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The Journal of Immunology, 2007, 179, 5819-5828
Copyright © 2007 by The American Association of Immunologists, Inc.
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Autoimmunity Stimulated by Adoptively Transferred Dendritic Cells Is Initiated by Both {alpha}beta and {gamma}{delta} T Cells but Does Not Require MyD88 Signaling1

David A. Martin2,*, Kang Zhang*, Justin Kenkel*, Grant Hughes*, Edward Clark{dagger},{ddagger}, Anne Davidson§ and Keith B. Elkon3,*,{ddagger}

* Division of Rheumatology, {dagger} Department of Microbiology, and {ddagger} Department of Immunology, University of Washington School of Medicine, Seattle, WA 98195; and § Feinstein Institute for Medical Research, North Shore-Long Island Jewish Health System, Manhasset, NY 11030


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Vaccination of nonautoimmune prone mice with syngeneic dendritic cells (DC) readily induces anti-DNA autoantibodies but does not trigger systemic disease. We observed that anti-DNA autoantibody generation absolutely required {alpha}beta T cells and that {gamma}{delta} T cells also contributed to the response, but that regulatory T cells restrained autoantibody production. Although both NZB/W F1 mice and DC vaccinated C57/BL6 mice produced autoantibodies against dsDNA, vaccinated mice had higher levels of Abs against H1 histone and lower levels of antinucleosome Abs than NZB/W F1 mice. Despite a 100-fold increase in IL-12 and Th1 skewing to a foreign Ag, OVA, synergistic TLR activation of DC in vitro failed to augment anti-DNA Abs or promote class switching beyond that induced by LPS alone. TLR stimulation was not absolutely required for the initial loss of B cell tolerance because anti-DNA levels were similar when wild-type (WT) or MyD88-deficient DC were used for vaccination or WT and MyD88-deficient recipients were vaccinated with WT DC. In contrast, systemic administration of LPS, augmented anti-DNA Ab levels and promoted class switching, and this response was dependent on donor DC signaling via MyD88. LPS also augmented responses in the MyD88-deficient recipients, suggesting that LPS likely exerts its effects on both transferred DC and host B cells in vivo. These results indicate that both the {alpha}beta and {gamma}{delta} subsets are necessary for promoting autoantibody production by DC vaccination, and that although TLR/MyD88 signaling is not absolutely required for initiation, this pathway does promote augmentation, and Th1-mediated skewing, of anti-DNA autoantibodies.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Although spontaneous lupus models provide insight into mechanisms of disease, by the time that disease features develop, it is difficult to unravel which abnormalities are primary and which are secondary or tertiary phenomena. Furthermore, the activation of multiple cell types makes it difficult to determine the role of any particular cell type in initiation of disease. In contrast, the ability to induce lupus-like autoantibodies by manipulation of a single cell type, the dendritic cell (DC),4 allows for dissection of the cell subsets and mechanisms responsible for loss of tolerance as well as determination of downstream effects. DC vaccination serves as a model of early loss of B cell tolerance for lupus (1, 2). Adoptive transfer of apoptotic cell-laden myeloid DC markedly accelerated disease in the lupus-prone New Zealand White (NZW) and New Zealand Black (NZB) mouse strain, NZB/W F1 (1). In contrast, DC vaccination of wild-type (WT) mice led to lupus type autoantibodies (including anti-dsDNA) and IgG deposition in the glomeruli, but clinical disease was not observed (2). Because subclass analysis revealed a high IgG1 to IgG2a ratio of anti-DNA, it was suggested that skewing of autoantibodies toward the IgG1 isotype most likely accounted for their lack of pathogenicity. IgG2 Abs are known to promote inflammatory responses by preferential engagement of activating Fc{gamma}RIV on macrophages, and furthermore, this skewing requires signaling via TLR and MyD88 (3, 4). In contrast, IgG1 Abs promote inhibitory signals through preferential engagement of Fc{gamma}RIIB.

How mature DC are able to break tolerance to self-Ags in the DC vaccination model and whether this is a T cell-dependent process are unknown. Autoreactive B cells may become activated in a T cell-independent manner (5), and DC have been shown to directly interact with and present unprocessed Ags to B cells (6, 7, 8). We therefore examined the roles of conventional {alpha}beta T cells as well as nonconventional {gamma}{delta} T cells in promoting autoantibody production following DC vaccination. In view of the requirements for TLR stimulation in class switching, as mentioned above, as well the role of TLR in the induction of inflammatory cytokines in lupus (9), we also explored the requirements for TLR stimulation in the induction and maturation of autoantibodies in this model.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice

Female 6- to 8-wk-old B6 (B6) mice were purchased from The Jackson Laboratory. Mice deficient in beta{delta} TCR (beta{delta}–/–), {gamma}{delta} TCR ({gamma}{delta}–/–), and {alpha}beta TCR ({alpha}beta–/–) on the C57BL/6 background were purchased from The Jackson Laboratory. MyD88-deficient mice used in this study were provided by Dr. S. Akira (Osaka University, Osaka, Japan), and OT-II mice were provided by M. Bevan (University of Washington, Seattle, WA). Sex- and age-matched female B6 mice (The Jackson Laboratory) were used as WT controls. A group of 10 NZB/W F1 females was purchased from The Jackson Laboratory. Starting at week 20 the mice were bled to test for autoantibodies every 2 wk, and their urine was tested weekly for proteinuria by dipstick (Multistick; Fisher Scientific International). Mice were followed until death. All mice were bred and maintained in a specific pathogen-free facility, and all procedures involving animals were approved by the Institutional Animal Care and Use Committee at the University of Washington (Seattle, WA).

