similar experiments for surface FasL on day 10 after activation. B, Temporal expression of surface FasL over 15 days. C, Representative quantitative PCR for FasL on day 13 after activation.

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Although Vδ1 cells are highly cytolytic to several cell types, they did not lyse immature myeloid DC (data not shown). As Fas expression was very high on DC (Fig. 4,A), this suggested an internal mechanism of resistance. c-FLIP is the natural inhibitor of Fas-induced cell death as it competes with caspase-8 for recruitment to FADD in the death-inducing signal complex (38). Levels of c-FLIP were thus examined by immunoblot from lysates of the precursor CD14+ monocytes and the resulting immature DC following 7 days culture with GM-CSF and IL-4. As shown in Fig. 4 B, the levels of c-FLIP were nearly undetectable in peripheral blood CD14+ monocytes but were greatly induced in the resulting DC.

FIGURE 4.

Immature myeloid DC express high levels of c-FLIP and are resistant to Fas-induced cell death. A, Expression of surface Fas by immature DC. B, Immunoblot of c-FLIP and control actin for cell lysates from Jurkat T cells stably transfected with c-FLIP, fresh CD14+ monocytes from peripheral blood, and immature myeloid DC derived from the monocytes after 7 days culture in GM-CSF and IL-4. C, Cytotoxicity measured by 51Cr release from labeled DC or Jurkat T cells not overexpressing c-FLIP, in the presence for 4 h of 293 mock-transfected cells, 293hFasL-expressing cells, anti-FLAG Ab alone, and soluble FLAG-tagged FasL (sFasL) oligomerized with anti-FLAG.

FIGURE 4.

Immature myeloid DC express high levels of c-FLIP and are resistant to Fas-induced cell death. A, Expression of surface Fas by immature DC. B, Immunoblot of c-FLIP and control actin for cell lysates from Jurkat T cells stably transfected with c-FLIP, fresh CD14+ monocytes from peripheral blood, and immature myeloid DC derived from the monocytes after 7 days culture in GM-CSF and IL-4. C, Cytotoxicity measured by 51Cr release from labeled DC or Jurkat T cells not overexpressing c-FLIP, in the presence for 4 h of 293 mock-transfected cells, 293hFasL-expressing cells, anti-FLAG Ab alone, and soluble FLAG-tagged FasL (sFasL) oligomerized with anti-FLAG.

Close modal

Consistent with the high levels of c-FLIP expression, DC were also extremely resistant to Fas-mediated cell death. As a positive control, Jurkat T cells that expressed high levels of surface Fas were very sensitive to cell death using either soluble FasL or 293 cells stably transfected with 293hFasL (Fig. 4,C). In contrast, Jurkat cells stably transfected with c-FLIP were highly resistant to Fas-induced death (data not shown). In a similar manner, day 7 immature DC resembled the c-FLIP-transfected Jurkat cells in that they were also almost completely resistant to Fas-mediated cell death by either means of FasL exposure (Fig. 4 C).

We have previously established in cell lines that c-FLIP can promote the activation of the MAPK, ERK, through its association with Raf-1 (39). Similarly c-FLIP can also augment NF-κB activity through association with TRAF1, TRAF2, and RIP1 (39). We therefore considered that ligation of Fas on DC might result in increased effector function, rather than cell death. Consistent with this view, stimulation of DC by soluble cross-linked FasL for 30 min resulted in rapid phosphorylation of ERK as well as the NF-κB inhibitor, IκBα (Fig. 5), which primes IκBα for ubiquitination and degradation.

FIGURE 5.

Rapid activation of ERK MAPK and NF-κB pathways in DC following Fas ligation. Immature DC were cultured for 30 min in the presence of anti-FLAG Ab alone or soluble FasL (sFasL; 200 ng/ml) plus anti-FLAG Ab (2 μg/ml). Cells were then lysed and lysates assessed by immunoblot for phospho-ERK vs total ERK (A), and phospho-IκBα vs total IκBα (B).

FIGURE 5.

Rapid activation of ERK MAPK and NF-κB pathways in DC following Fas ligation. Immature DC were cultured for 30 min in the presence of anti-FLAG Ab alone or soluble FasL (sFasL; 200 ng/ml) plus anti-FLAG Ab (2 μg/ml). Cells were then lysed and lysates assessed by immunoblot for phospho-ERK vs total ERK (A), and phospho-IκBα vs total IκBα (B).

