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The Journal of Immunology, 2006, 177: 5307-5316.
Copyright © 2006 by The American Association of Immunologists, Inc.

Absence of Innate MyD88 Signaling Promotes Inducible Allograft Acceptance1

Wendy E. Walker2,*, Isam W. Nasr2,*, Geoffrey Camirand*, Bethany M. Tesar*, Carmen J. Booth{dagger} and Daniel R. Goldstein3,*

* Department of Internal Medicine, Yale University School of Medicine; and {dagger} Section of Comparative Medicine, Yale University School of Medicine, New Haven, CT 06520


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Prior experimental strategies to induce transplantation tolerance have focused largely on modifying adaptive immunity. However, less is known concerning the role of innate immune signaling in the induction of transplantation tolerance. Using a highly immunogenic murine skin transplant model that resists transplantation tolerance induction when innate immunity is preserved, we show that absence of MyD88, a key innate Toll like receptor signal adaptor, abrogates this resistance and facilitates inducible allograft acceptance. In our model, absence of MyD88 impairs inflammatory dendritic cell responses that reduce T cell activation. This effect increases T cell susceptibility to suppression mediated by CD4+CD25+ regulatory T cells. Therefore, this study provides evidence that absence of MyD88 promotes inducible allograft acceptance and implies that inhibiting innate immunity may be a potential, clinically relevant strategy to facilitate transplantation tolerance.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Solid organ transplantation is the treatment of choice for end stage disease of various organs (1, 2). The success of transplantation has largely been due to improvements in both surgical techniques and immunosuppressive medications that impair the adaptive immune system (3). Despite this success, recipients of solid organ transplantation manifest excess mortality and morbidity due to the side effects of lifelong immunosuppression and graft loss from chronic rejection (4, 5). Hence, protocols that induce transplantation tolerance will be a great advance for transplantation as well as for the prevention and treatment of autoimmune disease.

Importantly, the majority of experimental studies examining tolerance induction have focused on adaptive immunity. However, the innate immune system, which provides the initial response to noxious stimuli (3), is essential for triggering the adaptive immune response. Less is known concerning the mechanisms by which the innate immune system is activated in alloimmunity. Elucidation of the role of innate immunity during allotransplantation could provide potential novel targets for promoting transplant tolerance (3).

Innate immunity represents the first line of defense in response to microbial invasion (6). TLRs are critical sentinel receptors of the innate system that sense the presence of conserved microbial motifs (6). Additionally, TLRs sense nonbacterial products such as RNA, DNA, and chromatin and may play a role in autoimmunity (7). TLRs are expressed on dendritic cells (DCs)4 and other cell types including epithelial and endothelial cells (6, 8), and to date at least 11 TLRs have been discovered. TLRs signal through specific adaptor proteins (e.g., MyD88), which leads to DC maturation and priming of naive T cells (8). Hence, innate TLR-driven immunity initiates adaptive immune function (9, 10).

Although the signals that promote innate immune system activation in alloimmunity are poorly understood, TLRs appear to be involved. For example, TLR-driven MyD88-dependent immunity is critical for the rejection of minor mismatched skin allografts by inducing DC maturation, priming of graft-reactive T cells, and inducing Th1 immunity (11). Additionally, innate immunity via TLR signaling plays an important role in ischemia reperfusion injury (12, 13, 14). In a fully allogeneic skin transplant model, MyD88 signaling was not required for rejection in untreated recipients, although Th1 cytokines were reduced and MyD88–/– recipients manifest a delayed ability to reject vascularized allografts (15). Nonetheless, TLR signaling through MyD88 may still contribute to the rejection response in fully mismatched recipients and inhibition of such signals could promote induction of tolerance. Alternatively, Th1 cytokines may be required for tolerance (16, 17). Furthermore, TLR activation may have disparate effects on regulatory T cell (T regs) function (9, 10). Thus, in the current study, we address the role of MyD88 signaling in the induction of transplant tolerance. To examine this, we used a fully allogeneic skin allograft model because such allografts, in contrast to cardiac and islet allografts, are more resistant to tolerance induction (18, 19, 20, 21). We found that absence of MyD88 allows long term skin allograft survival under cover of costimulatory blocking agents that are normally ineffective in a highly resistant skin allograft model (20). Importantly, MyD88-deficient recipients exhibit impaired DC inflammatory responses that reduce T cell proliferation and increase T effector susceptibility to the suppressive effects of CD4+CD25+ T regs.


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

B6.129/SvJ-MyD88tm1AK1 mice were generously provided by Dr. S. Akira (Osaka University, Osaka, Japan) and were backcrossed 10 times onto both the C57BL/6 (H2b) and BALB/c (H2d) backgrounds (designated as B6.MyD88–/– and BALB/c.MyD88–/– mice, respectively). Caspase 1-deficient mice (B6.129/SvJ-Caps-1tm1Flv), referred to as B6.ICE–/–, were provided by Dr. R. Flavell (Yale University, New Haven, CT). B6.129S7-RAG1TM1Mom (designated as B6.RAG–/–), B6, and BALB/c (H2d) mice were purchased from The Jackson Laboratory. CBA mice (H2k) were purchased from the National Cancer Institute. All mice were kept in pathogen-free conditions and prophylactically given sulfatrim antibiotics intermittently in the drinking water. The Yale University Institutional Animal Care and Use Committee approved use of animals in this study.

