In Vivo Maturation of Allo-Specific CD8 CTL and Prevention of Lupus-like Graft-versus-Host Disease Is Critically Dependent on T Cell Signaling through the TNF p75 Receptor But Not the TNF p55 Receptor

Kateryna Soloviova, Maksym Puliaiev, Mark Haas and Charles S. Via
J Immunol May 1, 2013, 190 (9) 4562-4572; DOI: https://doi.org/10.4049/jimmunol.1300091

Abstract

A third signal is required for maturation of effector CD8 CTL in addition to TCR and CD28 engagement. Inflammatory cytokines can provide a third signal; however, in nonpathogen settings (i.e., antitumor responses), the identity of the third signal is not clear. A useful model for in vivo CD8 CTL in the absence of exogenous pathogens is the alloantigen-driven parent-into F1 model of acute graft-versus-host disease (GVHD) characterized by a strong TNF-dependent donor antihost CD8 CTL T cell response. To determine whether TNF acts directly on donor T cells in a signal 3 manner, F1 mice received TNFR 1 (p55) knockout (KO) and/or TNFR 2 (p75) KO donor T cells. Donor p75 KO but not p55KO donor T cells failed to induce acute GVHD phenotype and instead induced a lupus-like chronic GVHD both short and long term because of quantitative and qualitative donor T cell defects, that is, reduced perforin, IFN-γ, and TNF production. Transfer of mixed or matched purified CD4 and CD8 T cells from wild type or p75KO donors demonstrated that optimal CTL maturation required p75 signaling in both CD4 and CD8 T cells. Despite defective p75KO CD4 help for CD8 CTL, p75KO CD4 help for B cells and autoimmunity was intact. These results provide a mechanism by which impaired CD8 CTL could contribute to reduced antiviral and antitumor responses and autoimmunity reported in patients receiving TNF blockers. Our results support the idea that selective p55 blockade may be beneficial by reducing inflammation without compromising CD8 CTL.

Introduction

In addition to TcR- and CD28-mediated signals, CD8 T cells require a third signal to mature into effector CTL and avoid apoptosis (1, 2). In pathogen models, IL-12 and IFN-α have been shown to act in a signal 3 manner and directly promote CD8 CTL effector function; however, the relative importance of either molecule varies with the pathogen (3). CD8 CTL play an important role in elimination of certain pathogens, typically accompanied by activation of the innate immune system and endogenous IL-12 or IFN-α. However, CD8 CTL arising in the nonpathogen setting (i.e., antitumor responses) likely occur in the absence of IL-12 and IFN-α, and may therefore use other molecules as a third signal.

A useful model for generating in vivo CD8 CTL in the absence of pathogen infection is the parent-into-F1 (p→F1) model in which homozygous parental strain T cells are transferred into normal nonirradiated F1 mice (reviewed in Refs. 4, 5). For example, the transfer of C57BL/6 (B6) splenocytes into B6D2F1 (BDF1) hosts (B6→BDF1) initially results in donor CD4 T cell recognition of F1 host allogeneic MHC class II and the provision of help to: 1) F1 B cells that then undergo polyclonal activation and expansion over the next 7 d; and 2) donor (B6) CD8 T cells that recognize allogeneic host MHC class I, mature into CTL effectors, and reverse host splenic lymphoproliferation by eliminating host splenocytes from days 7–14 after transfer (5).

Using this model, we have previously demonstrated that in vivo TNF blockade during the first week after donor cell transfer completely abrogated donor antihost CD8 CTL maturation and converted 2-wk phenotype from acute to chronic graft-versus-host disease (GVHD) (6). TNF blockade after day 7 did not alter acute GVHD phenotype (6), supporting the idea that TNF has a role in donor CD8 CTL maturation, possibly as a signal 3 function, separate from its well-known effector role.

TNF signaling in T cells results from TNF binding to one of two receptors: 1) TNFR1 or p55 expressed on nearly all cells and primarily associated with the inflammatory and apoptotic activity of TNF; and 2) TNFR2 or p75 whose expression is restricted to a subpopulation of immune cells and other cells, and is involved in T cell survival and costimulation (7–9). To determine whether the requirement for TNF in CD8 CTL maturation in p→F1 acute GVHD mice requires TNF signaling directly on donor T cells consistent with a signal 3 role (as opposed to acting indirectly through host APCs), we used the B6→BDF1 model of anti-F1 CD8 CTL generation and transferred donor T cells deficient in TNFR p75 and/or p55 into wild type (WT; TNFR intact) BDF1 hosts. Our results indicate a critical role for p75 signaling on donor CD4 and CD8 T cells for optimal CD8 effector CTL maturation.

Materials and Methods

Mice

Six- to 8-wk-old male WT B6 (H-2b), TNFR p55–deficient B6.Tnfrsf1atm1Imx/J (p55KO), p75-deficient B6.Tnfrsf1btm1Imx/J (p75KO) or double-deficient B6.Tnfrsf1atm1ImxTnfrsf1btm1Imx/J (p55/75KO) (all H-2b), and BDF1 (H-2b/d) mice were purchased from The Jackson Laboratory (Bar Harbor, ME). All animal procedures were preapproved by the Institutional Animal Care and Use Committee at the Uniformed Services University of Health Sciences.

Induction of GVHD

Single-cell suspensions of B6 WT or TNFR-deficient donor splenocytes were prepared as described previously (10) and transferred into age-matched BDF1 hosts by tail vein injection. The percentages of donor CD4 and CD8 T cells populations were analyzed by flow cytometry and donor inoculum adjusted before transfer to achieve the desired amount of donor CD4 and CD8 T cells as indicated in the text and respective figure legends. In some experiments, acute GVHD was induced using purified donor T cell subsets achieved through negative selection using Dynal mouse CD4 or CD8 negative isolation kits (Invitrogen, Carlsbad, CA), which deplete B cells, NK cells, monocytes/macrophages, dendritic cells (DCs), granulocytes, platelets, erythrocytes, and either CD8 or CD4, respectively. Purity was confirmed by flow cytometry and was typically >95%.

Flow cytometric analysis

Spleen cells were first incubated with anti-murine FcγRII/III mAb, 2.4G2 for 10 min, and then stained with saturating concentrations of Alexa Fluor 488–conjugated, allophycocyanin-conjugated, biotin-conjugated, PE-conjugated, FITC-conjugated, PerCPCy5.5-conjugated, and Pacific Blue-conjugated mAb against CD4, CD8, B220, H-2Kdd, I-Ad, CD11b, and CD11c purchased from either BD Biosciences (San Jose, CA), BioLegend (San Diego, CA), eBioscience (San Diego, CA), or Invitrogen (Carlsbad, CA). Biotinylated primary mAbs were detected using PE-Texas Red-streptavidin (BD Biosciences). Cells were fixed in 1% paraformaldehyde before reading.

Ex vivo intracellular staining for perforin, IFN-γ, and TNF was performed using Abs and reagents purchased from BD Pharmingen (San Diego, CA) or BioLegend, and staining performed according to the manufacturer’s instructions. Importantly, there was no in vitro restimulation or use of Golgi blocking agents. After completion of the staining protocol, cells were analyzed by flow cytometry immediately.

Multicolor flow cytometric analyses were performed using a BD LSRII flow cytometer (BD Biosciences). Gating strategy was as follows: lymphocytes were gated by forward and side scatter, and fluorescence data were collected for a minimum of 10,000 gated cells. Studies of donor T cells were performed on a minimum of 5000 cells collected using a lymphocyte gate that was positive for CD4 or CD8 and negative for MHC class I of the uninjected parent (H-2Kd−). B cells were gated as positive for B220 and either positive (host origin) or negative (donor origin) for MHC class II of the uninjected parent (I-Ad). Short-lived effector CD8 CTL were assessed as KRLG-1+, CCR7− gated donor CD8 T cells. Host DCs and macrophages were identified as I-Ad+ and CD11c+ or CD11b+, respectively, using a broad forward and side scatter gate.

Cytokine expression by real-time PCR

Real-time PCR was performed as described previously (11). In brief, splenocytes (1 × 107) were homogenized in 1 ml RNA-STAT-60 (Tel-Test, Friendswood, TX). cDNA was synthesized from mRNA using TaqMan Reverse Transcription Reagents kit (Applied Biosystems, Foster City, CA). Real-time PCR was performed using premade primers and probes from TaqMan Gene Expression Assays and TaqMan Universal PCR Master Mix (Applied Biosystems) for the following targets: IFN-γ, IL-10, IFN-γ, oligoadenylate synthetase-1 (OAS), and inducible protein-10 (IP-10) with 18s rRNA as an internal control. The calculation of relative gene expression differences was done by comparative 2−ΔΔCT method. The result was expressed as fold change in the experimental groups compared with uninjected BDF1 control. Mice were tested individually, and values are shown as group mean ± SE.