Close modal

To more effectively define whether Fas ligation on DC resulted in alterations in function, Fas was ligated on DC by three different methods: soluble cross-linked FasL, agonistic IgM anti-Fas Ab, and 293hFasL cells. Immature DC spontaneously produced very little IL-12, but following stimulation with soluble cross-linked FasL or agonistic anti-Fas Ab there was a significant increase in IL-12 production (Fig. 6,A). A particularly dramatic induction of IL-12 by DC was observed using the 293hFasL cell line, whereas the mock-transfected 293 cells did not augment IL-12 production (Fig. 6 B). The 293hFasL cells were not the source of IL-12 as none was detected in culture supernatants of 293hFasL cells or in intracellular staining of these same cells (data not shown). The findings were consistent in three separate experiments.

FIGURE 6.

Fas signals IL-12 production by DC. A, Immature DC were activated by anti-FLAG Ab alone, FLAG-tagged soluble FasL (sFasL) cross-linked by anti-FLAG, or agonistic IgM anti-Fas (CH-11), and supernatants examined after 24 h for IL-12 production by ELISA. B, Immature DC were stimulated with 293 cells alone, or 293 cells stably expressing FasL (293hFasL). Supernatants were examined after 24 h for IL-12 production. Shown are mean ± SD of IL-12 levels. Statistically significant increase (∗) of IL-12 production in the presence of the various forms of Fas stimulation (DC + sFasL p = 0.0032; DC + anti-Fas CH11 p = 0.0032; DC + 293hFasL p = 0.0086).

FIGURE 6.

Fas signals IL-12 production by DC. A, Immature DC were activated by anti-FLAG Ab alone, FLAG-tagged soluble FasL (sFasL) cross-linked by anti-FLAG, or agonistic IgM anti-Fas (CH-11), and supernatants examined after 24 h for IL-12 production by ELISA. B, Immature DC were stimulated with 293 cells alone, or 293 cells stably expressing FasL (293hFasL). Supernatants were examined after 24 h for IL-12 production. Shown are mean ± SD of IL-12 levels. Statistically significant increase (∗) of IL-12 production in the presence of the various forms of Fas stimulation (DC + sFasL p = 0.0032; DC + anti-Fas CH11 p = 0.0032; DC + 293hFasL p = 0.0086).

Close modal

Alterations in the surface phenotype of DC was also apparent after Fas stimulation. Following a 24 h stimulation by soluble FasL, the surface expression increased for both CD40 and CD86 (Fig. 7). A similar change of phenotype was also apparent using anti-Fas Ab (data not shown).

FIGURE 7.

FasL promotes DC surface expression of CD40 and CD86. Immature DC were cultured with anti-FLAG Ab alone, or with soluble FasL (sFasL) plus anti-FLAG for 16 h and then analyzed for surface expression of the indicated molecules. Results are expressed as mean fluorescence intensity (MFI) and were consistent in three separate experiments.

FIGURE 7.

FasL promotes DC surface expression of CD40 and CD86. Immature DC were cultured with anti-FLAG Ab alone, or with soluble FasL (sFasL) plus anti-FLAG for 16 h and then analyzed for surface expression of the indicated molecules. Results are expressed as mean fluorescence intensity (MFI) and were consistent in three separate experiments.

Close modal

Given the high and sustained levels of surface FasL expressed by the synovial Vδ1 clones, combined with their very low levels of CD40L expression, it was of interest to assess to what extent, if any, these γδ T cells would stimulate DC via FasL, and whether this would render the DC competent to activate the Vδ1 clones. For these studies we chose two representative Vδ1 clones, one that expresses very high levels of FasL (Bb01) and a second that expresses moderate and sustained levels of FasL (Bb03) (see Fig. 3,C). Both clones express negligible surface CD40L (Fig. 2 C). Each clone was mixed with an equal number of DC in the presence of B. burgdorferi sonicate for 18 h and then supernatants were assayed for IL-12 and IFN-γ secretion, followed by surface staining of CD86 on DC and CD25 up-regulation on the Vδ1 clones.