Skin transplantation and Ab treatment

Full-thickness trunk skin was transplanted from 6- to 8-wk donor mice (BALB/c-H2d) and were sutured on B6. recipients (H2b) as previously described (15). Rejection was defined as graft necrosis of the entire graft area. Invariably, recipients that accepted their allografts for >150 days demonstrated preserved graft size, hair growth, and had no cellular infiltration or evidence of inflammation demonstrated by histology (data not shown). The experimental tolerance protocol consisted of anti-CD40L (anti-CD154, clone MR1) 500 µg/mouse on days +2, +4, +6, and +8 relative to transplantation and CTLA-4 Ig 500 µg/mouse on day +2 relative to transplantation both via i.p. injection (reagents generously provided by C. Larsen, Emory University, Atlanta, GA). Anti-CD25 (clone PC61) was generously provided by D. Rothstein (Yale University), and 150 µg/mouse was given on day –2 i.p. before transplantation. We determined that this treatment depleted the CD4+CD25+ subpopulation by 90% and also specifically depleted Foxp3+ cells (data not shown). Isotype control Ig 150 µg/mouse (IgG1) was obtained from Bioexpress. rIL-6, monoclonal IL-6 inhibiting Ab (2 µg/ml) and isotype control were purchased from R & D systems.

Adoptive transfer

B6.RAG–/– mice that were transplanted with MyD88+/+ BALB/c skin grafts 2 wk previously were adoptively transferred, via i.v. tail vein injection with 2 x 105 CD4+CD25+ T regs) harvested from B6.MyD88–/– recipients that had accepted their BALB/c.MyD88–/– allografts for >150 days and cotransfused with 1 x 105 MyD88+/+ B6.CD4+CD25– T effectors from naive mice. Similar experiments were performed with CD4+CD25+ T regs purified from MyD88-sufficient groups at >150 days after transplantation. This group invariably had rejected their allografts at this time point.

ELISPOT

ELISPOT was performed as per previously published work (22). To assess the T cell recall response after transplantation, recipient spleen cells (0.1–1 x 106) were harvested at day +14 posttransplantation (or other indicated time points), purified for CD4+ and CD8+ T cells as described below, and cultured with 0.3–3 x 105 irradiated (28 Gy) donor stimulator spleen cells per well. Third party donor spleen cells and naive responder cell controls were included. For B6.MyD88–/– recipients, BALB/c.MyD88–/– stimulators were used, although similar results were obtained when MyD88+/+ stimulators were used for mutant responders and vice versa. For the DC ELISPOT assay, CD11c+ splenic cells were purified as described below at 1 wk after transplantation. This time point was identified as the optimal time to assess cytokine production in preliminary experiments using MyD88+/+-transplanted mice (data not shown). The DCs were plated (3 x 105/well) overnight in 96-well ELISPOT plates coated with the Ab of interest. No ex vivo stimulation was used for the DC experiments, and results were corrected for baseline levels of cytokine-producing cells via use of DCs from untransplanted control mice from the representative groups. All cells were cultured in RPMI, 10% FCS plus penicillin streptomycin during the ELISPOT procedure. Plates were read on a CTL automatic ELISPOT reader and analyzed using Immunospot 3.1 software (CTL). Results were expressed as spots per 1 x 106 cells or otherwise indicated.

Cell sorting and flow cytometric staining

After staining for cell surface markers, lymphoid cells were fixed and permeabilized overnight and then stained with Foxp3-specific PE-conjugated rat anti-mouse mAb (clone FJK-16S, IgG2a) or isotype control at 4°C according to the manufacturer’s instructions (eBiosciences). To purify CD4+ T effectors, spleen cells were harvested from mutant and MyD88+/+ mice, respectively, and purified via magnetic negative selection using the EasySep magnetic sorting system (StemCell Technologies). A similar protocol was used for purifying CD8+ T cells. The CD4+ cells were then purified for effectors or T regs (>95% purity) by staining with FITC-conjugated rat anti-mouse CD25, PE-conjugated rat anti-mouse CD45RB, and Cychrome-conjugated rat anti-mouse CD4 Abs (eBiosciences) and sorting the CD4+CD25–CD45RBhigh or CD4+CD25+CD45RBlow cells via FACS (FACSadvantage; BD Biosciences). DCs (CD11c+, purity >80% purity, assessed by flow cytometry) were purified via a positive selection magnetic protocol (EasySep; Stemcell). CD11c, markers of costimulatory molecules CD80 and CD86 and the chemokine receptor CCR7, were stained with relevant fluorescent mAbs or isotype controls (eBiosciences). Staining was assessed via flow cytometry. All flow cytometric data was acquired on a FACSCalibur flow cytometer, and data were analyzed using FlowJo software (Treestar).

MLR, generation and activation of bone marrow-derived DCs (BMDCs), and proliferative assays

Allogeneic BMDCs were harvested as per our previously published work where bone marrow cells are harvested; depleted of T, B, and NK cells; and cultured in the presence of GM-CSF (15). To perform the MLR to examine the effect of MyD88 signaling within T cells, irradiated allogeneic (BALB/c) BMDCs (1 x 103 or 1 x 104/well) were added to 1 x 105 B6.MyD88+/+ or MyD88–/–CD4+CD25– T effectors or CD8+ T cells for 2 days in 96-well plates in complete Bruff’s medium (Invitrogen Life Technologies). At this point, the cells were resuspended in RPMI 1640, 10% FCS with 1% penicillin and streptomycin, transferred to ELISPOT plates specific for IL-2, and cultured for another 20 h; the plates were then analyzed as described above. To assess the effect of MyD88-dependent cytokines on T cell proliferation in an MLR and on T reg-mediated allosuppression, B6.MyD88+/+ and MyD88–/– BMDCs were stimulated with LPS (50 ng/ml) in complete Bruff’s medium. After 12 h the supernatants were harvested (denoted as stimulated medium) and used to culture syngeneic MyD88+/+ T effectors (1 x 105/well) that were allostimulated with irradiated BALB/c spleen cells (1:1 ratio). In certain experiments, 1 x 105 MyD88+/+ B6. T regs (CD4+CD25+) were then added to the wells. Proliferation was measured after 72 h of culture. During the last 12 h of culture, [3H]thymidine was added to the wells, and DNA was harvested and analyzed by a scintillation counter as an indicator of cell proliferation (PerkinElmer Life Science). Control wells included: stimulators or T effectors in rest medium (i.e., medium that was harvested from DCs that were not TLR activated where the same dose of LPS used to stimulate DCs was added after the culture supernatants were collected and before the MLR); T effectors cultured in stimulated medium without allogeneic spleen cells; and media only wells.