Monoclonal Abs, regulatory T cell (Treg) depletion, and flow cytometry analysis

Directly conjugated Abs reactive against murine cell surface markers (MHC class II, CD86, CD40, and CD11c) were purchased from Biolegend. The secondary reagents for ELISA were purchased from Jackson ImmunoResearch Laboratories. Before staining of DC, Fc receptors were blocked with unlabeled polyclonal mouse IgG (Sigma-Aldrich). For multicolor analysis, 0.2 million cells were incubated for 30 min at 4°C with directly conjugated Abs, and flow cytometry was performed on a FACScanto (BD Pharmingen). Flow cytometry analysis was done using FlowJo software. Mice were depleted of Tregs using an anti-IL-2 mAb, S4B6, according to the protocol of Setoguchi et al. (10). In brief mice were given S4B6 mAb (500 µg) on days –7 and –3 before DC injection on day 0. Mice were given a second depletion on days 21 and 25 with S4B6. Treg depletions were confirmed from study animals in the peripheral blood.

Generation of bone marrow-derived DC, immunization protocol, and cytokine analysis

Bone marrow suspensions were prepared as described (11). Erythrocyte-depleted bone marrow isolated from long bones was resuspended at one million cells/ml of culture medium (RPMI 1640 medium, 10% FCS, 50 µM 2-ME, 20 µg/ml gentamicin) in the presence of 20 ng/ml murine GM-CSF. Immature DC were harvested on day 6 and shown to be >80% CD11c+, MHC class IIhigh, CD86/CD40+. In some in vitro experiments, immature bone marrow-derived DC (CD11c+/MHC class IIlow) were flow sorted to >99% purity with similar results.

Immature day-6 DC were matured by various TLRs, including LPS (10 ng/ml unless indicated otherwise), CpG oligodeoxynucleotide 1826 (1 µM), and in some experiments, agonistic anti-CD40 mAb (clone 3/23, 1 µg/ml) as per published reports (12, 13). Cytokine production following DC or T cell (see below) activation was quantified by ELISA for TNF, IL-12p70, IL-23p19, IFN-{gamma}, and IL-4 according to the manufacturer’s protocol (Opti-EIA; BD Pharmingen) and (14). The DC were harvested after 6 h of in vitro culture and washed three times with PBS. One million cells were i.v. injected per mouse via tail vein. Injections were performed every 2 wk for a total of four injections, and mice were sacrificed 2 wk after the final injection at week 8. In most experiments the mice were bled for serum retroorbitally at week 3 as an early time point following the second injection.

For congenic DC transfer and survival analysis, we used DC from CD45.1 allotype marked B6 mice (5 x 106 DC/mouse) and harvested at 48 h posttransfer. For this experiments, some mice received TLR matured DC and/or additional i.p. LPS at time 0 and 24 h before harvest. Analysis was performed on CD11c-selected DC isolated from spleens.

In vivo T cell activation assay

Immature DC were loaded with OVA and then either cultured in medium or matured by multiple TLR agonists (LPS, 20 ng/ml or CpG, 1 µM) and anti-CD40 (1 µg/ml) for 6 h. A total of 5 x 106 DC were injected into each mouse followed by 5 x 106 OT-II T cells (CD45.1 allotype) 1 day later. Mice were sacrificed after 1 wk and splenic OT-II T cells were restimulated in vitro with 1 µM OVA peptide (OVA323–339) or anti-CD3/CD28. The supernatants were analyzed for IL-4 (Th2) and IFN-{gamma} (Th1) cytokines by ELISA as discussed.

Detection of autoantibodies

Serum anti-DNA IgG levels were quantified by sandwich ELISA as described (2). Briefly, polystyrene microtiter plates were coated with calf thymus DNA (Sigma-Aldrich) overnight at 4°C. After blocking of the plates with 1% OVA, test sera were added and incubated for 2 h at 1/100 dilution, washed, and reacted with alkaline phosphatase-conjugated goat anti-mouse IgG (Sigma-Aldrich) at a dilution of 1/2000. The reaction was developed with p-nitrophenyl phosphate, and the OD at 405 nm was determined. Serum IgG subclasses were quantified as described, except that the samples were reacted with goat Ab specific for IgG1 or IgG2a at 1/1000 dilution (Jackson ImmunoResearch Laboratories). For anti-histone assays, ELISA plates were coated overnight with H1 or H2A/H2B histones (Roche) at 10 µg/ml overnight, and anti-histone IgG was detected as discussed for IgG anti-DNA. For the DNA/H2A/H2B ELISA, the histones were added onto plates precoated with DNA. For anticardiolipin IgG levels, polystyrene plates were coated with cardiolipin by incubation with 100 µg/ml bovine cardiolipin (Avanti Polar Lipids) in 100% ethanol overnight at 4°C until completely evaporated. Plates were blocked with 1% BSA in PBS, and serum was added at 1/100 dilution. Anticardiolipin Abs were detected with alkaline phosphatase-conjugated goat anti-mouse IgG (Southern Biotechnology Associates). For all autoantibody assays, the results were expressed in arbitrary ELISA units relative to a standard positive sample derived from a pool of 9-mo-old MRL/lpr mice.

Histopathology

The kidneys were removed at the time that the mice were harvested. One kidney was fixed with 10% buffered formalin, embedded in paraffin, and sectioned before staining with H&E.