As shown in Fig. 8,A, DC production of IL-12 was greatly augmented by the FasLhigh Vδ1 clone, Bb01, and to a lesser extent by the FasLmoderate Vδ1 clone, Bb03. The ability to induce IL-12 production by DC was extensively inhibited by blocking anti-FasL Ab in a dose-dependent manner, though not by isotype control IgG. The degree of inhibition by anti-FasL was also proportional to the level of FasL expressed by the Vδ1 clone. Thus, stimulation of DC IL-12 by Bb01 was blocked nearly 75% with anti-FasL at 20 μg/ml, whereas the same concentration of anti-FasL blocked IL-12 induced by Bb03 × 50%. The effectiveness of the blocking anti-FasL Ab was demonstrated by its ability to completely inhibit cytolysis of Jurkat T cells by the 293hFasL cells (data not shown). These findings were consistent in five experiments. In addition, IFN-γ is a known stimulatory cytokine for IL-12 production by DC (50). Given that the synovial Vδ1 clones produce IFN-γ, it was not surprising that inhibition of IFN-γ also partly blocked IL-12 production by the DC (Fig. 8,A). In a similar manner, CD86 expression by DC, which was induced by the Vδ1 clones, was blocked by anti-FasL though not by anti-IFN-γ or control IgG (Fig. 8,B). Consistent with the low level CD40L expression by the Vδ1 clones, blocking anti-CD40L had no effect on either CD86 expression (Fig. 8 C) or IL-12 production (data not shown) in this system. These findings were consistent in three separate studies.

FIGURE 8.

Vδ1 clones stimulate DC via FasL. FasLhigh Vδ1 clone Bb01 and FasLmoderate Vδ1 clone Bb03 were cultured for 16 h in the presence of immature DC, B. burgdorferi, and either murine IgG (mIgG), blocking anti-FasL at the indicated concentrations, or blocking IFN-γ, CD40L, or TNF-α (each 20 μg/ml) and supernatants assessed for IL-12 production (A) by Vδ1 clones using ELISA, or up-regulation of surface CD86 expression by (B and C) DC as assessed by FACS. The findings are representative of three experiments. Shown are the mean ± SD for DC for IL-12 production. Statistically significant decrease (∗) in IL-12 production between control murine IgG vs anti-FasL at 20 μg/ml (Bb01 p = 0.0077 and Bb03 p = 0.067), 10 μg/ml (Bb01 p = 0.0110 and Bb03 p = 0.0129), and anti-IFN-γ at 20 μg/ml (Bb01 p = 0.0098 and Bb03 p = 0.0051).

FIGURE 8.

Vδ1 clones stimulate DC via FasL. FasLhigh Vδ1 clone Bb01 and FasLmoderate Vδ1 clone Bb03 were cultured for 16 h in the presence of immature DC, B. burgdorferi, and either murine IgG (mIgG), blocking anti-FasL at the indicated concentrations, or blocking IFN-γ, CD40L, or TNF-α (each 20 μg/ml) and supernatants assessed for IL-12 production (A) by Vδ1 clones using ELISA, or up-regulation of surface CD86 expression by (B and C) DC as assessed by FACS. The findings are representative of three experiments. Shown are the mean ± SD for DC for IL-12 production. Statistically significant decrease (∗) in IL-12 production between control murine IgG vs anti-FasL at 20 μg/ml (Bb01 p = 0.0077 and Bb03 p = 0.067), 10 μg/ml (Bb01 p = 0.0110 and Bb03 p = 0.0129), and anti-IFN-γ at 20 μg/ml (Bb01 p = 0.0098 and Bb03 p = 0.0051).

Close modal

As noted earlier, DC cultured in the presence of B. burgdorferi can induce the Vδ1 clones to up-regulate surface CD25 and to secrete IFN-γ. Expression of CD25 by Vδ1 cells was greatly reduced in the presence of anti-FasL compared with control IgG (Fig. 9,A). In a manner similar to the stimulation of DC, the inhibition of CD25 expression by anti-FasL was proportional to the expression of FasL by the Vδ1 clone. Thus, induction of surface CD25 on FasLhigh Bb01 was more effectively blocked by anti-FasL than was CD25 expression on FasLmoderate Bb03. In contrast to anti-FasL, neither anti-IFN-γ (Fig. 9,A) nor anti-IL-12 (Fig. 9,B) had any effect on CD25 expression by the Vδ1 clones in three separate experiments, whereas anti-IL-12 very efficiently inhibited production of IFN-γ (Fig. 9 C). Thus, despite the absence of significant CD40L expression by synovial Vδ1 clones, there is nonetheless a strong mutual stimulation between DC and Vδ1 cells that is driven largely by FasL. This extends the number of cell types that, in the presence of high levels of c-FLIP, can divert Fas-mediated signals from a death pathway toward activation and effector function.