To perform the T reg-suppressive assays in response to CD3 activation, increasing numbers of CD4+CD25+ T regs (1 x 104, 1 x 105, 1 x 106) were added to 96-well plates containing 5 x 104 CD4+CD25– T effectors that were stimulated with anti-CD3, 0.3–3 µg/ml (Sigma-Aldrich) along with 5 x 104 irradiated, syngeneic, splenic feeder cells in complete medium. After 3 days, [3H]TdR was added to the wells to monitor cell proliferation as described above. Controls included T regs stimulated with anti-CD3, anti-CD3-stimulated effectors only and T effectors and T regs alone without stimulation, respectively.

Statistical analysis

Survival analysis between groups was calculated using the log rank method. Comparison of means was performed using a two-tailed t test. Repeated measures were assessed using a two-way ANOVA. All results were generated using GraphPad Prism software. Statistical significance was considered as p < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Absence of MyD88 synergizes with costimulatory receptor blockade and facilitates allograft acceptance

To test the hypothesis that an absence of MyD88 facilitates transplantation tolerance, we used a fully allogeneic murine skin allograft model that resists the tolerizing effects of costimulatory receptor blockade (specifically anti-CD154 and CTLA4-Ig, both of which inhibit signal 2 from APCs to T cells) (23) in wild-type recipients (24). Hence, we administered costimulatory receptor blockade to transplant groups that were either sufficient (wild type) or deficient in MyD88. In the absence of MyD88 (i.e., BALB/c MyD88–/– skin [H2d]->B6.MyD88–/– recipients [H2b]) the vast majority of transplant recipients (10 of 13) accepted their skin allografts indefinitely (>200 days), and graft rejection was significantly delayed when recipients are only MyD88 deficient (i.e., BALB/c MyD88+/+->B6.MyD88–/–) (Fig. 1A). In contrast, MyD88+/+ recipients that received MyD88+/+ skin allografts and the tolerance protocol rejected their allografts with a median survival time (MST) of 27 days (p < 0.0001 vs MyD88-deficient group).


Figure 1
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  		FIGURE 1. Absence of MyD88 facilitates graft acceptance mediated by costimulatory receptor blockade in a highly immunogenic, fully allogeneic skin transplant model. Skin allograft acceptance is induced in the absence of MyD88 and after perioperative treatment with costimulatory receptor blockade (anti-CD40L and CTLA 4 Ig) ({diamond}, MST >200 days) vs when MyD88 signaling is sufficient in both donor (BALB/c) and recipient (B6) ({square}, MST 27 days, p < 0.0001). Graft rejection is also prolonged by the tolerance protocol when only the recipient is deficient of MyD88 ({circ} MST >150 days, p = 0.01) but not when only the donor is deficient of MyD88–/– ({triangleup}) (MST = 18 days). Recipients that manifest functionally impaired IL-1 and IL-18 (caspase-1 deficient or ICE–/–; •) signaling remained resistant to the effects of costimulatory blockade (MST = 36 days p = 0.01 vs either MyD88–/– recipient group.

 
Consistent with a prior report, the absence of MyD88 did not delay graft rejection when the tolerance protocol was not administered (MST 12 days in MyD88-sufficient and -deficient groups, respectively) (15). Although untreated MyD88-deficient recipients mount an effective immune response similar to that exhibited by MyD88+/+ recipients (15), MyD88-deficient recipients are much more susceptible to tolerogenic effects of costimulatory blockade.

Although MyD88 is downstream of TLRs, it is also downstream of IL-1 and IL-18 (25). Hence, to determine whether graft acceptance in the MyD88–/– recipients was dependent on defective signaling via IL-1 or IL-18, we used caspase-1-deficient recipients that cannot signal via IL-1 and IL-18 (11, 26). The results show that these recipients remained resistant to the effects of costimulatory blockade (MST 36 days, p < 0.01 vs MyD88–/– recipient group) (Fig. 1). This suggests that graft acceptance in the MyD88–/– recipients was independent of the IL-1 and IL-18 signaling.

MyD88 signaling is important for DC inflammatory responses and migration after transplantation and treatment with costimulatory receptor blockade

Activation of DCs with TLR agonists causes them to produce proinflammatory cytokines, which are important for initiating adaptive immunity (26). This occurs principally via priming Th1 immune responses and inhibiting the effect of T reg cells (9, 26, 27, 28). Our prior studies have shown that MyD88 signaling within DCs is important for Th1 alloimmunity (11, 15); however, the impact of MyD88 signaling on DC production of proinflammatory cytokines during allotransplantation has not been previously documented (11, 15). We next examined whether splenic DCs produced proinflammatory cytokines after transplantation and whether these responses were altered in the absence of MyD88 and/or treatment with costimulatory blockade. Toward this end, we purified DCs from animals 7 days after transplantation with or without treatment with costimulatory blockade and tested their production of IL-6 and TNF-{alpha} via ELISPOT. DCs from MyD88+/+-transplanted recipients in the absence of costimulatory blockade demonstrated a substantial IL-6 and TNF-{alpha} production. Either administration of costimulatory blockade or absence of MyD88 alone significantly impaired DC proinflammatory cytokine production as compared with untreated MyD88+/+ transplant recipients (Fig. 2A). These responses were further impaired in MyD88–/– recipients treated with costimulatory blockade (Fig. 2A). This indicates that costimulatory blockade and an absence of MyD88 act synergistically to impair DC proinflammatory cytokine production after transplantation.