Kidney histopathology

Kidney tissue was fixed in 10% buffered formalin and processed for routine paraffin embedding and histological sectioning. Three-micrometer-thick sections, stained with periodic acid–Schiff stains, were blindly scored by a renal pathologist (M.H.). The severity of proliferative glomerulonephritis was determined according to a previously described semiquantitative (1+ to 4+) scoring system developed for a murine model of lupus nephritis (12, 13).

Serological studies

Mice were bled at the times indicated, and sera were tested by ELISA for the presence of IgG Abs to ssDNA as described previously (11).

Statistical analysis

Statistical comparisons (t test and ANOVA) were performed using Prism 5.0 (GraphPad Software, San Diego, CA).

Results

Acute GVHD phenotype at day 14 is absolutely dependent on donor T cell signaling through the p75 receptor

At 14 d after donor cell transfer, B6→F1 acute GVHD mice exhibit a cytotoxic phenotype characterized by significant engraftment of both CD4 and CD8 donor T cells and profound elimination of host splenocytes mediated by donor CD8 CTL specific for host MHC I (reviewed in Refs. 5, 14). By contrast, DBA/2J→F1 chronic GVHD mice exhibit a stimulatory phenotype at day 14 characterized by substantial engraftment of CD4 but not CD8 donor T cells and significant expansion of host B cells, T cells, and APC versus control F1 mice. F1 mice received donor T cells deficient in either p55, p75, or both p55/p75 TNFR and phenotype assessed at day 14 to address whether signaling through the donor T cell TNFR is required for donor CD8 CTL maturation and the development of a cytotoxic phenotype. Donor splenocytes were adjusted so that all mice received comparable numbers of CD8 T cells. Compared with uninjected control F1 mice (Fig. 1A–F, column 1 from left), WT→F1 acute GVHD mice (Fig. 1A–F, column 2 from left) exhibit the characteristic day 14 cytotoxic phenotype, that is, significant reductions in total splenocytes (Fig. 1A), host B cells (Fig. 1B), CD11b+ APCs (Fig. 1C), conventional (CD11c+) DCs (cDCs; Fig. 1D), and CD4 T cells (Fig. 1E). Host CD8 T cells are not significantly reduced versus uninjected control F1 mice (Fig. 1F), consistent with their greater resistance to elimination by donor CTL relative to other host splenocyte populations (15). As expected, WT donor CD4 and CD8 T cells are present in roughly equivalent numbers (Fig. 1G, 1H, column 1). The pattern seen for WT→F1 mice was not significantly altered in p55KO→F1 mice (Fig. 1A–F, column 5) with the exception of greater donor CD8 T cell engraftment (Fig. 1H) consistent with a vigorous donor antihost response (16). By contrast, p75KO→F1 mice exhibit a significant increase in all host splenocyte populations, not only compared with WT→F1 but also and importantly compared with uninjected control F1 mice (Fig. 1A–F, column 4). The increase in host cells seen for p75KO→F1 mice indicates that initial donor CD4-driven host expansion from days 1–7 (15) is present; however, the elimination phase from days 7–14 mediated by donor CD8 CTL is defective, consistent with a stimulatory phenotype and suggestive of chronic GVHD long term. The results for p55/p75KO→F1 mice (Fig. 1, column 3) either mirror those of p75KO→F1 mice (Figs. 1A, 1B, 1D, 1E) or are intermediate between WT→F1 and p75KO→F1 mice (Fig. 1C, 1F). In the p→F1 model, donor T cell engraftment peaks at approximately day 10 followed by contraction, particularly for donor CD8 T cells (15). The significantly greater engraftment of donor CD4 and CD8 T cells in p75KO→F1 versus WT→F1 mice (Fig. 1G, 1H) in the setting of a failure to eliminate host cells is suggestive of impaired CTL effector function or delayed maturation, or both.

FIGURE 1.
FIGURE 1.

Donor T cell signaling through TNFR p75 is critical for induction of acute GVHD phenotype at day 14. Normal (TNFR intact) BDF1 mice were either uninjected or injected with splenocytes from B6 WT, B6 p55KO, B6 p75KO, or B6 p55/p75KO donor cells. At 14 d after donor cell transfer, F1 spleens were assessed for donor and host lymphocyte populations by flow cytometry as described in Materials and Methods. Group means ± SE are shown for (A) total splenocytes, (B) host B cells, (C) host CD11b+ APCs, (D) host cDCs (CD11c+), (E) host CD4 T cells, (F) host CD8 T cells, (G) donor CD4 T cells, and (H) donor CD8 T cells (n = 5 mice/group). Donor splenocytes were examined by flow cytometry and adjusted so that all groups received ∼4 × 106 donor CD8 T cells and 6.7 ± 1.2 × 106 donor CD4 T cells. *p < 0.0, **p < 0.01, ***p < 0.005.

Spleens from the cohort shown in Fig. 1 were also tested for cytokine gene expression by real-time PCR (Fig. 2). Cytokine genes important in CD8 CTL effector function such as IFN-γ, IP-10, and OAS are increased in WT→F1 mice (Fig. 2A–C) as previously demonstrated (17, 18). Increased IL-10 gene expression (Fig. 2D) is also typical of day 14 B6 WT→F1 mice GVHD (18) and contributes to both the exhausted CTL phenotype (19) and memory CD8 generation (20). Cytokine gene expression in p55KO→F1 mice does not differ significantly from WT→F1 mice; however, p75KO→F1 mice exhibit reduced values for all cytokines and reach significance for all except OAS versus WT→F1 mice. Taken together, the results of Figs. 1 and 2 indicate that donor T cell p75 but not p55 signaling is critical for the induction of a day 14 acute GVHD cytotoxic phenotype consistent with impaired donor CD8 CTL elimination of host cells.

FIGURE 2.
FIGURE 2.

Cytokine gene expression typical of day 14 acute GVHD is defective in p75KO→F1 mice. Spleens harvested from the cohort described in Fig. 1 at day 14 were also tested for cytokine gene expression by real time-PCR as described in Materials and Methods. As in Fig. 1, mice were tested individually and values expressed as group mean ± SEM are shown for: (A) IFN-γ, (B) OAS, (C) IP-10, and (D) IL-10. **p < 0.01, ***p < 0.005.

Defective donor T cell p75 signaling converts acute GVHD to chronic GVHD long term

To address whether the day 14 stimulatory phenotype seen in Fig. 1 for p75KO→F1 mice is indicative of a long-term chronic GVHD phenotype or whether donor CD8 CTL maturation is merely delayed and mice eventually exhibit acute GVHD (i.e., elimination of host splenocytes), F1 mice were injected using a protocol similar to that in Fig. 1 and mice followed long term. Despite using a low donor cell dose to reduce acute GVHD-related mortality and permit long-term survival, deaths were seen in all GVHD groups. The overall number of deaths per group was greater in WT→F1 (4/8) and in p55KO→F1 (3/7) versus that of p75KO→F1 (1/7) or p55/075KO→F1 (1/8); however, the differences did not reach statistical significance.

At 20 wk, splenocytes from surviving mice were analyzed by flow cytometry (Fig. 3). Both WT→F1 and p55KO→F1 mice (Fig. 3, columns 2, 5) exhibit a typical long-term acute GVHD phenotype (15); that is., significant elimination of host B cells (Fig. 3A) with repopulation by donor B cells (Fig. 3B) and elimination of host CD11b APC (Fig. 3C), cDC (Fig. 3D), CD4 (Fig. 3E), and CD8 T cells (Fig. 3F) with demonstrable engraftment of both donor CD4 (Fig. 3G) and CD8 T cells (Fig. 3H). p55KO→F1 mice did not exhibit a phenotype significantly different from WT→F1 with the exception of a mild but significant reduction in donor B cells repopulation. Interestingly, engraftment of p55KO donor T cells is also significantly reduced versus WT→F1 donor cells, yet the elimination of all host cell populations is comparable with that of WT→F1, indicating that reduced p55KO donor T cell engraftment is not associated with impaired acute GVHD phenotype long term. By contrast, p75KO→F1 mice (Fig. 3, column 4) exhibit host B cell expansion (Fig. 3A) with little detectable repopulation by donor B cells (Fig. 3B), no significant elimination of host CD11b+ APC, DC, CD4, or CD8 T cells versus control F1 (Fig. 3C–F). Engraftment of donor p75KO CD4 and CD8 T cells is significantly reduced versus WT →F1 donor T cells (Fig. 3G, 3H). Together, these features are consistent with a chronic GVHD phenotype (5, 17, 18). Double knockout (KO) p55/p75KO→F1 mice for the most part resemble that of p75KO→F1 mice and do not exhibit a clearly distinct phenotype.