FIGURE 9.

DC activate Vδ1 cells via Fas signaling and IL-12 production. FasLhigh Vδ1 clone Bb01 and FasLmoderate Vδ1 clone Bb03 were activated with DC pulsed with B. burgdorferi in the presence of control murine IgG (mIgG), or blocking Abs to human FasL, IFN-γ, or IL-12. After 16 h culture, the Vδ1 clones were examined for surface CD25 expression (A and B) and IFN-γ production (C) in supernatants using ELISA. Statistically significant decreases (∗) in CD25 expression by Vδ1 clone plus DC plus B. burgdorferi in the presence of blocking anti-FasL at 20 μg/ml (Bb01 p = 0.0008 and Bb03 p = 0.0010), 10 μg/ml (Bb01 p = 0.0010 and Bb03 p = 0.0019), 5 μg/ml (Bb01 p = 0.0061 and Bb03 p = 0.0041). Differences with blocking anti-IFN-γ or anti-IL-12 were not significant. Statistically significant inhibition (∗) of anti-IFN-γ production by anti-IL-12 (p = 0.0247).

FIGURE 9.

DC activate Vδ1 cells via Fas signaling and IL-12 production. FasLhigh Vδ1 clone Bb01 and FasLmoderate Vδ1 clone Bb03 were activated with DC pulsed with B. burgdorferi in the presence of control murine IgG (mIgG), or blocking Abs to human FasL, IFN-γ, or IL-12. After 16 h culture, the Vδ1 clones were examined for surface CD25 expression (A and B) and IFN-γ production (C) in supernatants using ELISA. Statistically significant decreases (∗) in CD25 expression by Vδ1 clone plus DC plus B. burgdorferi in the presence of blocking anti-FasL at 20 μg/ml (Bb01 p = 0.0008 and Bb03 p = 0.0010), 10 μg/ml (Bb01 p = 0.0010 and Bb03 p = 0.0019), 5 μg/ml (Bb01 p = 0.0061 and Bb03 p = 0.0041). Differences with blocking anti-IFN-γ or anti-IL-12 were not significant. Statistically significant inhibition (∗) of anti-IFN-γ production by anti-IL-12 (p = 0.0247).

Close modal

The current findings extend the potential functions of the subset of γδ T cells expressing the Vδ1 TCR through its high and sustained expression of FasL. Immature myeloid DC are resistant to Fas-mediated cell death due to their up-regulated expression of c-FLIP during differentiation from peripheral blood monocytes in the presence of GM-CSF and IL-4. c-FLIP prevents Fas signaling from propagating down the caspase cascade, and diverts it toward activation of ERK and NF-κB. This results in the secretion of IL-12 by DC and up-regulation of surface markers such as CD86 that are important to activation of naive T cells in the adaptive immune response. This renders DC capable of promoting CD25 up-regulation on the Vδ1 cells in the presence of B. burgdorferi. A mutual costimulation thus results between Vδ1 cells and DC, which involves to a large extent interactions of Fas/FasL rather than CD40/CD40L.

A growing number of cell types have been shown to respond to Fas signaling with increased proliferation or differentiation rather than cell death. This includes fibroblast growth (42), cardiac myocyte hypertrophy (51), neurite outgrowth (41), hepatocyte regeneration (43), and growth of certain tumors (52). However, the signaling pathway leading to this unexpected effect was not defined. We have previously observed that increased expression of the death receptor inhibitor c-FLIP is able to block activation of the caspase cascade and hence cell death by Fas, and simultaneously divert signals toward the NF-κB and ERK MAPK pathways (39). c-FLIP is homologous to caspase-8 in containing two N-terminal death effector domains, but lacks the critical C-terminal cysteine at the caspase enzymatic pocket, rendering c-FLIP enzymatically inert. As such, c-FLIP functions as a competitive inhibitor of caspase-8 recruitment to FADD following Fas ligation (38). The ability of c-FLIP to activate the NF-κB and ERK pathways derives from the ability of c-FLIP to bind to RIP1, TRAF1, TRAF2, and Raf1 (39). The model of Fas signaling through c-FLIP thus may not require caspase activation. This idea is consistent with our finding that blocking caspase activity did not alter Fas-induced production of IL-12 or IL-12 up-regulation of CD86 (data not shown). Based on these qualities c-FLIP is an ideal candidate for switching Fas signals from the caspase cascade toward those associated with growth or differentiation. This would also be consistent with the finding that freshly isolated peripheral blood CD14+ monocytes express very low levels of c-FLIP and are very sensitive to Fas-induced death. This is consistent with recent reports that T cells can kill activated macrophages via Fas-FasL interactions (53). By contrast, the immature DC derived from monocytes have high levels of c-FLIP and are extremely resistant to Fas-induced death.