Figure 2
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  		FIGURE 2. Absence of MyD88 leads to defective DC proinflammatory cytokine and migratory responses after skin transplantation in the absence and presence of costimulatory (Costim) blockade. A, CD11c+ DCs were purified from the spleens of MyD88+/+ or MyD88–/– B6 recipients 7 days after receiving a BALB/c skin allograft in the presence or absence of costimulatory blockade. Cells were then analyzed for IL-6 or TNF-{alpha} production via ELISPOT. Results demonstrate that DCs isolated from MyD88+/+ untreated transplant recipients produced substantial levels of IL-6 and TNF-{alpha}. Either absence of MyD88 or treatment with costimulatory blockade reduced DC proinflammatory cytokine responses as compared with untreated transplanted MyD88+/+ recipients. The effect was further inhibited in MyD88–/– recipients treated with costimulatory blockade. The results shown are corrected for baseline cytokine secretion produced by nontransplanted mice from the respective groups. Data for naive mice are: IL-6: MyD88+/+ 58 spots/3 x 105 DCs; MyD88–/– 57 spots/3 x 105; TNF-{alpha}: MyD88+/+ 15 spots/3 x 105, MyD88–/– 1 spot/3 x 105. Data are representative of three independent experiments with three mice per group per experiment. B, Draining lymph nodes (inguinal and axillary) were harvested from MyD88+/+ or MyD88–/– recipients after transplantation and treatment with costimulatory blockade. The lymph nodes were subsequently analyzed via flow cytometry for CD11c and CCR7 staining. Gating on the CD11c+ population, lymph nodes from either naive, untransplanted MyD88+/+ or MyD88–/– mice manifest similar expression of CCR7. DCs from MyD88+/+ untreated transplant recipients up-regulated CCR7 as compared with naive controls. This response was impaired in CD11c+ DCs from MyD88–/– transplant recipients in the presence or absence of costimulatory blockade. Black thick line, MyD88+/+; thin gray line, MyD88–/–; thin black line, isotype control. Results are from two independent experiments with three mice per group per experiment. C, The absolute number of mature DCs (defined as either CD11c+CD80high or CD11c+CD86high) was quantified via flow cytometry in the draining lymph nodes of allografts at 1-wk after transplantation after treatment with costimulatory blockade. The absolute number of mature DCs was calculated as the product of the total number of lymphocytes times the proportion of CD11c+CD80high or -86high as determined by flow cytometry. The results demonstrate that in MyD88+/+ transplant recipients treated with costimulatory blockade mature DCs accumulate in the draining lymph node whereas this accumulation is impaired in MyD88–/– counterparts. In agreement with prior reports (11 15 ), similar findings were obtained in the absence of immune modulation (data not shown).

 
In our prior work, we found that in MyD88+/+-transplanted recipients that did not receive costimulatory blockade, mature DCs (defined as either CD11c+CD80high or -86high) accumulate in the draining lymph nodes of allografts, whereas this effect was compromised in MyD88–/– recipients (15). These effects persisted in the presence of costimulatory blockade and were associated with an impaired ability of MyD88–/– DCs to up-regulate the chemokine homing receptor CCR7 (Fig. 2, B and C).

Absence of MyD88 leads to reduced CD4+ and CD8+ T cell priming after transplantation and treatment with costimulatory receptor blockade

Because MyD88 signaling was important for DC inflammatory responses during transplantation and because DC inflammatory responses are important for T cell priming and generation of T effectors (6, 29), we next examined whether MyD88 signaling altered CD4+ and CD8+ T cell priming after transplantation. Hence, we measured the number of both CD4+ and CD8+ IL-2- and IFN-{gamma}-producing T cells either in the absence of immune modulation or after administration of costimulatory blockade. Either administration of costimulatory blockade or MyD88 deficiency alone led to impaired allospecific CD4+ and CD8+ T cell responses (Fig. 3, A and B), with the possible exception of the IL-2 response of wild-type recipients treated with costimulatory blockade (Fig. 3B). The combination of administration of costimulatory blockade and MyD88 deficiency manifest the most defective response indicating synergy between these two factors (Fig. 3, A and B). Finally, we found that defective cytokine responses within unsorted spleen cells persisted in MyD88-deficient recipients that had accepted their allografts for >150 days post transplantation (denoted as long term acceptors) as measured via ELISPOT (Fig. 3C). However, spleen cells from these recipients were not completely anergic and were able to produce a response to nonspecific stimulation with Con A (Fig. 3C).


Figure 3
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  		FIGURE 3. Absence of MyD88 impairs T cell priming after transplantation and treatment with costimulatory receptor blockade. MyD88-sufficient groups or deficient groups were transplanted and were either treated with costimulatory receptor blockade or left untreated. Spleen cells from these groups were harvested at 2 wk posttransplantation (A and B) or 150 days posttransplantation (C) and sorted via negative magnetic selection for CD4+ or CD8+ T cells (or left unsorted) as described in Materials and Methods. These cells were then stimulated ex vivo with donor irradiated spleen cells and analyzed via ELISPOT. BALB/c. MyD88–/– spleen cells were used as stimulators for B6.MyD88–/– recipients that received BALB/c.MyD88–/– skin grafts, although similar results were obtained when MyD88–/– T cells were stimulated by irradiated MyD88+/+ donor spleen cells and vice versa (data not shown). In agreement with our prior work (15 ), naive responders did not elicit a response to donor-specific spleen cells and responders were nonresponsive when stimulated with irradiated donor third party (CBA) spleen cells (data not shown). A, Treatment with costimulatory blockade or absence of MyD88 reduces the number of IFN-{gamma} CD4+- and CD8+-producing T cells. This effect is reduced further in MyD88–/– recipients treated with costimulatory blockade. B, Similar results were found with IL-2. C, Unsorted spleen cells were harvested from long term (>150 days posttransplant) B6.MyD88–/– acceptors and analyzed for responsiveness to irradiated donor stimulator spleen cells via ELISPOT. The data demonstrate that long term acceptors were hyporesponsive for all cytokines tested (IL-2, IL-6, TNF-{alpha}, and IFN-{gamma}; <10 spots/1 x 106 spleen cells) yet were able to respond to nonspecific T cell activation with Con A (>1500 spots/1 x 106 spleen cells). Spot counts for each cytokine appear above each histogram bar. (MyD88+/+ recipients of MyD88+/+ allografts that were treated with costimulatory receptor blockade demonstrated a detectable response at 150 days after transplantation: IFN-{gamma}, 836 ± 33 spots/1 x 106 cells; IL-2, 546 ± 25 spots/1 x 106 cells.)