Statistical analyses

Statistical comparisons were compared by Student’s t test for normally distributed, and the Mann-Whitney rank sum test for non-normally distributed data. Values of p < 0.05 were considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Anti-dsDNA autoantibody production is absolutely T cell-dependent and has a positive requirement for both {alpha}beta and {gamma}{delta} subsets but is negatively regulated by Treg

Using a previously described protocol of DC vaccination that uses four i.v. injections of LPS-matured myeloid DC to induce lupus type autoantibodies (2), we first asked whether autoantibody production following DC transfer into normal B6 mice was T cell-dependent. In some experiments, LPS (10 µg) was given i.p. at the time of each injection to augment autoantibody responses (see below). beta{delta} TCR-deficient mice (beta{delta}–/–) that lack both the {alpha}beta and {gamma}{delta} subsets of T cells were used as DC recipients. In striking contrast to WT mice, DC vaccination of beta{delta}–/– mice failed to induce autoantibodies as determined by fluorescent antinuclear autoantibodies (data not shown) or anti-DNA ELISA (either total IgG, or IgG1 or IgG2a subclass) (Fig. 1, A–C). The Ab levels were essentially equivalent to uninjected control mice. These results indicate that loss of B cell tolerance leading to the production of autoantibodies in this model is absolutely T cell-dependent. Similar results were seen in a second set of mice following DC transfer in the absence of additional TLR stimulation (data not shown).


Figure 1
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  		FIGURE 1. Conventional {alpha}beta T cells ({alpha}beta) are absolutely required for autoantibody production following DC vaccination. A–C, Sera from WT or beta{delta} TCR-deficient (beta{delta}–/–) mice obtained at 8 wk following four DC injections (1 million LPS-matured DC per injection via tail vein) in combination with i.p. LPS (10 µg) were tested for anti-dsDNA autoantibodies, including total (A), IgG2a (B), and IgG1 (C) by ELISA at a dilution of 1/100. Results are expressed in arbitrary ELISA units (AEU) relative to a standard positive sample derived from a pool of aged MRL/lpr mice. D–F, Sera from WT or {alpha}beta TCR-deficient ({alpha}beta–/–) mice obtained 2 wk after the fourth injection at week 8 as above were tested for total (D), IgG2a (E), and IgG1 (F) anti-dsDNA Abs by ELISA. Shown are serum autoantibody levels as measured by ELISA, and expressed in arbitrary ELISA units. Each symbol represents an individual mouse. Horizontal line represents mean value, and controls were from saline-injected age-matched littermates. *, p < 0.01; **, p < 0.001 between WT and beta{delta} or {alpha}beta TCR-deficient mice. Data shown are representative of two independent experiments.

 
The majority of peripheral T cells are comprised of conventional T cells that have {alpha}beta receptors, but nonconventional T cells ({gamma}{delta}, NK, and NKT) have also been implicated in systemic autoimmunity. We therefore examined the relative contributions of {alpha}beta and {gamma}{delta} T cells to autoantibody production following DC vaccination using {alpha}beta TCR-deficient ({alpha}beta–/–) and {gamma}{delta} TCR-deficient ({gamma}{delta}–/–) mice as recipients. Abs directed against dsDNA (Fig. 1, D–F) as well as antinuclear Abs and anticardiolipin IgG (data not shown) were reduced in the {alpha}beta–/–, similar to the beta{delta}–/– mice at week 8. These results confirm the absolute requirement of conventional {alpha}beta T cells for autoantibody production in this DC transfer model.

Peng and Craft (15) previously reported that MRL/lpr mice deficient in conventional {alpha}/beta+ T cells developed milder disease characterized by the production of autoantibodies predominantly of the IgG1 subtype and attributed this autoimmunity to {gamma}{delta} T cells. When {gamma}{delta}–/– mice were vaccinated with DC, autoantibodies directed against DNA, including total IgG, IgG2a and IgG1 isotypes, were significantly reduced compared with controls at week 8 (Fig. 2, A–C), indicating that {gamma}{delta} T cells also contributed to autoantibody production in these mice. The {gamma}{delta}–/– mice developed low titers of autoantibodies that were consistently and significantly above control mice, indicating that, unlike the {alpha}beta–/– mice, low levels of anti-DNA could be generated in the absence of {gamma}{delta} T cells. Although Abs directed against cardiolipin were also reduced in the {gamma}{delta}–/– mice, this did not reach statistical significance (Fig. 2D). WT, beta{delta}–/–, and {alpha}beta–/– mice all failed to develop significant proteinuria or evidence of glomerulonephritis on light microscopy on sacrifice (data not shown).


Figure 2
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  		FIGURE 2. {gamma}{delta} T cells ({gamma}{delta}) contribute to anti-dsDNA autoantibody production following DC vaccination. A–C, Sera levels of IgG anti-dsDNA in WT and {gamma}{delta} TCR-deficient ({gamma}{delta}–/–) mice following DC vaccination at 8 wk were tested for total (A) and IgG2a (B), and IgG1 (C) anti-dsDNA or IgG anticardiolipin (D) by ELISA at a dilution of 1/100. Results are expressed in arbitrary ELISA units (AEU) relative to a standard positive sample derived from a pool of aged MRL/lpr mice as described in Fig. 1. Each symbol represents an individual mouse. Horizontal line represents mean value. *, p < 0.05; **, p < 0.005. ns, Not significant. E and F, Mice deficient in {gamma}{delta} T cells develop contact-dependent dermatitis. A representative rash observed on the back (E) along with histopathology of involved skin (F) demonstrating perivascular mononuclear cell infiltrate is shown. H&E stain was at original magnification of x40.