Our findings are consistent with those of Rescigno et al. (40) who also observed that FasL stimulation of human or mouse DC increased production of IL-12 and up-regulated MHC class II, CD40, and CD86. However, those studies did not suggest a signaling pathway nor did they show any interaction of DC with γδ T cells. Because both CD4+ and CD8+ T cells express FasL transiently upon activation, this mechanism of DC activation may be more widely applied than to just γδ T cells. Lack of Fas expression by DC may contribute in part to the susceptibility of Fas-deficient lpr mice to certain infections such as with Toxoplasma gondii (54, 55).

Cell-to-cell contact between synovial Vδ1 cells and DC was necessary to produce maximal activation of each cell type. However, soluble factors subsequently produced by both cells were instrumental in achieving full activation. Thus, IFN-γ production by Vδ1 cells contributed considerably to IL-12 production but not CD86 up-regulation by DC. Conversely, IL-12 production by DC promoted secretion of IFN-γ but not CD25 expression by the Vδ1 cells. This cytokine interplay may be partly responsible for the previously reported ability of γδ T cells to promote a Th1 cytokine environment in a Fas-dependent manner during the immune response to Coxsackievirus (56). Furthermore, in contrast to activated αβ T cells, CD40L had little influence in the interaction between Vδ1 cells and DC as the Vδ1 cells expressed negligible levels of CD40L and blocking this had no effect of the ability of the Vδ1 cells to activate immature DC.

The transgenic expression of FasL by different tissues was initially considered as a potentially effective method to deplete the immune system of T cells reactive with that tissue. In many of these cases the opposite result was observed, with augmented tissue injury. Tumor cells expressing FasL were rejected even faster than the parent clone, due in part to activation of macrophages via FasL to secrete chemokines that recruited polymorphonuclear neutrophils (57). A related study of NOD mice observed that β islet cells transgenically expressing FasL were more sensitive to diabetogenic T cells in a FasL-dependent manner (58). These FasL+ β islet cells also resulted in more rapid rejection of pancreases transplanted under the kidney capsule of allogeneic mice. Thus, FasL expression, either normally by activated T cells, or ectopically in target tissues, can lead to enhanced activation of responding T cells, possibly through stimulation of DC and subsequent recruitment of other inflammatory cells.

γδ T cells likely make important contributions in the response to various infections as well as in autoimmune conditions. These include protective roles in infectious models of tuberculosis (22), listeriosis (20), malaria (23), leishmaniasis (21), and salmonellosis (24). In humans the Vγ2Vδ2 subset of γδ T cells increases substantially in response to infection with Mycobacterium tuberculosis, Brucella melitensis, Listeria monocytogenes, and Ehrlichia chaffeensis (59), and has been shown to respond to small nonpeptide Ags that include alkylphosphate (16, 60), alkylamine (17), and bisphosphonate (17). Whereas Vγ2Vδ2 T cells represent the major γδ cells in peripheral blood (61), those expressing the Vδ1 TCR are distributed largely in tissues, including intestine, spleen, and inflamed joints (62). Vδ1 cells are also increased during infection with HIV (63, 64) and Plasmodium (65). In the latter case this may also represent depletion or anergy of Vδ2 cells that were activated by the parasite’s alternative pathway of isoprenoid synthesis. It is not currently known how synovial Vδ1 cells maintain high levels of FasL. This T cell subset nonetheless represents an important link between the innate and adaptive immune responses through Fas stimulation of DC.

We thank Colette Charland for technical assistance with flow cytometry and Timothy Hunter in the Vermont Cancer Center for assistance with real-time PCR.

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 was supported by Grants AR43520 and AI 45666 (to R.C.B.) and P30CA22435 (to Vermont Cancer Center) from the National Institutes of Health.

3

Abbreviations used in this paper: DC, dendritic cell; FasL, Fas ligand; FADD, Fas-associated death domain protein; TRAF, TNFR-associated factor; 293hFasL, human FasL; CT, threshold cycle.

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