 
No evidence of impaired in vitro intrinsic T cell defects in the absence of MyD88

The results in the previous paragraph could be explained either by defective MyD88–/– DC responses leading to reduced T cell priming or an intrinsic defect in MyD88–/– T cells. To examine whether absence of MyD88 impaired intrinsic T cell function, MyD88+/+ or MyD88–/– CD4+ or CD8+ T cells were stimulated with allogeneic spleen cells or activated nonspecifically with CD3 stimulation. The results indicate that MyD88–/– T cells functioned equally as well as MyD88+/+ counterparts in these assays (Fig. 4, A and B). Hence, the data indicate that absence of MyD88 does not intrinsically impair T cell function.


Figure 4
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  		FIGURE 4. No evidence of intrinsic T cell defects in MyD88–/– T cells vs. MyD88+/+ counterparts. A, Preserved CD4+ and CD8+ responses in naive B6.MyD88–/– T cells vs. MyD88+/+ counterparts in response to allogeneic stimulation. B6.CD4+CD25– T cells or CD8+ T cells were purified (as described in Materials and Methods) from naive MyD88+/+ or MyD88–/– mice and stimulated with irradiated BALB/c BMDCs. The numbers of IL-2-producing T cells were measured by ELISPOT. No significant differences were noted between MyD88+/+ and MyD88–/– groups at the different DC doses. Unstimulated T cells did not produce a response (data not shown). B, Preserved CD4+ and CD8+ T cell responses after activation with plate-bound CD3. MyD88+/+ or MyD88–/– CD4+ and CD8+ T cells were purified and then activated with increasing doses of CD3. Results were obtained by [3H]TdR incorporation.

 
Synergy between an absence of MyD88 and treatment with costimulatory blockade is donor specific

To determine whether the synergy between treatment with costimulatory receptor blockade and an absence of MyD88 led to donor-specific graft acceptance, long term B6.MyD88–/– recipients that had accepted their BALB/c MyD88–/– allografts for >150 days (denoted as acceptors) were rechallenged with either donor-specific (MyD88–/–) BALB/c skin allografts or MyD88+/+, third party skin allografts (CBA, H2k). The results demonstrate that donor-specific rechallenge allografts exhibited markedly prolonged survival (MST >100 days), whereas the third party allografts were rapidly rejected (MST 14 days, p = 0.003; Fig. 5). Similar results were obtained when rechallenge grafts were from MyD88-sufficient donors (i.e., BALB/c MyD88+/+ rechallenge grafts, MST >30 days, p = 0.004 vs CBA, p = 0.1 vs MyD88–/– donor rechallenge grafts; Fig. 5). These data demonstrate that the acceptance of allografts mediated by costimulatory receptor blockade in the absence of MyD88 is donor specific.


Figure 5
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  		FIGURE 5. Graft acceptance after treatment with costimulatory blockade in the absence of MyD88 is donor specific. B6.MyD88–/– long term acceptors of mutant BALB/c allografts were rechallenged with a MyD88–/– donor-specific (i.e., BALB/c) skin allograft or a third party, MyD88+/+ (CBA) allograft. Results demonstrate that third-party allografts ({square}, MST 14 days) were rejected in an accelerated fashion as compared with donor-specific MyD88–/– rechallenge grafts ({circ}, MST >100 days, p = 0.003). Mice rechallenged with MyD88+/+ BALB/c skin grafts ({blacktriangledown}) also manifested extended allograft survival vs third-party grafts, (i.e., BALB/c. MyD88+/+ rechallenge grafts, MST >30 days, p = 0.004 vs CBA, p = 0.1 vs MyD88–/– donor rechallenge grafts).

 
Synergy between an absence of MyD88 and treatment with costimulatory blockade that leads to graft acceptance occurs via a CD25-dependent regulatory mechanism

Because long term MyD88–/– acceptors demonstrated delayed rejection of donor-specific challenge grafts at a point when the effects of costimulatory blockade have likely dissipated, this suggests that immune regulation is one of the mechanisms of graft acceptance in these recipients. Given that CD4+CD25+ T regs are important for tolerance induction (30), we asked whether long term allograft acceptance induced by costimulatory receptor blockade in the absence of MyD88 involves a regulatory mechanism (30). B6.MyD88–/– mice were treated with either an inhibiting anti-CD25 Ab, which has been recently shown to functionally inactivate CD4+CD25+ T regs (31), or with an isotype control before transplantation with BALB/c.MyD88–/– skin and administration of costimulatory receptor blockade. The results demonstrate that anti-CD25 treatment abrogates graft acceptance in MyD88–/–-transplanted recipients and recovers alloimmunity (Fig. 6, A and B), indicating that CD4+CD25+ T regs are important for graft acceptance in these recipients. Experiments in MyD88+/+ recipients that received costimulatory blockade demonstrated that rejection kinetics was not altered by CD25 inhibition (anti-CD25-treated MyD88+/+ group MST, 22 days, n = 4, p = 0.2 vs MyD88+/+ isotype-treated group). Additionally, we were able to transfer graft acceptance by isolating purified CD4+CD25+ T regs from long term B6.MyD88–/– acceptors and adoptively cotransferring the T regs with MyD88+/+CD4+CD25– T effectors into B6.RAG–/– recipients that were previously transplanted with a MyD88+/+ BALB/c skin allograft (Fig. 6C).