FIGURE 3.
FIGURE 3.

Long term, p75KO→F1 mice exhibit chronic rather than acute GVHD. Donor and host combinations are as described for Fig. 1. Mice were assessed at 20 wk by flow cytometry for the following splenic lymphocyte subpopulations: (A) host B cells, (B) donor B cells, (C) host CD11B+ APCs, (D) host cDCs (CD11C+), (E) host CD4 T cells, (F) host CD8 T cells, (G) donor CD4 T cells, and (H) donor CD8 T cells. Mice received unfractionated donor splenocytes containing 7.1 ± 0.7 × 106 CD4 T cells and 4.6 ± 0.6 × 106 CD8 T cells. For normal F1, n = 5; WT→F1 and p55 →F1, n = 4; p75KO→F1, n = 6; and p55/p75KO→F1, n = 7. *p < 0.0, **p < 0.01, ***p < 0.005.

Cardinal features of chronic GVHD in the p→F1 model are lupus-like autoantibodies and immune complex glomerulonephritis (4, 5). For BDF1 hosts, severe lupus-like ICGN is seen after transfer of DBA/2J parental cells (4+ proteinuria, nephrotic syndrome) (13), whereas a mild nephritis follows transfer of B6 CD4 T cell donors (≤2+ proteinuria), unless higher donor cell doses (≥15 × 106 CD4 T cells) are used (10, 21). Accordingly, we observed that proteinuria was ≤2+ and glomerular scores were ≤ 1+ for all groups, with no significant differences between groups (data not shown) and consistent with the relatively low numbers of donor CD4 T cells transferred (<8 × 106). Nevertheless, despite the low intensity of the chronic GVHD reaction, striking and significant differences were seen in serum anti-ssDNA Ab, a nonspecific marker of host B cell activation characteristic of chronic GVHD (10). Only p75KO→F1 and p55/p75KO→F1 mice exhibited significantly elevated serum anti-ssDNA Ab levels versus control F1 as early as 2 wk and persisting through week 14 (Fig. 4). The two curves for p75KO→F1 and p55/p75KO→F1 were not significantly different, further supporting the idea that p55/p75KO→F1 mice have a phenotype similar to p75kO→F1 mice. No significant elevations over uninjected controls were seen for WT→F1 or p55KO→F1 mice.

FIGURE 4.
FIGURE 4.

Lupus-like autoantibodies are seen only in F1 mice receiving p75KO donor T cells. Sera was tested for anti-ssDNA Abs on the cohort described in Fig. 3 at the indicated times as described in Materials and Methods. ***p < 0.005.

Taken together, these results demonstrate that both WT→F1 and p55KO→F1 mice exhibit an acute GVHD phenotype long term, whereas p75KO→F1 and p55/p75KO→F1 mice exhibit a chronic GVHD phenotype consistent with a defect in donor CD8 effector maturation and function in the setting of donor CD4-driven host B cell expansion and autoimmunity.

CD8 CTL effector phenotype requires donor T cell signaling only through the p75 TNFR

In B6 WT→F1 mice, donor CD8 numbers peak at approximately day 10 and mediate the decline in host lymphocyte expansion (15). By contrast, chronic GVHD in this model results from a failure of donor CD8 CTL maturation in the context of intact donor CD4-driven host splenocyte expansion. Thus, the foregoing results demonstrating that p75KO→F1 mice exhibit a chronic GVHD phenotype short and long term strongly support a defect in p75KO donor CD8 CTL effector function coupled with intact donor CD4 T cell help for host B cells. GVHD was induced using a protocol similar to Fig. 1 and mice were assessed at the peak of donor CD8 CTL expansion (day 10), to determine the integrity of initial maturation of donor CD8 T cells (15). Because p75/p55KO→F1 mice did not exhibit a unique phenotype but were either intermediate or similar to p75KO→F1 mice, this group is omitted to focus on the dichotomy between p55KO→F1 versus p75KO→F1 mice. Compared with control F1 mice (Fig. 5A–F, column 1), WT→F1 mice (Fig. 5A–F, column 2) exhibit: 1) a mild increase in total splenocytes (Fig. 5A); 2) significant elimination of host B cells (Fig. 5B); 3) expansion of host DCs and CD11b+ APCs (Fig. 5C, 5D); 4) reduction of host CD4 T cells (Fig. 5E), but not host CD8 T cells (Fig. 5F); and 5) engraftment of both CD4 and CD8 B6 WT donor T cells (Fig. 5G, 5H, column 1). This pattern confirms previous work at day 10 demonstrating that host B cells and CD4 T cells are more sensitive to elimination and are eliminated earlier than host APCs or CD8 T cells (15). Values for p55KO→F1 mice (Fig. 5A–F, column 4, Fig. 5G, 5H, column 3) do not differ significantly from WT→F1 mice. By contrast, p75KO→F1 mice (Fig. 5A–F, column 3) exhibit a significant increase in total splenocytes, host B cells, host DCs, host CD11b+ APCs, and host CD4 T cells (Fig. 5A–E) compared with either WT→F1 or p55KO→F1 mice. The increase in host cells seen in p75KO→F1 mice is an indicator of impaired donor CD8 CTL elimination of intact donor CD4-driven host splenocyte expansion. Interestingly, p75KO→F1 mice exhibit significantly greater donor CD4 engraftment versus WT→F1 or p55R KO→F1 mice (Fig. 5G), yet CD8 T cell engraftment is mildly but significantly reduced versus WT CD8 T cells (Fig. 5H), consistent with a mild but significant quantitative defect in donor CD8 T cell expansion. A second day 10 experiment gave similar results; however, the reduction in engrafted donor CD8 T cells for p75KO→F1 mice did not reach statistical significance as it did in Fig. 5H (data not shown).

FIGURE 5.
FIGURE 5.

Peak day 10 donor CD8 T cell engraftment is defective in p75KO→F1 mice. Acute GVHD was induced in BDF1 hosts after the transfer of B6 WT, p55KO, or p75KO donor splenocytes containing 7.15 ± 0.35 × 106 CD4 and 4.35 ± 0.45 × 106 CD8 T cells. F1 hosts were assessed by flow cytometry on day 10 for: (A) total splenocytes, (B) host B cells, (C) host cDC, (D) host CD11b+ APCs, (E) host CD4 T cells, (F) host CD8 T cells, (G) donor CD4 T cells, and (H) donor CD8 T cells. Results are shown as group mean ± SE (n = 5 mice/group). *p < 0.0, **p < 0.01, ***p < 0.005.

Defective p75KO donor CD8 CTL elimination of host cells can be partially compensated by transferring greater numbers

We have previously demonstrated that transferring standard dose donor cells with qualitative CTL defects (i.e., perforin or FasL defective) results in an intermediate phenotype at day 14 because of impaired CTL elimination of host splenocytes (22, 23). Higher doses of qualitatively defective donor cells improved but did not fully correct the impaired acute GVHD phenotype (22, 23). Thus, qualitative defects in CD8 T cell function can be partially but not fully compensated by quantitative measures. To address whether the impaired elimination of host cells in p75KO→F1 mice is due solely to the quantitative defect (shown in Fig. 5H as a significant reduction in peak donor CD8 T cell numbers) or whether a qualitative aspect also contributes, we examined the effect of higher donor cell transfers. F1 mice received either: 1) low-dose WT or p75KO donor cells (doses comparable with those used in Figs. 1–5), or 2) high-dose p75KO donor cells and mice assessed at day 14. Compared with uninjected F1 mice, low-dose WT→F1 exhibited the expected cytotoxic phenotype (Fig. 6, columns 1, 2) consisting of profound elimination of host B cells (Fig. 6A), no increase in host CD11c+ cells (Fig. 6B), and engraftment of both donor CD4 and CD8 T cells subsets (Fig. 6C, 6D). Similarly, low-dose p75KO→F1 (Fig. 6, column 3) reproduced the day 14 phenotype shown in Fig. 1A, that is, a failure to eliminate host B cells or CD11c+ cells (Fig. 6A, 6B), normal engraftment of donor CD4 T cells (Fig. 6C) coupled with a mild but significant reduction in donor CD8 T cell engraftment. High-dose p75KO→F1 mice (Fig. 6, column 4) exhibited significantly greater elimination of host B cells versus low-dose p75KO→F1 mice (Fig. 6A); however, B cell numbers were still significantly greater than that of WT→F1 mice consistent with impaired donor CD8 CTL killing. High-dose p75KO→F1 mice did not exhibit significantly greater killing of host DCs (Fig. 6B), host CD11b+APC, or host CD4 T cells (data not shown). Importantly, failure to correct host killing in high-dose p75KO→F1 mice occurred in conjunction with donor CD4 and CD8 T cell engraftment levels that were comparable with or significantly greater those of WT→F1 mice (Fig. 6C, 6D, columns 1, 3). These results confirm our day 14 data in Fig. 1 in which p75KO→F1 mice exhibit defective elimination of donor CD4 expanded host B cell and APC (Fig. 1A–C) despite significantly greater donor CD8 T cell engraftment compared with WT→F1 mice (Fig. 1H). Thus, transferring donor cell numbers sufficient to engraft p75KO donor cells in numbers greater than or equal to those of WT donor cells improves but does not fully correct the defects in host splenocyte elimination. These results support the conclusion that defective donor T cell p75 signaling results in qualitative defects in effector CTL function in addition to quantitative defects.