 
Interestingly, when {gamma}{delta} TCR-deficient mice (B6 background) were evaluated following the second injection at week 3, we observed the development of patchy skins lesions with hair loss over the upper back (Fig. 2, E and F). These lesions, which were not observed in untreated C7BL/6.{gamma}{delta}–/– mice, disappeared when nitrile gloves were substituted for latex, suggesting that latex stimulated the dermatitis. The dermatitis was similar to that displayed by the {gamma}{delta}-deficient NOD and FVB strains (16). Because the dermatitis was not observed in beta{delta}–/– mice (lacking both {alpha}beta and {gamma}{delta} T cells) that received DC, these results suggest that activated DC provoked the allergic response that is {alpha}beta T cell mediated but usually suppressed by {gamma}{delta} T cells.

To further analyze T cell control of autoantibody production following DC vaccination, we depleted circulating Treg by two injections of anti-IL-2 mAb (10). Circulating Foxp3+ cells showed an ~8-fold depletion when analyzed at either day 10 (Fig. 3A) or day 30 (data not shown) by this method. DC vaccination following depletion of Treg induced a modest increase in total anti-DNA levels at 8 wk compared with mice injected with DC and isotype control Ab (p = 0.05) or anti-IL-2 alone (Fig. 3B). Analysis of subclasses revealed that the dominant increase in anti-DNA was IgG1, which was highly significantly increased compared with mice injected with DC and isotype control mAb (Fig. 3C, p = 0.008). IgG2a levels were unaffected (Fig. 3D). Therefore, Tregs influence the quantity but not the quality of anti-DNA induced in this model. Similar to control mice, we observed neither proteinuria nor significant levels of glomerulonephritis at week 8 in the Treg-depleted mice (data not shown).


Figure 3
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  		FIGURE 3. Treg depletion augments IgG1 anti-DNA responses. A, Treg depletion (CD4+, FoxP3+) 10 days after administration of the anti-IL-2 mAb S4B6. Representative flow cytometry plots of peripheral blood from mice treated with S4B6 or normal rat IgG in the presence or absence of DC injections as indicated. Anti-dsDNA (B), IgG1 anti-DNA (C), and anti-IgG2a anti-DNA (D) levels in Treg-depleted (S4B6) or control IgG-treated recipient mice at 8 wk following four vaccinations of mature bone marrow-derived DC. Results are expressed in arbitrary ELISA units (AEU) relative to a standard positive sample derived from a pool of aged MRL/lpr mice. The mice received DC transfers following S4B6-mediated Treg depletion or treatment with control rat IgG as indicated. Each symbol represents an individual mouse. Controls were from Treg-depleted littermates that did not receive DC. *, p = 0.05; **, p < 0.009.

 
Antichromatin autoantibodies induced by DC vaccination produce high titer anti-H1, but low titer anti-H2A/H2B/DNA responses

The evolution of autoantibodies to the components of chromatin that include DNA, nucleosomes (including H2A/H2B/DNA) and individual histones have been reported in both human systemic lupus erythematosus (SLE) and in murine models (17, 18, 19, 20, 21). This response is known to be T cell-driven and, over time, results in epitope spreading and development of high affinity Abs of the IgG2a subclass against multiple components of chromatin (22). Because we conclusively demonstrated that the anti-DNA response following DC vaccination was T cell-dependent, we asked whether the humoral immune response against chromatin evolved in a similar way to spontaneous lupus in NZB/W F1 mice. Female NZB/W F1 mice develop autoantibodies and resultant nephritis between 5 and 9 mo of age, although stochastic factors determining anti-DNA levels and clinical severity of disease in individual mice during this window remain undefined (23). In a cohort of unmanipulated NZB/W F1 mice (n = 10), we observed the development of high titer anti-DNA, antinucleosome (H2A/H2B/DNA) and anti-H2A/H2B IgG between 26 and 36 wk of age, although there was wide variation in titers and time of onset between individual mice. After four injections, DC-vaccinated mice produced Abs to these same three Ags, albeit at significantly lower levels (Fig. 4, A–C). In contrast, DC-vaccinated mice produced higher, albeit not statistically significant, levels of Abs to histones H1 than 30 wk old NZB/W F1 mice (Fig. 4D). Therefore, qualitative analysis of antichromatin autoantibodies following DC vaccination reveal a moderate anti-DNA and anti-histone H1 response but failure to progress to high titer anti-DNA or antinucleosome responses, which others have shown to be pathogenic (24).


Figure 4
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  		FIGURE 4. Fine specificities of antichromatin autoantibodies in DC-vaccinated and NZB/W F1 mice. A–D, Sera from a cohort of NZB/W F1 mice (n = 10, 18–36 wk of age) or from DC vaccinated mice (n = 8) at 8 wk following four injections of DCs (as in Fig. 1) were tested for Abs to dsDNA (A), nucleosomes (H2A/H2B/DNA) (B), histone H2A/H2B (C), or histone H1 (D). Results shown are serum autoantibody levels as measured by ELISA and expressed as the mean ± SE in arbitrary ELISA units (AEU) relative to a standard positive sample derived from a pool of aged MRL/lpr mice. To facilitate more consistent presentation of the NZB/W cohort, the autoantibody levels are presented in weeks before the onset of 3+ proteinuria (>300 mg/dL). NZB/W Ab levels to DNA, nucleosome, and H2A/H2B were significantly higher at the 0 week time point compared with DC-vaccinated mice when analyzed by the Mann-Whitney rank sum test. Anti-H1 levels were higher in DC-vaccinated mice but did not reach statistical significance (p = 0.06).