Figure 6
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  		FIGURE 6. Synergy between costimlatory receptor blockade and absence of MyD88 leading to allograft acceptance occurs via a CD25-regulatory mechanism. A, Anti-CD25 treatment ({square}) before transplantation abrogates BALB/c.MyD88–/– graft acceptance in B6.MyD88–/– recipients treated with costimulatory receptor blockade (p < 0.01 vs isotype control-treated group {diamond}). B, Anti-CD25 treatment before skin transplantation and treatment with costimulatory receptor blockade in B6.MyD88–/– recipients ({blacksquare}) led to recovery of T cell alloimmune responses as compared with B6.MyD88–/– recipients treated with isotype control ({square}) (p = 0.03) at 2 wk after transplantation as measured by ELISPOT. Data are representative of two independent experiments. C, CD4+CD25+ T cells purified from B6.MyD88–/– who were long term acceptors (i.e., 150 days after transplantation) of BALB/c.MyD88–/– allografts induced graft acceptance when adoptively cotransferred with CD4+CD25–MyD88+/+ T effectors (Teff) into B6.RAG–/– recipients that were transplanted previously with a MyD88+/+ BALB/c allograft. CD4+CD25+ T regs (2 x 105) were cotransferred with 1 x 105 CD4+CD24– T effectors via i.v. tail vein injection. This impaired the ability of T effectors to induce graft rejection ({blacktriangleup}, MST >180 days) vs transplanted B6.RAG–/– recipients that were transferred with T effectors only ({blacksquare}, MST 22 days, p < 0.01). Coinfusion of 2 x 105 CD4+CD25–MyD88–/– T effectors and 1 x 105 MyD88+/+ T effectors (•) did not delay the ability of MyD88+/+ T effectors to induce graft rejection (MST 19 days, p = 0.3 vs infusion of MyD88+/+ T effectors alone).

 
Absence of MyD88 does not intrinsically augment T reg function

To examine whether MyD88 deficiency itself leads to augmented suppressive CD4+CD25+ T reg function, we measured the suppressive ability of either MyD88+/+ or MyD88–/– T regs both in vitro and in vivo. Our results show that MyD88–/– T regs do not possess superior ability to suppress in vitro T cell activation (Fig. 7A) or in vivo alloimmunity (Fig. 7B) as compared with MyD88+/+ counterparts. Furthermore, we did not find evidence that the absence of MyD88 leads to increased numbers of splenic Foxp3+ cells in naive mice (FoxP3 is a transcription factor found in naturally occurring CD4+CD25+ T regs (30)) (Fig. 7, C and D). Nor did we find evidence of increased numbers of CD4+CD25+ T cells within MyD88–/– host spleen cells either pre- or posttransplantation as compared with MyD88+/+ counterparts (data not shown). These findings indicate that MyD88 deficiency does not intrinsically enhance CD4+CD25+ T reg-suppressive function or the generation of T regs.


Figure 7
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  		FIGURE 7. MyD88 deficiency does not enhance the intrinsic suppressive ability of T regs. A, CD4+CD25+ T regs were FACs sorted from naive B6.MyD88–/– or B6.My88+/+ mice, and their ability to suppress T effector proliferation was measured by thymidine incorporation assay. B6.MyD88–/– T regs ({diamond}) manifest equal suppressive in vitro function vs MyD88+/+ CD4+CD25+ T regs ({square}) on nonspecifically activated (via CD3) CD4+CD25– T effectors. B, Survival curve showing that CD4+CD25+ T regs purified from MyD88–/– recipients transplanted with a BALB/c.MyD88–/– allograft (150 days after transplantation) and treated with costimulatory receptor blockade manifest a similar ability to impair CD4+CD25– T effectors in inducing allograft rejection as compared with MyD88+/+ CD4+CD25+ T regs purified from similarly treated MyD88-sufficient transplant recipients. T regs (2 x 105) purified from either MyD88–/– ({blacktriangleup}) or MyD88+/+ recipients ({diamondsuit}) were adoptively cotransferred, via i.v. tail vein injection, with 1 x 105 MyD88+/+CD4+CD25– T effectors into a B6.RAG–/– recipient previously transplanted with a MyD88+/+ BALB/c skin allograft. Both p < 0.01 vs survival in recipients infused with T effectors alone ({blacksquare}). C, Flow cytometric dot plots showing equal expression of Foxp3 in MyD88–/– vs MyD88+/+ spleen cells. Proportions are shown in lower right quadrant. D, Histogram plot showing equal expression of Foxp3 within the CD4+CD25+ subpopulation in naive MyD88–/– and MyD88+/+ hosts. Blue, MyD88–/–; red, MyD88+/+; green, isotype control. Histograms are gated on the CD4+CD25+ subpopulation. Flow cytometric dot plots of CD4+ and CD25+ cells within the Foxp3+ subpopulation in naive MyD88–/– and MyD88+/+ mice. (Spleen cells from naive MyD88–/– and MyD88+/+ mice were of similar size and cellularity; data not shown.). WT, wild type; FL2-H, fluorescence.

 
MyD88-dependent proinflammatory cytokines from TLR-activated DCs augment T cell proliferation during allogeneic stimulation

Prior reports indicate that proinflammatory cytokines are important for recovery of T cell function in immunosenescent cells and promoting autoimmunity (32, 33). We next examined whether proinflammatory cytokines from TLR activated (via LPS) DCs augmented the ability of MyD88+/+ T effectors to respond to allogeneic stimulation. To examine the role of MyD88 signaling within TLR-activated DCs, we harvested culture media from either B6.MyD88+/+ or MyD88–/– BMDCs after LPS stimulation. In agreement with prior studies (34), proinflammatory cytokines IL-6 and TNF-{alpha}, present in MyD88+/+ supernatants, were abrogated in supernatants from MyD88–/– DCs (data not shown). These media were then used to culture syngeneic T effector cells that were stimulated with allogeneic BALB/c spleen cells. The results demonstrated that culture media harvested from LPS-stimulated MyD88+/+ DCs augmented proliferation as compared with T effectors allostimulated in the presence of media harvested from LPS stimulated MyD88–/– BMDCs (Fig. 8A). Media from this latter group did not augment T cell proliferation above control levels generated from media harvested from representative non-TLR-activated DCs (Fig. 8A). We also found that media harvested from LPS-stimulated MyD88+/+ DCs augmented T cell proliferation in response to allostimulation in a dose-dependent manner and were significantly superior to media harvested from LPS-stimulated MyD88–/– BMDCs (Fig. 8B).