FIGURE 6.
FIGURE 6.

Defective donor CD8 CTL killing by p75KO donor T cells is only partially corrected by normalizing engraftment. Acute GVHD was induced in BDF1 hosts after the transfer of low-dose B6 WT or p75KO splenocytes (6.6–6.8 × 106 CD4 and 4.8–4.9 × 106 CD8 T cells) or high-dose p75KO donor cells (8.8 × 106 CD4 and 6.4 × 106 CD8 T cells). F1 hosts were assessed at day 14 by flow cytometry for lymphocyte subsets as described in Fig. 1 and shown as: (A) host B cells, (B) host cDCs, (C) donor CD4, and (D) donor CD8 T cells. Results are shown as group mean ± SE (n = 5 mice/group except for high-dose p75KO→F1, n = 3). *p < 0.0, **p < 0.01, ***p < 0.005.

Optimal donor CD8 CTL killing requires p75 signaling by both donor CD4 and CD8 T cells

The foregoing results demonstrate that optimal CD8 CTL elimination of host splenocytes requires TNFR signaling through p75 on donor T cells. It is not clear whether this effect is mediated by p75 signaling through CD8 T cells alone, through CD4 T cells alone, or through both. To address this question, we transferred purified donor CD4 and CD8 T cells from both WT and p75KO donors into WT BDF1 mice in all four possible matched or mixed combinations and assessed phenotype during peak donor CD8 CTL expansion and maturation (day 10). As shown in Fig. 6, defects in p75KO donor cells can be improved using larger numbers of donor cells. Therefore, to better demonstrate possible mild contributory defects in CD4 T cells, we transferred subthreshold numbers of CD4 donor cells in conjunction with numbers of CD8 T cells at or above threshold (6.0 ± 0.3 × 106 CD4 and CD8 T cells), resulting in a CD4/CD8 ratio of 1:1 rather than the typical 2:1. Using host B cell elimination as a surrogate marker of donor antihost CD8 CTL function (15, 24), F1 hosts receiving matched WT donor T cells, that is, WT CD4 + WT CD8 →F1 (matched WT control), exhibit a mild reduction in host B cells versus control uninjected F1 mice (Fig. 7A, columns 1, 2; p value is NS), consistent with the day 10 readout at which point donor CD8 CTL maturation typically has reversed the initial donor CD4-mediated host B cell expansion; however, elimination of host B cells is not complete until approximately day 14 (15). By contrast, B cells in p75KO CD4 + p75KO CD8→F1 mice (matched p75 control) are significantly elevated versus uninjected control F1 (Fig. 1A, columns 3 and 1), indicative of an intact donor CD4-driven expansion phase, but a failure of donor CD8 CTL-mediated elimination phase consistent with a stimulatory chronic GVHD phenotype and confirming our results in Fig. 1A using unfractionated p75KO donor splenocytes. For F1 mice receiving mixed donor T cells, that is, WT CD4 + p75KO CD8→F1 or p75KO CD4 + WT CD8→F1 (Fig. 7A, columns 4, 5), both groups exhibit B cell elimination values comparable with matched WT CD4 + WT CD8→F1 control (Fig. 7A, column 2), and are significantly reduced versus the matched p75KO CD4 + p75KO CD8→F1 control (Fig. 7A, column 3). These results support a role for p75 signaling in both CD4 and CD8 T cells.

FIGURE 7.
FIGURE 7.

Optimal donor CD8 CTL function requires p75 signaling in both CD4 and CD8 T cells. CD4 and CD8 T cells were purified from WT and p75KO donor spleens, and combined as described in Materials and Methods. BDF1 mice received 6.0–6.7 × 106 CD4 and 5.7–6.2 × 106 CD8 T cells that were either matched or mixed according to donor strain. At day 10, host spleens were analyzed by flow cytometry for numbers of (A) host B cells, (B) donor CD4 T cells, (C) donor CD8 T cells, and (D) KLRG-1+ donor CD8 T cells. Results are shown as group mean ± SE (n = 5 mice/group for all groups). *p < 0.0, **p < 0.01, ***p < 0.005.

Specifically, the defective host B cell killing (increased host B cell numbers versus control F1) seen for p75KO CD4 + p75KO CD8→F1 mice (Fig. 7A, column 3) can be significantly improved by substituting either WT CD4 T (Fig. 7A, column 4) or WT CD8 T cells (Fig. 7A, column 5). Conversely, B cell killing for WT CD4 + WT CD8→F1 (Fig. 7A, column 2) is not significantly altered by the substitution of only one p75KO T cell subset (Fig. 7A, column 4 or 5). The results in Fig. 7A demonstrate that defective p75 signaling in both CD4 or CD8 T cells impairs optimal CD8 CTL maturation as measured by host B cell killing; however, the defect in each subset can be significantly mitigated by pairing with the appropriate WT subset.

This conclusion is further supported by donor engraftment results. CD4 T cell engraftment levels (Fig. 7B) support a critical role for CD4 p75 signaling in optimal CD4 engraftment. Engraftment of WT CD4 T cells does not differ significantly whether paired with WT or p75KO CD8 T cells (Fig. 7B, columns 1, 3). Engraftment of p75KO CD4 engraftment is significantly reduced versus WT CD4 regardless of the source of CD8 T cells. Specifically, both p75KO CD4 + p75KO CD8→F1 and p75RKO CD4 + WT CD8→F1 mice (Fig. 7B, columns 2, 4) exhibit a significant reduction in donor CD4 T cell engraftment versus F1 mice receiving WT CD4 T cells in either combination (WT CD4 + WT CD8→F1 or WT CD4 + p75KO CD8→F1; Fig. 7B, columns 1, 3). Thus, p75 signaling is critical for optimal donor CD4 T cell engraftment and is not altered by mixed pairing of CD8 T cells.

Similarly, donor CD8 T cell engraftment levels (Fig. 7C) support a role for intrinsic CD8 p75 signaling in optimal engraftment. Compared with the matched WT control (Fig. 7C, column 1), matched p75KO control mice (Fig. 7C, column 2) exhibit significantly reduced engraftment of donor CD8 T cells. These results are consistent with the day 10 peak CD8 engraftment results demonstrating a mild but significant reduction in CD8 engraftment for p75KO→F1 mice (Fig. 5H). They differ from our day 14 results in Fig. 1 using unfractionated cells possibly as a consequence of the greater numbers of donor cells used in that experiment. Defective CD8 T cell engraftment seen in the matched p75KO control mice (Fig. 7C, column 2) is mildly but not significantly boosted when paired with WT CD4 T cells (Fig. 7C, column 3), indicative of an intrinsic p75-mediated CD8 defect. Of note, engraftment of WT CD8 T cells is mildly but not significantly reduced when paired with p75KO CD4 T cells versus WT CD4 T cells (Fig. 7C, column 1 versus 4), consistent with a mild helper defect for p75KO CD4 T cells.