 
Synergistic or repetitive TLR signaling enhances Th1 skewing but does not induce pathogenic autoantibodies

Abs of the IgG2 isotype have potent effector function through Fc{gamma}R engagement and/or complement activation and are predominantly responsible for lupus nephritis, whereas IgG1 preferentially engages inhibitory Fc{gamma}RIIB (4). Because DC vaccination in normal mice led to a reduced ratio of IgG2a to IgG1 anti-DNA Abs compared with MRL/lpr mice (2) and failed to evolve to high titer anti-nucleosome responses (Fig. 4), we attempted several strategies to manipulate the innate and/or adaptive immune response to produce pathogenic Abs. LPS activation of DC is thought to induce a Th1 response, presumably through induction of IL-12, which promotes IFN-{gamma} production by T cells (25). Because it was recently demonstrated that combinations of TLR agonists can augment IL-12p70 up to 100-fold (12, 13), we attempted to more intensively skew cytokines toward the induction of IL-12.

Synergistic DC activation by LPS plus CpG markedly enhanced the production of IL-12p70, TNF, and IL-23p19 but not type I IFN or IL-10 by DC (Fig. 5A and data not shown) and also altered the Th1/Th2 skewing for T cells responsive to a model test Ag in vivo. First, to validate that synergistic T cell activation does not reduce DC survival in vivo, we analyzed DC numbers using a congenic marker (CD45.1) to follow untreated or TLR-matured DC after adoptive transfer. As shown in Fig. 5B, DC survival as measured by recovery of transferred cells, increased 2-fold following TLR maturation stimuli, given either in vitro or i.p. compared with nonstimulated DC 72 h after transfer. To determine whether the enhanced IL-12 cytokine production can, in fact, skew T cell responses to Th1, OT-II TCR transgenic T cells and OVA-loaded DC were adoptively transferred to B6 mice, mice were sacrificed 7 days later and cytokine responses analyzed in an in vitro restimulation assay. In contrast to unstimulated cells, which showed relatively high IL-4 and low IFN-{gamma} production, synergistic TLR activation reversed this profile with lower IL-4 and markedly increased IFN-{gamma} production in response to anti-CD3/CD28 or OVA stimulation (Fig. 5, C and D). Although synergistic TLR stimulation of the DC ex vivo does produce Th1 skewing in vivo, autoreactive T cells may not behave in the same way. Together, these experiments demonstrate that synergistic TLR activation not only induces massive IL-12 up-regulation, but maintains or increases DC survival and promotes Th1 responses, at least in response to a non self Ag presented by the DCs.


Figure 5
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  		FIGURE 5. TLR agonists promote DC survival and Th1 skewing in response to a model Ag. A, Synergistic stimulation of DC by combinations of TLR agonists. Day 6 bone marrow-derived DC were stimulated in vitro with LPS alone or in combination with CpG 1826 (1 µM) and agonistic anti-CD40 (1 µg/ml) as indicated. Supernatants were tested at 24 h for TNF, IL-23, and IL-12p70 by ELISA. The results are presented as the mean of duplicates of one or four representative experiments as described in Materials and Methods. B, Day-6 CD45.1 myeloid DC were either untreated immature DC (iDC) or DC matured by synergistic TLR stimuli (mDC) and adoptively transferred into CD45.2 hosts (5 x 106 DC/mouse). Some mice also received i.p. LPS. After 72 h, the splenic CD45.1 DC were enumerated by flow cytometry. Results are the mean recovery of CD45.1+/CD11c+ DC from two separate experiments. C and D, Th1 skewing of T cell responses in vivo following combination TLR activation. Mice were injected with OVA-peptide loaded DC followed 1 day later by 5 x 106 OT-II T cells specific to OVA protein. Before peptide loading, the DC were either untreated or matured with TLR as indicated. One group received additional in vivo TLR stimulation (IP LPS) as indicated. Seven days later, spleens were harvested and T cell cytokine responses were measured by ex vivo stimulation by anti-CD3/CD28 or OVA peptide (1 µM). Shown are the concentrations of IFN-{gamma} or IL-4 in culture supernatants.

 
To test whether a similar approach could alter the Th1 to Th2 ratio of autoreactive T cells leading to IgG2a anti-DNA Abs, mice were vaccinated with DC activated by LPS alone, LPS in combination with CpG, or LPS and CpG plus agonistic anti-CD40 Ab. Despite the substantial increase in DC IL-12p70 production in vitro, the synergistically activated DC stimulated anti-DNA Abs at levels similar to LPS alone, and no differences were observed in total or IgG2a anti-DNA or the relative ratios of IgG2a and IgG1 anti-DNA (Fig. 6, A and B, and data not shown). No significant differences in the total numbers of splenic B cell, T cell, and DC populations, levels of proteinuria (<100 µg/ml) nor renal deposition of IgG were observed between experimental groups (data not shown). These results indicate that TLR-mediated enhancement of DC IL-12 as well as other proinflammatory cytokines is insufficient to induce isotype switching and changes in the quality of antichromatin autoantibodies or nephropathology in B6 mice. Thus, additional checkpoints on class switching to pathogenic autoantibodies are present.


Figure 6
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  		FIGURE 6. Hyperinduction of Th1 cytokines by stimulation of DC with multiple TLR agonists in vitro fails to enhance IgG2a subclass autoantibodies. A and B, Anti-dsDNA Abs induced by adoptive transfer of LPS matured or synergistically activated (LPS/CpG or LPS/CpG/anti-CD40) bone marrow-derived DC into syngeneic mice. Sera obtained at week 8 were tested for total IgG anti-DNA (B) or IgG2a anti-DNA (C) by ELISA and expressed in arbitrary ELISA units (AEU) relative to a standard positive sample derived from a pool of aged MRL/lpr mice. Each symbol represents an individual mouse. Horizontal line represents the mean, and controls from saline-injected age-matched littermates are shown. C, Mice received four i.v. injections of mature (LPS stimulated) DC, as in Fig. 1, alone (DC), together with i.p. LPS (10 µg) (DC + IP LPS), or i.p. LPS alone (IP LPS). At 8 wk, total anti dsDNA IgG autoantibodies were measured by ELISA. **, p < 0.005.