Figure 8
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  		FIGURE 8. Proinflammatory cytokines produced by TLR-stimulated DCs augment T cell proliferation during allogeneic stimulation and impair suppression of alloimmunity mediated by T regs. Culture supernatants were harvested from either TLR-activated (via LPS) B6.MyD88+/+ (denoted as MyD88+/+ media) or MyD88–/– (denoted as MyD88–/– media) BMDCs and used to culture 1 x 105 MyD88+/+ CD4+CD25– T effectors that were allostimulated with 1 x 105 BALB/c spleen cells. Media harvested from non-TLR-activated MyD88+/+ DCs (denoted as rest media) was supplemented with LPS after DC removal. Proliferation was measured after 72 h of culture via [3H]TdR incorporation. Each figure (A–E) is derived from separate independent experiments. A, Proliferation of MyD88+/+ T effectors was augmented in the presence of media harvested from TLR activated MyD88+/+ BMDCs as compared with media harvested from TLR-activated MyD88–/– DCs or control rest media. All media used in this experiment was diluted 1/2 with fresh media. B, Dose response effect of media harvested from TLR activated MyD88+/+ BMDCs or MyD88–/– BMDCs. Figure shows that MyD88+/+ allostimulated T effectors manifest increased proliferation in the presence of media harvested from MyD88+/+ BMDCs vs MyD88–/– BMDCs across the range of dilutions. C, Recombinant IL-6 supplementation (38 ng/ml) of media harvested from TLR-activated MyD88–/– BMDCs significantly augmented T effector proliferation during allostimulation as compared with nonsupplemented media harvested from TLR-activated MyD88–/– BMDCs. D, IL-6 mAb inhibition (2 µg/ml) significantly reduced T effector proliferation during allostimulation when added to media harvested from TLR-activated MyD88+/+ BMDCs as compared with unmodified media harvested from TLR-activated MyD88+/+ BMDCs or media treated with isotype control. E, MyD88+/+ B6.CD4+CD25+ T regs were added to the wells at a 1:1 ratio with T effectors. In the absence of T regs, media harvested from MyD88+/+ TLR-activated DCs (MyD88+/+ media) increased T effector proliferation significantly compared with media from MyD88–/– TLR-activated DCs (p = 0.01). The ability of T regs to suppress T effector proliferation during allostimulation was impaired in the presence of MyD88+/+ media (*, 40% suppression) as compared with MyD88–/– media (**, 80% suppression, p = 0.005).

 
Given that prior studies have identified IL-6 as an important cytokine for T cell activation and the induction of autoimmunity (33, 35, 36), we wished to determine whether IL-6 was an active component of the media harvested from TLR-activated MyD88+/+ BMDCs, which augmented T cell proliferation during allostimulation. Hence, we performed experiments in which we supplemented media from MyD88–/– TLR-activated DCs with rIL-6. The dose of IL-6 used was determined by the difference in IL-6 concentration (38 ng/ml) measured by ELISA in the culture media produced by TLR-activated MyD88+/+ and MyD88–/– DCs, respectively. The results show that the addition of IL-6 to the MyD88–/– media significantly augmented the ability of T effectors to proliferate in response to allostimulation (Fig. 8C). Next, we incubated T effectors in media harvested from TLR-activated MyD88+/+ BMDCs and added an IL-6-inhibitory Ab during allostimulation. The results of this experiment demonstrate that IL-6 inhibition reduced T effector proliferation (Fig. 8D). Hence, the above data indicate factors produced by TLR-activated DCs augment T cell proliferation in vitro; this augmentation is MyD88-dependent and appears to involve IL-6 as one of the active components.

Media harvested from TLR-activated MyD88+/+ DCs inhibit T reg-mediated suppression of alloimmunity

A prior study has found that MyD88-dependent cytokines produced from DCs modify T effector cell susceptibility to suppression mediated by T regs during nonspecific T cell activation (9). Given that our data provide evidence that MyD88 signaling is important for DC production of inflammatory cytokines after transplantation and that these MyD88-dependent cytokines augment T cell proliferation during allostimulation, we next examined whether these cytokines altered the ability of T regs to suppress T effector proliferation in response to allogeneic stimulation with BALB/c spleen cells. Hence, MyD88+/+ B6.CD4+ CD25+ T regs were added to cultures of B6.CD4+CD25– T effectors, which were then allostimulated with BALB/c spleen cells in either culture supernatants derived from TLR-activated MyD88-sufficient or -deficient DCs. The results demonstrate that in the presence of media from MyD88+/+ TLR-activated DCs, wild-type T regs manifest an inferior ability to inhibit proliferation of syngeneic MyD88+/+ T effector cells in response to allostimulation. In contrast, T regs manifest a superior ability to suppress T effector function when cultured in the presence of media harvested from MyD88–/– DCs (Fig. 8E). These data demonstrate that MyD88-dependent cytokines prevent T reg-mediated suppression of allogeneic stimulation in vitro.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
In our study, we provide evidence that absence of MyD88 synergizes with costimulatory receptor blockade and induces long term transplant acceptance in a highly immunogenic skin allograft model (37). We found that lack of MyD88 signaling in our model impairs DC-inflammatory responses and T cell priming (Figs. 2 and 3). We also show that donor-specific rechallenge allografts demonstrate a marked prolonged survival in MyD88–/– recipients that had accepted their primary allograft for >150 days as compared with third party allografts (Fig. 5). Because immunoregulation is a possible explanation for this result, we examined the role of T regs in our model. Our results indicate that graft acceptance induced by costimulatory receptor blockade in the absence of MyD88 was dependent on CD4+CD25+ T regs since graft acceptance was transferable after adoptive transfer of MyD88-deficient CD4+CD25+ T regs (Fig. 6C). Furthermore, functional CD25 inhibition (31) abrogated graft acceptance (Fig. 6A) and recovered alloimmunity (Fig. 6B) in the MyD88-deficient recipients. This result implies that in our model, graft acceptance mediated by costimulatory receptor blockade requires the presence of T regs.