Lastly, KLRG-1 expression is a marker of CD8 CTL effector maturation and KLRG-1pos/CCR7neg CD8 T cells represent short-lived effector cells (SLEC) (19). As shown in Fig. 7D, SLEC numbers are profoundly defective in p75KO matched control mice (Fig. 7D, column 2) versus that of the matched WT control (Fig. 7D, column 1) with values below the limits of detection. Reduced p75KO CD8 SLEC numbers seen in p75KO matched mice (Fig. 7D, column 2) are mildly improved but not corrected by pairing with normal CD4 help (WT CD4 + p75KO CD8→F1, column 3) consistent with a significant intrinsic defect in p75RKO CD8 T cells. Conversely, WT SLEC CD8 numbers seen in WT control mice are moderately but significantly reduced when paired with p75KO CD4 T cells versus WT CD4+ T cells (Fig. 7D, columns 1, 4), further supporting a mild but contributory role for CD4 p75 signaling in providing help for normal CD8 CTL effector maturation. Of note, mice receiving mixed WT CD4 and p75KO CD8 donor T cells exhibit significantly reduced CD8 engraftment and KLRG-1 upregulation versus WT matched control (Fig. 7C, 7D, columns 1 and 3); however, this was not associated with a significant impairment of host B cell killing (Fig. 7A, column 4), indicating the mild nature of the p75KO defect when only one subset is affected.

Taken together, the results in Fig. 7 demonstrate that p75 signaling is critical for optimal donor CD4 and CD8 engraftment, for optimal CD4 help for CD8 CTL elimination of B cells, and for optimal effector marker upregulation (KLRG-1).

p75KO donor T cell signaling is required for optimal CD4 and CD8 intracellular cytokine expression

Splenic donor T cells from the cohort shown in Fig. 7 were analyzed by flow cytometry for intracellular markers characteristic of effector maturation, that is, TNF, IFN-γ, and perforin (Fig. 8). Representative staining plots are shown in Supplemental Fig. 1. These markers typically peak between days 7 and 10, and accordingly, WT matched controls exhibit detectable numbers of cytokine expressing cells for all cytokines except perforin expressing CD4 T cells (Fig. 8, column 1). Compared with the WT matched control, p75 matched control mice (Fig. 8, column 2 versus 1) exhibit significant reductions in the numbers of: 1) CD4 T cells expressing TNF and IFN-γ (Fig. 8A, 8C); and 2) CD8 T cells expressing TNF, IFN-γ, and perforin (Figs. 8B, 8D, 8E). Numbers of perforin expressing CD4 T cells were too small for all groups to allow meaningful conclusions (data not shown). The results for the two matched control groups demonstrate that when both donor T cell subsets are p75 defective, significant qualitative defects in both donor CD4 and CD8 T cell effector maturation can be seen.

FIGURE 8.
FIGURE 8.

Intact p75 signaling is required for optimal donor CD4 and CD8 intracellular expression of effector CTL markers. Spleens from the cohort in Fig. 7 were examined on day 10 by flow cytometry for intracellular expression of TNF (A, B), IFN-γ (C, D), and perforin (E) on donor CD4 (A, C) or CD8 (B, D, E) T cells. *p < 0.0, **p < 0.01, ***p < 0.005.

Mixing of donor T cell subsets improves but does not correct the defects seen for matched p75KO donor cells, further supporting the idea that both CD4 and CD8 p75 signaling is required for optimal CD8 effector maturation. In general, mixed subsets (Fig. 8, columns 3, 4) exhibit values that are intermediate between the matched WT or p75KO controls (Fig. 8, columns 1, 2). In particular, pairing of WT CD4 T cells with p75KO CD8 T cells (Fig. 8, column 3) results in a significant boost in all CD4 and CD8 parameters versus matched p75KO control (Fig. 8, column 2); however, for the most part these values do not reach those of the matched WT control (Fig. 8A–D, column 1). Thus, WT CD4 T cells can boost but not normalize defective p75KO CD8 T cell IFN-γ, TNF, and perforin expression (Fig. 8B, 8D, 8E, columns 3 versus 2), indicating that even with a normal source of CD4 help, p75KO CD8 T cells remain intrinsically abnormal.

Similarly, p75KO CD4 T cells exhibit a mild-to-moderate defect in their help for CD8 T cells as shown by pairing of p75KO CD4 with WT CD8 T cells and compared with values of the WT control (Fig. 8, columns 1, 4). This helper defect is best seen as a significant reduction in WT CD8 expression of IFN-γ and a nonsignificant reduction in perforin expression when paired with p75 CD4 T cells versus WT CD4 T cells (Fig. 8D, 8E, columns 1, 4). There was also a significant reduction in p75KO CD4 + WT CD8→F1 TNF expression in CD8 T cells; however, the mean value was increased because of mouse-to-mouse variability. The results in Fig. 8 further support the conclusion from Fig. 7 that optimal quantitative and qualitative CD8 CTL effector maturation requires p75 signaling by both CD4 and CD8 T cells.

Discussion

CD8 CTL generation in the p→F1 model has been previously shown to be critically dependent on TNF (6). In B6→BDF1 mice, anti-TNF mAb treatment during days 0–5 prevented donor CD8 CTL maturation and converted phenotype from acute to chronic GVHD, whereas TNF blockade on day 7 had little effect, supporting the idea that TNF is critical in CD8 CTL initiation rather than acting later at the effector phase. It was not clear from that study, however, whether TNF acted directly on donor T cells in a signal 3/costimulatory role or acted indirectly, perhaps through host APCs. Our results confirm the former possibility by demonstrating that TNF signaling through donor T cell TNFR p75, but not p55, is absolutely required for CD8 CTL effector maturation. CD4 T cell signaling through p75 is required for optimal CD4 help for CD8 maturation, and CD8 T cell signaling through p75 is required for optimal CTL effector function. Interestingly, direct measurement of p75KO donor cell quantitative and qualitative parameters, for example, donor T cell engraftment and intracellular effector markers, revealed only mild defects for mice receiving unfractionated donor cells. By contrast, indirect parameters of donor CD8 CTL function, for example, elimination of host splenocytes, revealed profound and long-lasting defects such that at 2 wk, p75KO→F1 mice exhibited a stimulatory phenotype rather than the expected cytotoxic phenotype that persisted long term with p75KO→F1 mice exhibiting chronic rather than acute GVHD. These results are consistent with previous work demonstrating that host B cell depletion is a more sensitive indicator of in vivo donor CD8 CTL activity than are CTL assays performed in vitro, ex vivo, or even in vitro (15, 24).

Because chronic GVHD in this model is mediated solely by donor CD4 T cells, whereas acute GVHD requires both CD4 and CD8 donor T cells (5), the conversion from acute to chronic GVHD in p75KO→F1 mice indicates a significant defect in donor p75KO CD8 T cell maturation. In support of this idea, dose–response studies demonstrated that transferring sufficient donor cells to achieve p75KO T cell engraftment greater than that of WT donor cells improved but did not correct the profound defects in host B cell elimination in p75KO→F1 mice. These results provide indirect evidence for a significant qualitative defect in p75KO CD8 effector function. Direct evidence for a p75KO donor cell qualitative defect was best seen at donor cell doses just above the threshold for GVHD induction and consisted of defective intracellular expression of IFN-γ and TNF for both CD4 and CD8 T cells, and defective perforin expression for donor CD8 T cells. Together, these results support the conclusion that p75 signaling in both Ag-specific CD4 and CD8 T cell subsets is critical for Ag-specific CD8 CTL maturation and effector function. By contrast, no defects were seen in acute GVHD phenotype short or long term when p55KO donor cells were transferred. These results appear to be at variance with results in a murine model of acute GVHD using irradiated bone marrow transplant recipients and demonstrating that p55, but not p75, signaling is critical for acute GVHD (25). This discrepancy likely reflects the strong inflammatory cytokine storm and inflammation elicited by the conditioning regimen in that model and the well-known role of p55 signaling in TNF-mediated inflammation (26, 27). By contrast, in the p→F1 model, there is no conditioning regimen and no similar cytokine storm, thus mitigating the role of p55-mediated inflammation.

It has been previously demonstrated that the CD8 CTL antipathogen response requires a third signal in addition to TcR stimulation and CD28-mediated costimulation (1, 3). Two well-recognized signal 3 molecules are IL-12 and IFN (1, 3). TNFR superfamily members also influence T cell activation and maturation by providing costimulation signals separate from CD28 signaling. OX-40, 4-B11, CD27 HVEM, CD30, and GITR can all provide signals important in the initiation of immune responses and in long-lived memory (28). In addition, TNFR p75 signaling can provide a costimulatory signal for T cells (28) that ensures optimal IL-2 production and T cell survival during clonal expansion and enhances memory pool formation by conferring protection from apoptosis (29–31). As a result, p75KO T cells have been reported to exhibit a reduced accumulation of effector cells as shown by reduced numbers of Ag-specific intracellular IFN-γ–expressing cells and reduced clearance of Listeria monocytogenes (29). Our results, although consistent with these quantitative defects, differ in that we also demonstrate a qualitative defect in p75KO CD8 CTL killing and further, that this defect is not corrected by normalization of p75KO T cell numbers.