 
Persistent TLR signaling induced by repeated i.p. injections of TLR ligands was shown to break CD8 T cell tolerance following the vaccination of Ag-loaded DC (26). To test whether systemic autoimmunity could be augmented by persistent TLR stimulation in vivo, we combined IV DC vaccination with simultaneous LPS (TLR4) i.p. stimulation, and observed significantly enhanced levels of total (Fig. 6C) and IgG2a anti-DNA as well as mild elevation in antinuclear Ab titers (data not shown). These results demonstrate that delivery of a second TLR stimulus has a much more potent effect in vivo than in vitro, suggesting a possible effect on cells other than the donor DC.

Initial loss of B cell tolerance following DC vaccination does not require signaling through MyD88 but this pathway does impact class switching

Although immune complex mediated TLR activation augments inflammation (27), it is not known whether TLR activation is necessary for initiation of autoimmunity in SLE and what cell types may be effected (28). We therefore compared the quantity and quality of anti-DNA responses following vaccination of MyD88 deficient DCs into WT hosts and vaccination of WT DCs into MyD88–/– hosts. MyD88 is an adapter that is required for signaling by all of the TLRs except for TLR3, which exclusively uses the Trif adaptor, and TLR4, which uses both MyD88 and Trif (25). Because we have previously reported that TLR stimulation of the donor DC by LPS augmented costimulatory molecule expression as well as cytokine production resulting in significantly increased anti-DNA Ab production in recipient mice (2), we first asked whether costimulatory molecule up-regulation on MyD88–/– DC in the absence of cytokine production, would be sufficient to induce autoantibodies upon transfer into WT recipients.

We confirmed (29) that LPS activation of MyD88–/– DC, signaling exclusively through Trif, up-regulated CD86, CD40, and MHC class II expression equivalent to WT, but failed to produce significant quantities of proinflammatory cytokines, including IL-12 and TNF (Fig. 7, A and B and data not shown). Following vaccination with either WT or MyD88–/– donor DC matured in vitro with LPS, equivalent levels of total anti-DNA autoantibodies were observed at week 8 (Fig. 7C), suggesting that donor DC cytokine production was not essential for the initial loss of B cell tolerance in response to mature DC vaccination. However, whereas vaccination of WT DC with an in vivo TLR stimulus (i.p. LPS) resulted in a significant increase in the total IgG anti-DNA response, MyD88–/– DC failed to induce the same response to the in vivo TLR stimulus (Fig. 7C). Furthermore, WT mice vaccinated with MyD88–/– DC, either alone or in the presence of additional i.p. LPS, failed to increase IgG2a anti-DNA levels (Fig. 7D) but maintained levels of IgG1 anti-DNA, resulting in a reduced ratio of anti-DNA IgG2a to IgG1 in these same mice (Fig. 7E and data not shown). In conclusion, although increased costimulatory molecule and MHC (presumably with self Ag) expression was sufficient to initiate the production of anti-DNA Abs by MyD88–/– DCs, MyD88 expression in the donor DCs was necessary for in vivo LPS augmentation of total IgG anti-DNA levels and for switching to the IgG2a subclass.


Figure 7
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  		FIGURE 7. Donor DC MyD88 is dispensable for initiation of anti-DNA Ab production, but is required for subclass switching to IgG2a. A, MyD88-deficient DC up-regulate CD86 in response to LPS but not CpG. Results are the mean fluorescence intensity (MFI) of CD86 of day-6 WT or MyD88–/– bone marrow-derived DC following overnight exposure to medium, LPS, or CpG as indicated. B, MyD88 is required for production of IL-12p70 in response of LPS. IL-12p70 levels were measured by cytokine ELISA from WT or MyD88–/– bone marrow-derived DC as in A. For both A and B, data are representative of three experiments performed. C–E, Serum total IgG (C), IgG2a (D), and IgG1 (E) anti-dsDNA levels in WT recipient mice at 8 wk following four vaccinations of mature bone marrow-derived DC from either WT or MyD88–/– mice. Results are expressed in (AEU) relative to a standard positive sample derived from a pool of aged MRL/lpr mice. The mice received DC transfers either alone or with additional i.p. LPS 24 h following each DC transfer as indicated. Each symbol represents an individual mouse. Controls were from i.p. LPS-injected littermates. *, p < 0.01; **, p < 0.005. ns, Not significant.

 
To address the role of TLR signaling in the non-DC compartment, we vaccinated MyD88–/– hosts with mature WT DC. In the absence of in vivo TLR stimulation, WT and MyD88–/– recipients developed equivalent levels of total IgG anti-DNA and IgG1 subtypes (Fig. 8, A ands B and data not shown). We did note, however, a significant reduction in levels of IgG2a anti-DNA and anticardiolipin IgG in MyD88–/– recipient mice (Fig. 8, C and D). To determine whether in vivo LPS could rescue the IgG2a class switch defect, MyD88–/– recipients were vaccinated with WT DC and boosted with i.v. administered LPS. Under these conditions, the WT and MyD88–/– hosts produced equivalent moderate levels of anti-DNA autoantibodies, including total IgG, IgG2a, and IgG1 subtypes as well as anticardiolipin autoantibodies (Fig. 8, E–H, and data not shown). These findings indicate that activation of TLR pathways are not absolutely required for initial loss of B cell tolerance in response to DC vaccination, but that lack of signaling through host MyD88 impairs B cell isotype switching to IgG2a, consistent with a previous report (3). Together with the results in Fig. 7, MyD88 signaling on both the donor DC and recipient B cells are therefore required for switching to IgG2a anti-DNA Abs, although we cannot exclude a role for LPS activation of other cell types.