We found that an absence of MyD88 enhanced neither the generation nor the intrinsic function of T regs (Fig. 7). We provide a possible explanation to this issue by demonstrating that absence of MyD88 impairs DC inflammatory responses after transplantation (Fig. 2). We present evidence that the functional consequences of this impaired inflammatory response lead to reduced in vitro T effector proliferation during allostimulation (Fig. 8, A and B). We also show that MyD88-dependent cytokines produced by TLR-activated DCs impair T reg mediated suppression of allostimulated T effectors (Fig. 8E). This effect occurred without altering the nature or number of T regs in this assay. We noted that a minority of MyD88–/– recipients that had accepted their allografts for >150 days subsequently rejected rechallenge donor-specific grafts. A possible explanation for this is that the effects of costimulatory blockade may have dissipated at this point and therefore the ischemia reperfusion injury upon rechallenge may have perturbed the cytokine environment and tipped the balance from tolerance to immunity. Overall, our data suggest that absence of MyD88 alters the inflammatory cytokine environment, allowing T effectors to proliferate less and be more easily suppressed by T regs. This occurs without changing the intrinsic function or generation of T regs.

Our study is compatible with a previous in vitro report that demonstrated that IL-6 produced by TLR-activated DCs allowed effector T cells to become resistant to the suppressive effects of T regs, although the authors of this study found that this phenomenon could not be entirely explained by IL-6 alone (9). We show that in an alloimmune setting IL-6 is an important factor in augmenting T cell proliferation, although our data do not exclude that other inflammatory cytokines may also be involved in this effect. Future in vivo studies are required to examine the role of specific proinflammatory cytokines in transplantation tolerance induction, although it is possible that there may be a high degree of redundancy between cytokines. This is supported by studies conducted several years ago that documented redundancy in cytokines involved in mediating acute allograft rejection (17, 38).

Although our study has identified CD4+CD25+ T regs as an important mediator of graft acceptance in our model, other mechanisms such as clonal deletion, anergy, and ignorance may also play a role. Future studies are required to address these issues. Lastly, there are many other innate immune receptors and pathways that are TLR independent (39, 40). Future studies will be required to investigate the impact of these other innate immune pathways on transplantation tolerance induction.

It has been known for decades that skin allografts are a far more stringent barrier for graft acceptance as opposed to other organs (3, 19, 20). Our data suggest that increased activation of the innate immune system by skin allografts may be one of the explanations for this finding given that other studies have found that vascularized and islet allografts are more susceptible to tolerance induction (3, 19, 20, 41). The ligands that activate innate immunity during solid organ transplantation are yet to be elucidated. Possibilities include substances that are endogenous derived, exogenous to the host or synergy between multiple activators. The role of exogenous substances in alloimmunity remains unclear because long term, accepted allografts without signs of inflammation can also undergo rejection (37, 42). However, a recent study has provided evidence that exogenous innate ligands given systemically may abrogate tolerance induction (43). We used LPS in our study as a practical experimental tool to activate TLRs on DCs and produce sufficient quantities of cytokines. LPS is a well established activator of the MyD88 pathway in nontransplant settings (34). Clearly, future studies are needed to determine the nature and identity of the activators of DCs during the response to transplantation.

The potential clinical implications of our study are that transiently inhibiting innate immune pathways may provide targets to promote the efficacy of transplant tolerance protocols. This is potentially important given the rising appreciation of innate immunity in ischemia reperfusion injury (12, 13, 14). The other implication of our study is that an acute inflammatory response, for example an infection, which can augment innate immune signaling, may interfere with the efficacy of transplant tolerance protocols, leading to a break of tolerance and initiation of immunity. Indeed, there are prior reports that heterologous immunity to viral Ags prevents transplantation tolerance (44). This is supported by a recent study that demonstrated that lymphocytic choriomeningitis viral infection, which likely activates many components of innate immunity, impaired the ability of an allograft prolonging treatment consisting of donor cell transfusion and anti-CD154 to extend allograft survival (43).

In conclusion, our study provides proof-of-principle evidence that inhibiting specific innate immune signaling pathways may synergize with therapies that have the potential to induce transplantation tolerance. This information may be relevant to future clinical strategies aimed at inducing transplantation tolerance.


   Acknowledgments
 
We thank Fred Kantor and David Rothstein (both of Yale University) for their careful reading of the manuscript.


   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 was supported by National Institutes of Health Grant AI064660 (to D.R.G.). Back

2 Contributed equally to this work. Back

3 Address correspondence and reprint requests to Dr. Daniel R. Goldstein, 333 Cedar Street, 3 FMP, P.O. Box 208017, New Haven, CT 06520-8018. E-mail address: daniel.goldstein{at}yale.edu Back

4 Abbreviations used in this paper: DC, dendritic cell; SPC, spleen cell; T reg, regulatory T cell; BMDC, bone marrow-derived DC; MST, median survival time. Back

Received for publication February 6, 2006. Accepted for publication July 31, 2006.


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J. Immunol., August 1, 2008; 181(3): 1849 - 1858.
[Abstract] [Full Text] [PDF]

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A. C. Shirali and D. R. Goldstein
Tracking the Toll of Kidney Disease
J. Am. Soc. Nephrol., August 1, 2008; 19(8): 1444 - 1450.
[Abstract] [Full Text] [PDF]

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