A second novel finding of our study regards the qualitative defect in p75KO CD4 T cell function. As discussed earlier, p75KO→F1 mice exhibit chronic rather than acute GVHD because of defective CD8 CTL maturation. Chronic GVHD in the p→F1 model is mediated solely by donor CD4 T cell recognition allogeneic host MHC class II and the provision of cognate CD4 T cell help to host B cells resulting in B cell hyperactivity and autoantibody production (32). Although p75KO CD4 T cells are defective in their help for CD8 CTL maturation, they are nevertheless able to provide help to host B cells and induce lupus-like autoantibodies and chronic GVHD. Thus, p75 signaling is much more important for CD4 T cell help to CD8 CTL than it is for CD4 help to B cells. This functional disparity in CD4 T cell help is reminiscent of our findings with another TNFR super family member, Fas, in which Fas defective donor CD4 T cells exhibit significantly impaired help for CD8 CTL but no detectable defect in their ability to provide help to B cells and drive chronic, lupus-like GVHD (10).

Lastly, our results may have relevance to patients receiving therapeutic TNF blockers. Our demonstration of a critical role for p75 signaling in optimal in vivo CD8 CTL function raises the concern that TNF blockers may impair the CD8 CTL contribution to antiviral and antitumor responses. For example, CD8 CTL are strongly linked to recovery from EBV infection (33), and TNF is critical for the development of HBV-specific CTL (34). Incidents of reactivation of EBV, HBV, varicella zoster, and less commonly, hepatitis C have been reported in patients on TNF blockers (35–37). Although the numbers are small, in the absence of long-term prospective studies, a definitive statement regarding the safety of TNF blockers in viral conditions cannot be made, and in the case of HBV, prophylactic antiviral therapy and close clinical monitoring have been suggested (38, 39).

Regarding tumors, estimating the cancer risk of TNF blockers separate from that of the underlying condition and the concomitant use of immunosuppressives is complicated by the long latency of tumors, their relative low incidence, and methodological concerns, for example, discrepancies between observational studies and meta-analyses (40, 41). Accordingly, reports of an increase in malignancies (42), particularly hematological malignancies (43), in patients receiving TNF blockers have not been uniformly confirmed (40, 44). Nevertheless, evidence supporting an increased risk for cancer, particularly lymphomas, in children and adolescents receiving TNF blockers (45), prompted the U.S. Food and Drug Administration to issue a warning (46). TNF signaling particularly through the p75 has been shown to be critical to antitumor T cell responses (47). Moreover, Fas and perforin pathways are important in normal lymphocyte homeostasis (48–50). Thus, impairment of p75 T cell signaling by TNF blockers could impair T cell–mediated tumor surveillance.

Lastly, our results may be relevant to the lupus-like autoimmunity reported in patients receiving TNF blockers (47, 51, 52) and suggest a possible mechanism by which this could occur. In both humans and mice, CD4 T cells are necessary and sufficient for lupus development (53–57), and CD8 T cells, to include CTL (5, 58–63), act to downregulate lupus. Based on our results demonstrating that p75 signaling is important for CD8 effector maturation, but not for CD4 T cell help to B cells, it is possible that patients who experience a lupus-like tolerance break (i.e., CD4-driven B cell hyperactivity) while on TNF blockers may exhibit an impairment in the ability of down-regulatory CD8 T cells to restore tolerance without a concomitant impairment of CD4 T cells to provide help to autoreactive B cells and autoantibody production. For this scenario to occur, patients would first need to encounter a lupus trigger, which may account for the relative low frequency of autoimmunity in the setting of TNF blockers. Once tolerance is lost, the balance between lupus and CD8-mediated downregulation may be shifted such that autoimmunity is favored. The possible benefit of therapeutic selective p55 or p75 blockade was put forth roughly a decade ago (64) and remains under active study (9, 65). Moreover, soluble TNF efficiently activates only p55, whereas membrane TNF is largely responsible for p75 activation (9, 66). By extension, our results support the conclusion that it is membrane bound rather than soluble TNF binding to donor T cell p75 that is critical for CD8 CTL maturation in the absence of pathogens. In light of the beneficial role of CD8 T cells in infections, tumors, and autoimmunity, our results demonstrating a critical role for p75 signaling in CTL maturation provide support for further investigation of TNF blockers that selectively inhibit p55 and spare p75, to include differential targeting of membrane versus soluble TNF.

Disclosures

The authors have no financial conflicts of interest.

Footnotes

  • This work was supported by National Institutes of Health Grant AI047466 (to C.S.V.).

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    BDF1
    B6D2F1
    B6
    C57BL/6
    cDC
    conventional dendritic cell
    DC
    dendritic cell
    GVHD
    graft-versus-host disease
    IP-10
    inducible protein 10
    KO
    knockout
    OAS
    oligoadenylate synthetase-1
    p→F1
    parent-into-F1
    p55
    TNFR 1
    p75
    TNFR 2
    p55KO
    B6.Tnfrsf1atm1Imx/J
    p75KO
    B6.Tnfrsf1btm1Imx/J
    p55/p75KO
    B6.Tnfrsf1atm1ImxTnfrsf1btm1Imx /J
    SLEC
    short-lived effector cell
    WT
    wild type.

  • Received January 10, 2013.
  • Accepted February 25, 2013.