Figure 8
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  		FIGURE 8. Host MyD88 is dispensable for initiation of anti-DNA and anticardiolipin Ab production, but is required for subclass switching to IgG2a following DC vaccination. A–F, Sera levels of anti-dsDNA IgG in WT and MyD88–/– mice following DC vaccination at 8 wk for total (A and E), IgG1 (B and F), IgG2a (C and G) anti-dsDNA and anticardiolipin (D and H) were tested by ELISA as described in Fig. 1. Results are expressed in arbitrary ELISA units (AEU) relative to a standard positive sample derived from a pool of aged MRL/lpr mice. Each symbol represents an individual mouse, and horizontal line represents mean value. WT or MyD88–/– recipient mice received transfers of DC alone (A–D) or DC with i.p. LPS (10 µg) (DC + IP LPS) (E–H) 24 h after each DC transfer as indicated. **, p < 0.005.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Many immune abnormalities have been implicated in the predisposition to murine lupus (30). In at least three cases, abnormalities in DC themselves have been shown to orchestrate lupus-like diseases. These examples include accelerated maturation of DC in the sle3 congenic strain (31), DC-specific overexpression of the antiapoptotic baculovirus p35 protein (32) and DC-specific SOCS1 deficiency (33). Because we and others have shown that repetitive immune stimulation by mature DC readily induce lupus autoantibodies (1, 2), this model affords an opportunity to dissect the cellular and molecular pathways required to break B cell tolerance as well as elucidation of the factors required to generate pathogenic Abs. We report that T cells are absolutely required for generation of lupus autoantibodies following DC vaccination and that both {alpha}beta and {gamma}{delta} T cells contribute to the autoimmune process.

We previously showed that LPS-activated DC, but not activated macrophages (that produced much higher levels of TNF-{alpha} and IL-1), induced high levels of anti-dsDNA IgG in normal mice (2). Furthermore, high titer anti-dsDNA production was lost following paraformaldehyde fixation of DC. Together, these observations suggested that DC play a specific role, most likely presentation of self Ag to T cells (2). We confirmed that T cells were absolutely required for autoantibody production in this model and that the conventional {alpha}/beta T cell subset plays the dominant role in providing the necessary help to autoreactive B cells. However, because B6 mice lacking {gamma}{delta} T cells also had reduced autoantibody levels following DC vaccination, {gamma}{delta} T cells must also promote autoantibody production, consistent with previous studies in MRL mice (15).

{gamma}{delta} T cells, typically enriched in epithelial tissues, are among the earliest T cells to respond to infection and immunization (34). They may recognize Ags on mature DC and "prime" {alpha}beta T cells for high titer responses, in either a Th1 or Th2 fashion (35). In their absence, therefore, overall autoantibody responses may be expected to be lower as was observed in this study. We therefore favor the model that initial recognition of self Ags through nonclassical (e.g., CD1d) presentation of Ags is by {gamma}{delta} T cells, consistent with the proposed role for this T cell subset in recognition of self Ags (36). Similar to the {gamma}{delta}-deficient mice used in this study, mice deficient in CD1d developed exacerbated dermatitis and reduced Th2 cytokines on the MRL background (37). Because human V{delta}2 T cells have been shown themselves to present Ag, we cannot entirely exclude a role for direct Ag presentation by murine {gamma}{delta} T cells assuming that they can take up and process Ag derived from DC as demonstrated for human {gamma}{delta} T cells (38).

We observed the development of contact dermatitis in {gamma}{delta}–/–, but not beta{delta}–/–, mice vaccinated with mature DC, consistent with the local suppressive effects of {gamma}{delta} T cells on conventional T cells in the skin (16). Notably, the dermatitis appeared only following transfer of mature DC into {gamma}{delta}-deficient B6 mice, indicating that in the absence of {gamma}{delta} T cells the transferred DC promoted inflammation. Various DC subsets, including plasmacytoid DC and both immature and mature myeloid DC, have been shown to modulate allergic disease, including dermatitis (39), presumably through the alteration of Th1, Th2, and/or Treg subsets by local factors, including TSLP (40) and RANK ligand (41). It will be of interest to determine which of these mechanisms is impacted by the absence of inhibitory skin {gamma}{delta}+ T cells.

The evolution of autoantibodies in lupus, including the components of chromatin, has been studied in both mice and humans. Earlier reports suggested that the initial autoimmune response is primarily directed against conformational epitopes on chromatin and H2A/H2B/DNA subnucleosome (22). More recent preliminary studies suggest that the loss of tolerance to the linker histones, including H1, is an early and possibly initial event in the pathogenesis of autoimmunity to chromatin (21). Although DC vaccinated mice developed moderate titers of anti-dsDNA Abs as previously reported (2), their antinucleosomal autoantibodies never exceeded levels observed in 6-mo-old, prenephritic NZB/W F1 mice and were significantly lower than nephritic mice. They did, however, develop high levels of anti-H1 Abs. This observation suggests that repetitive DC stimulation by itself is unable to facilitate epitope spreading from the exposed linker (H1) histones to nucleosomal core Ags. High-titer antinucleosomal autoantibodies may be particularly pathogenic due to the ability to bind heparan sulfate and nucleosomes trapped in the glomerular basement membrane (24, 42). Thus the absence of renal pathology in DC vaccinated mice compared with the NZB/W F1 may be due to both qualitative (autoantibody evolution) as well as quantitative differences.