References

  1. ↵
    1. Mescher M. F.,
    2. J. M. Curtsinger,
    3. P. Agarwal,
    4. K. A. Casey,
    5. M. Gerner,
    6. C. D. Hammerbeck,
    7. F. Popescu,
    8. Z. Xiao
    . 2006. Signals required for programming effector and memory development by CD8+ T cells. Immunol. Rev. 211: 81–92.
    OpenUrlCrossRefPubMed
  2. ↵
    1. Curtsinger J. M.,
    2. M. Y. Gerner,
    3. D. C. Lins,
    4. M. F. Mescher
    . 2007. Signal 3 availability limits the CD8 T cell response to a solid tumor. J. Immunol. 178: 6752–6760.
    OpenUrlAbstract/FREE Full Text
  3. ↵
    1. Curtsinger J. M.,
    2. M. F. Mescher
    . 2010. Inflammatory cytokines as a third signal for T cell activation. Curr. Opin. Immunol. 22: 333–340.
    OpenUrlCrossRefPubMed
  4. ↵
    1. Eisenberg R. A.,
    2. C. S. Via
    . 2012. T cells, murine chronic graft-versus-host disease and autoimmunity. J. Autoimmun. 39: 240–247.
    OpenUrlCrossRefPubMed
  5. ↵
    1. Via C. S.
    2010. Advances in lupus stemming from the parent-into-F1 model. Trends Immunol. 31: 236–245.
    OpenUrlCrossRefPubMed
  6. ↵
    1. Via C. S.,
    2. A. Shustov,
    3. V. Rus,
    4. T. Lang,
    5. P. Nguyen,
    6. F. D. Finkelman
    . 2001. In vivo neutralization of TNF-alpha promotes humoral autoimmunity by preventing the induction of CTL. J. Immunol. 167: 6821–6826.
    OpenUrlAbstract/FREE Full Text
  7. ↵
    1. Aggarwal B. B.,
    2. T. E. Eessalu,
    3. P. E. Hass
    . 1985. Characterization of receptors for human tumour necrosis factor and their regulation by gamma-interferon. Nature 318: 665–667.
    OpenUrlCrossRefPubMed
    1. Leist M.,
    2. F. Gantner,
    3. S. Jilg,
    4. A. Wendel
    . 1995. Activation of the 55 kDa TNF receptor is necessary and sufficient for TNF-induced liver failure, hepatocyte apoptosis, and nitrite release. J. Immunol. 154: 1307–1316.
    OpenUrlAbstract
  9. ↵
    1. Faustman D.,
    2. M. Davis
    . 2010. TNF receptor 2 pathway: drug target for autoimmune diseases. Nat. Rev. Drug Discov. 9: 482–493.
    OpenUrlCrossRefPubMed
  10. ↵
    1. Puliaeva I.,
    2. R. Puliaev,
    3. A. Shustov,
    4. M. Haas,
    5. C. S. Via
    . 2008. Fas expression on antigen-specific T cells has costimulatory, helper, and down-regulatory functions in vivo for cytotoxic T cell responses but not for T cell-dependent B cell responses. J. Immunol. 181: 5912–5929.
    OpenUrlAbstract/FREE Full Text
  11. ↵
    1. Soloviova K.,
    2. M. Puliaiev,
    3. A. Foster,
    4. C. S. Via
    . 2012. The parent-into-F1 murine model in the study of lupus-like autoimmunity and CD8 cytotoxic T lymphocyte function. Methods Mol. Biol. 900: 253–270.
    OpenUrlCrossRefPubMed
  12. ↵
    1. Passwell J.,
    2. G. F. Schreiner,
    3. M. Nonaka,
    4. H. U. Beuscher,
    5. H. R. Colten
    . 1988. Local extrahepatic expression of complement genes C3, factor B, C2, and C4 is increased in murine lupus nephritis. J. Clin. Invest. 82: 1676–1684.
    OpenUrlCrossRefPubMed
  13. ↵
    1. Foster A. D.,
    2. M. Haas,
    3. I. Puliaeva,
    4. K. Soloviova,
    5. R. Puliaev,
    6. C. S. Via
    . 2010. Donor CD8 T cell activation is critical for greater renal disease severity in female chronic graft-vs.-host mice and is associated with increased splenic ICOS(hi) host CD4 T cells and IL-21 expression. Clin. Immunol. 136: 61–73.
    OpenUrlCrossRefPubMed
  14. ↵
    1. Puliaeva I.,
    2. K. Soloviova,
    3. M. Puliaiev,
    4. T. Lang,
    5. R. Puliaev,
    6. C. S. Via
    . 2011. Enhancement of suboptimal CD8 cytotoxic T cell effector function in vivo using antigen-specific CD80 defective T cells. J. Immunol. 186: 291–304.
    OpenUrlAbstract/FREE Full Text
  15. ↵
    1. Puliaev R.,
    2. I. Puliaeva,
    3. L. A. Welniak,
    4. A. E. Ryan,
    5. M. Haas,
    6. W. J. Murphy,
    7. C. S. Via
    . 2008. CTL-promoting effects of CD40 stimulation outweigh B cell-stimulatory effects resulting in B cell elimination and disease improvement in a murine model of lupus. J. Immunol. 181: 47–61.
    OpenUrlAbstract/FREE Full Text
  16. ↵
    1. Foster A. D.,
    2. K. Soloviova,
    3. I. Puliaeva,
    4. M. Puliaiev,
    5. R. Puliaev,
    6. F. Finkelman,
    7. C. S. Via
    . 2011. Donor CD8 T cells and IFN-gamma are critical for sex-based differences in donor CD4 T cell engraftment and lupus-like phenotype in short-term chronic graft-versus-host disease mice. J. Immunol. 186: 6238–6254.
    OpenUrlAbstract/FREE Full Text
  17. ↵
    1. Puliaeva I.,
    2. R. Puliaev,
    3. C. S. Via
    . 2009. Therapeutic potential of CD8+ cytotoxic T lymphocytes in SLE. Autoimmun. Rev. 8: 219–223.
    OpenUrlCrossRefPubMed
  18. ↵
    1. Rus V.,
    2. A. Svetic,
    3. P. Nguyen,
    4. W. C. Gause,
    5. C. S. Via
    . 1995. Kinetics of Th1 and Th2 cytokine production during the early course of acute and chronic murine graft-versus-host disease. Regulatory role of donor CD8+ T cells. J. Immunol. 155: 2396–2406.
    OpenUrlAbstract
  19. ↵
    1. Parish I. A.,
    2. S. M. Kaech
    . 2009. Diversity in CD8(+) T cell differentiation. Curr. Opin. Immunol. 21: 291–297.
    OpenUrlCrossRefPubMed
  20. ↵
    1. Foulds K. E.,
    2. M. J. Rotte,
    3. R. A. Seder
    . 2006. IL-10 is required for optimal CD8 T cell memory following Listeria monocytogenes infection. J. Immunol. 177: 2565–2574.
    OpenUrlAbstract/FREE Full Text
  21. ↵
    1. Rus V.,
    2. V. Nguyen,
    3. R. Puliaev,
    4. I. Puliaeva,
    5. V. Zernetkina,
    6. I. Luzina,
    7. J. C. Papadimitriou,
    8. C. S. Via
    . 2007. T cell TRAIL promotes murine lupus by sustaining effector CD4 Th cell numbers and by inhibiting CD8 CTL activity. J. Immunol. 178: 3962–3972.
    OpenUrlAbstract/FREE Full Text
  22. ↵
    1. Shustov A.,
    2. I. Luzina,
    3. P. Nguyen,
    4. J. C. Papadimitriou,
    5. B. Handwerger,
    6. K. B. Elkon,
    7. C. S. Via
    . 2000. Role of perforin in controlling B-cell hyperactivity and humoral autoimmunity. J. Clin. Invest. 106: R39–R47.
    OpenUrlCrossRefPubMed
  23. ↵
    1. Via C. S.,
    2. P. Nguyen,
    3. A. Shustov,
    4. J. Drappa,
    5. K. B. Elkon
    . 1996. A major role for the Fas pathway in acute graft-versus-host disease. J. Immunol. 157: 5387–5393.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Puliaev R.,
    2. P. Nguyen,
    3. F. D. Finkelman,
    4. C. S. Via
    . 2004. Differential requirement for IFN-gamma in CTL maturation in acute murine graft-versus-host disease. J. Immunol. 173: 910–919.
    OpenUrlAbstract/FREE Full Text
  25. ↵
    1. Hill G. R.,
    2. T. Teshima,
    3. V. I. Rebel,
    4. O. I. Krijanovski,
    5. K. R. Cooke,
    6. Y. S. Brinson,
    7. J. L. Ferrara
    . 2000. The p55 TNF-alpha receptor plays a critical role in T cell alloreactivity. J. Immunol. 164: 656–663.
    OpenUrlAbstract/FREE Full Text
  26. ↵
    1. Sun Y.,
    2. I. Tawara,
    3. T. Toubai,
    4. P. Reddy
    . 2007. Pathophysiology of acute graft-versus-host disease: recent advances. Transl. Res. 150: 197–214.
    OpenUrlCrossRefPubMed
  27. ↵
    1. Ferrara J. L.,
    2. J. E. Levine,
    3. P. Reddy,
    4. E. Holler
    . 2009. Graft-versus-host disease. Lancet 373: 1550–1561.
    OpenUrlCrossRefPubMed
  28. ↵
    1. Watts T. H.
    2005. TNF/TNFR family members in costimulation of T cell responses. Annu. Rev. Immunol. 23: 23–68.
    OpenUrlCrossRefPubMed
  29. ↵
    1. Kim E. Y.,
    2. J. J. Priatel,
    3. S. J. Teh,
    4. H. S. Teh
    . 2006. TNF receptor type 2 (p75) functions as a costimulator for antigen-driven T cell responses in vivo. J. Immunol. 176: 1026–1035.
    OpenUrlAbstract/FREE Full Text
    1. Kim E. Y.,
    2. H. S. Teh
    . 2001. TNF type 2 receptor (p75) lowers the threshold of T cell activation. J. Immunol. 167: 6812–6820.
    OpenUrlAbstract/FREE Full Text
  31. ↵
    1. Kim E. Y.,
    2. H. S. Teh