Because DC vaccination in the autoimmune prone NZB/W mice leads to marked acceleration of renal failure and death (1), there may be critical regulatory checkpoints between initial autoantibody production and pathogeneic autoantibody production that are present in C57BL/6 mice but absent in NZB/W F1 mice. Activators of innate immunity, including DC vaccination (1, 2), IFNs (27, 43), or overexpression of the yaa gene (44, 45) may only serve to accelerate disease but are not sufficient for induce severe tissue damage in normal strain backgrounds. Whereas innate activation alone does not overcome tolerance checkpoints, C57BL/6 mice develop severe SLE under circumstances affecting B cell and T cell regulation (46, 47). Because it takes NZB/W F1 mice ~7 mo to develop nephritis, more prolonged exposure might be required and future experiments will be directed toward longer provocations.

Although we had previously established that, in normal mice, immune responses to self Ags could be enhanced by adoptive transfer of mature DC, we did not observe significant end-organ damage. Failure to induce tissue injury was attributed to the relatively high IgG1 to IgG2a anti-DNA Ab ratio observed (2). To overcome this checkpoint at the level of the adaptive immune response, we used a strategy to enhance DC production of IL-12 and hence skew T cell responses to Th1 (IFN-{gamma}), which promotes IgG2a responses. Despite the clear in vitro increase of IL-12p70 levels from the combination of CpG and LPS as reported by others (12, 13), as well as Th1 skewing to a model test Ag, OVA in vivo, this strategy failed to alter anti-DNA IgG subclass skewing or to cause tissue injury. This result is similar to that observed by Evel-Kabler et al. (33) who also failed to induce autoimmunity following the transfer of WT DC stimulated with exogenous TLR ligands or adenovirus overexpressing IL-12 and indicates that powerful additional tolerogenic influences exist on autoreactive T cells, B cells, or both.

The role of TLRs in promoting autoimmunity has been demonstrated in numerous murine models of both systemic as well as organ-specific disease (26, 48, 49, 50). Consistent with these studies, we observed that providing a second in vivo TLR stimulus following DC vaccination significantly enhanced autoantibody production. To more precisely dissect the requirement for TLR stimulation in the initial loss of tolerance, we evaluated autoantibody production using MyD88 deficient DC or MyD88–/– hosts. WT mice that received MyD88–/– DC (that express high levels of costimulatory molecules, but lack cytokine production) and MyD88–/– hosts that received WT DC produced anti-DNA levels equivalent to controls. These observations indicate that MyD88 dependent signaling in neither donor DC nor host B cells is required for the initial autoantibody response. However, in the absence of MyD88 signaling in either donor DC or host B cells, IgG2a anti-DNA levels were reduced, consistent with the results of some (3) but not other (51, 52) studies indicating a crucial role for TLR/MyD88 in the production of class switched IgG2 autoantibodies. Together, these results demonstrate that Ag presentation by DC that express high levels of costimulatory molecules is sufficient to break tolerance; however, the development of IgG2a autoantibodies requires signaling through MyD88. The requirement for a B cell class switch factor may also explain why synergistic activation of DCs was insufficient to produce pathogenic anti-DNA Abs.

In conclusion, DC vaccination induces autoantibodies with many similarities to spontaneous models of murine lupus. Autoantibody production requires both {alpha}beta and {gamma}{delta} T cells, and can be enhanced by TLR signaling, although this latter pathway is most important for the development of IgG2a subclass autoantibodies. Normal mice fail to develop clinical disease following DC vaccination. This result is explained, in part, by a skewing of anti-chromatin Abs to the suppressive IgG1 isotype and also by differences in the fine specificities of these Abs. In comparison to NZB/W F1 sera, DC vaccinated mice had responses that were at least equivalent against histone H1 but diminished to nucleosomes. This mouse model may therefore be most relevant to a significant population of individuals with serologic but not clinical manifestations of SLE (including relatives of bona fide patients or drug-induced SLE) (22) that apparently maintain critical regulatory pathways that prevent production of pathogenic autoantibodies. Although Treg appeared to control the magnitude of the anti-DNA response, other critical regulatory pathways that control isotype switching to IgG2a subclass, remain to be discovered.


   Acknowledgments
 
We thank Drs. Jolanta Kowalewska and Melanie Kuechle for help with renal and skin pathology, respectively, Dr. Shizuo Akira for sharing of the MyD88–/– mice, and Dr. John Hardin and members of the Elkon lab for helpful comments.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


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

1 This work is supported by Grant K08AR052804-02 (to D.A.M.) and Grant AR48796 (to K.B.E.) from National Institute of Arthritis and Musculoskeletal and Skin Diseases. D.A.M. is also the recipient of an Howard Hughes Medical Institute Physician-Scientist Early Career Award. Back

2 Current address: Amgen Inc., Mail Stop AW2-D 3262, 1201 Amgen Court West, Seattle, WA 98119-3105. Back

3 Address correspondence and reprint requests to Dr. Keith B. Elkon, Division of Rheumatology, University of Washington, 1959 NE Pacific Avenue, Box 356428, Seattle, WA 98195. E-mail address: elkon{at}u.washington.edu Back

4 Abbreviations used in this paper: DC, dendritic cell; NZB, New Zealand Black; NZW, New Zealand White; Treg, regulatory T cell; SLE, systemic lupus erythematosus; WT, wild type. Back

Received for publication August 3, 2007. Accepted for publication August 24, 2007.


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 Materials and Methods
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 Disclosures
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