    . 2004. Critical role of TNF receptor type-2 (p75) as a costimulator for IL-2 induction and T cell survival: a functional link to CD28. J. Immunol. 173: 4500–4509.
    OpenUrlAbstract/FREE Full Text
  32. ↵
    1. Morris S. C.,
    2. R. L. Cheek,
    3. P. L. Cohen,
    4. R. A. Eisenberg
    . 1990. Autoantibodies in chronic graft versus host result from cognate T-B interactions. J. Exp. Med. 171: 503–517.
    OpenUrlAbstract/FREE Full Text
  33. ↵
    1. Rickinson A. B.,
    2. D. J. Moss
    . 1997. Human cytotoxic T lymphocyte responses to Epstein-Barr virus infection. Annu. Rev. Immunol. 15: 405–431.
    OpenUrlCrossRefPubMed
  34. ↵
    1. Kasahara S.,
    2. K. Ando,
    3. K. Saito,
    4. K. Sekikawa,
    5. H. Ito,
    6. T. Ishikawa,
    7. H. Ohnishi,
    8. M. Seishima,
    9. S. Kakumu,
    10. H. Moriwaki
    . 2003. Lack of tumor necrosis factor alpha induces impaired proliferation of hepatitis B virus-specific cytotoxic T lymphocytes. J. Virol. 77: 2469–2476.
    OpenUrlAbstract/FREE Full Text
  35. ↵
    1. Kim S. Y.,
    2. D. H. Solomon
    . 2010. Tumor necrosis factor blockade and the risk of viral infection. Nat. Rev. Rheumatol. 6: 165–174.
    OpenUrlCrossRefPubMed
    1. Shale M. J.
    2009. The implications of anti-tumour necrosis factor therapy for viral infection in patients with inflammatory bowel disease. Br. Med. Bull. 92: 61–77.
    OpenUrlAbstract/FREE Full Text
  37. ↵
    1. Brunasso A. M.,
    2. M. Puntoni,
    3. A. Gulia,
    4. C. Massone
    . 2011. Safety of anti-tumour necrosis factor agents in patients with chronic hepatitis C infection: a systematic review. Rheumatology (Oxford) 50: 1700–1711.
    OpenUrlAbstract/FREE Full Text
  38. ↵
    1. Carroll M. B.,
    2. M. A. Forgione
    . 2010. Use of tumor necrosis factor alpha inhibitors in hepatitis B surface antigen-positive patients: a literature review and potential mechanisms of action. Clin. Rheumatol. 29: 1021–1029.
    OpenUrlCrossRefPubMed
  39. ↵
    1. Carroll M. B.,
    2. M. I. Bond
    . 2008. Use of tumor necrosis factor-alpha inhibitors in patients with chronic hepatitis B infection. Semin. Arthritis Rheum. 38: 208–217.
    OpenUrlCrossRefPubMed
  40. ↵
    1. Solomon D. H.,
    2. E. Mercer,
    3. A. Kavanaugh
    . 2012. Observational studies on the risk of cancer associated with tumor necrosis factor inhibitors in rheumatoid arthritis: a review of their methodologies and results. Arthritis Rheum. 64: 21–32.
    OpenUrlCrossRefPubMed
  41. ↵
    1. Askling J.,
    2. K. Fahrbach,
    3. B. Nordstrom,
    4. S. Ross,
    5. C. H. Schmid,
    6. D. Symmons
    . 2011. Cancer risk with tumor necrosis factor alpha (TNF) inhibitors: meta-analysis of randomized controlled trials of adalimumab, etanercept, and infliximab using patient level data. Pharmacoepidemiol. Drug Saf. 20: 119–130.
    OpenUrlCrossRefPubMed
  42. ↵
    1. Bongartz T.,
    2. A. J. Sutton,
    3. M. J. Sweeting,
    4. I. Buchan,
    5. E. L. Matteson,
    6. V. Montori
    . 2006. Anti-TNF antibody therapy in rheumatoid arthritis and the risk of serious infections and malignancies: systematic review and meta-analysis of rare harmful effects in randomized controlled trials. JAMA 295: 2275–2285.
    OpenUrlCrossRefPubMed
  43. ↵
    1. Nair B.,
    2. G. Raval,
    3. P. Mehta
    . 2007. TNF-alpha inhibitor etanercept and hematologic malignancies: report of a case and review of the literature. Am. J. Hematol. 82: 1022–1024.
    OpenUrlCrossRefPubMed
  44. ↵
    1. Leombruno J. P.,
    2. T. R. Einarson,
    3. E. C. Keystone
    . 2009. The safety of anti-tumour necrosis factor treatments in rheumatoid arthritis: meta and exposure-adjusted pooled analyses of serious adverse events. Ann. Rheum. Dis. 68: 1136–1145.
    OpenUrlAbstract/FREE Full Text
  45. ↵
    1. Diak P.,
    2. J. Siegel,
    3. L. La Grenade,
    4. L. Choi,
    5. S. Lemery,
    6. A. McMahon
    . 2010. Tumor necrosis factor alpha blockers and malignancy in children: forty-eight cases reported to the Food and Drug Administration. Arthritis Rheum. 62: 2517–2524.
    OpenUrlCrossRefPubMed
  46. ↵
    FDA. 2009. Cancer warnings required for TNF blockers. Available at: http://www.fda.gov/NewsEvents/Newsroom/PressAnnouncements/ucm175803.htm. Accessed: March 15, 2013.
  47. ↵
    1. Alessandri C.,
    2. R. Scrivo,
    3. F. R. Spinelli,
    4. F. Ceccarelli,
    5. L. Magrini,
    6. R. Priori,
    7. G. Valesini
    . 2007. Autoantibody production in anti-TNF-alpha-treated patients. Ann. N. Y. Acad. Sci. 1110: 319–329.
    OpenUrlCrossRefPubMed
  48. ↵
    1. de Saint Basile G.,
    2. A. Fischer
    . 2001. The role of cytotoxicity in lymphocyte homeostasis. Curr. Opin. Immunol. 13: 549–554.
    OpenUrlCrossRefPubMed
    1. Kägi D.,
    2. B. Odermatt,
    3. T. W. Mak
    . 1999. Homeostatic regulation of CD8+ T cells by perforin. Eur. J. Immunol. 29: 3262–3272.
    OpenUrlCrossRefPubMed
  50. ↵
    1. Badovinac V. P.,
    2. A. R. Tvinnereim,
    3. J. T. Harty
    . 2000. Regulation of antigen-specific CD8+ T cell homeostasis by perforin and interferon-gamma. Science 290: 1354–1358.
    OpenUrlAbstract/FREE Full Text
  51. ↵
    1. Soforo E.,
    2. M. Baumgartner,
    3. L. Francis,
    4. F. Allam,
    5. P. E. Phillips,
    6. A. Perl
    . 2010. Induction of systemic lupus erythematosus with tumor necrosis factor blockers. J. Rheumatol. 37: 204–205.
    OpenUrlFREE Full Text
  52. ↵
    1. Williams E. L.,
    2. S. Gadola,
    3. C. J. Edwards
    . 2009. Anti-TNF-induced lupus. Rheumatology (Oxford) 48: 716–720.
    OpenUrlAbstract/FREE Full Text
  53. ↵
    1. Peng S. L.,
    2. S. Fatenejad,
    3. J. Craft
    . 1996. Induction of nonpathologic, humoral autoimmunity in lupus-prone mice by a class II-restricted, transgenic alpha beta T cell. Separation of autoantigen-specific and -nonspecific help. J. Immunol. 157: 5225–5230.
    OpenUrlAbstract/FREE Full Text
    1. Mohan C.,
    2. S. Adams,
    3. V. Stanik,
    4. S. K. Datta
    . 1993. Nucleosome: a major immunogen for pathogenic autoantibody-inducing T cells of lupus. J. Exp. Med. 177: 1367–1381.
    OpenUrlAbstract/FREE Full Text
    1. Koshy M.,
    2. D. Berger,
    3. M. K. Crow
    . 1996. Increased expression of CD40 ligand on systemic lupus erythematosus lymphocytes. J. Clin. Invest. 98: 826–837.
    OpenUrlCrossRefPubMed
    1. Koh D. R.,
    2. A. Ho,
    3. A. Rahemtulla,
    4. W. P. Fung-Leung,
    5. H. Griesser,
    6. T. W. Mak
    . 1995. Murine lupus in MRL/lpr mice lacking CD4 or CD8 T cells. Eur. J. Immunol. 25: 2558–2562.
    OpenUrlCrossRefPubMed
  57. ↵
    1. Santoro T. J.,
    2. J. P. Portanova,
    3. B. L. Kotzin
    . 1988. The contribution of L3T4+ T cells to lymphoproliferation and autoantibody production in MRL-lpr/lpr mice. J. Exp. Med. 167: 1713–1718.
    OpenUrlAbstract/FREE Full Text
  58. ↵
    1. Fan G. C.,
    2. R. R. Singh
    . 2002. Vaccination with minigenes encoding V(H)-derived major histocompatibility complex class I-binding epitopes activates cytotoxic T cells that ablate autoantibody-producing B cells and inhibit lupus. J. Exp. Med. 196: 731–741.
    OpenUrlAbstract/FREE Full Text
    1. Karpouzas G. A.,
    2. A. La Cava,
    3. F. M. Ebling,
    4. R. R. Singh,
    5. B. H. Hahn
    . 2004. Differences between CD8+ T cells in lupus-prone (NZB x NZW) F1 mice and healthy (BALB/c x NZW) F1 mice may influence autoimmunity in the lupus model. Eur. J. Immunol. 34: 2489–2499.
    OpenUrlCrossRefPubMed
    1. Hahn B. H.,
    2. F. Ebling,
    3. R. R. Singh,
    4. R. P. Singh,
    5. G. Karpouzas,
    6. A. La Cava
    . 2005. Cellular and molecular mechanisms of regulation of autoantibody production in lupus. Ann. N. Y. Acad. Sci. 1051: 433–441.
    OpenUrlCrossRefPubMed
    1. Kim H. J.,
    2. B. Verbinnen,
    3. X. Tang,
    4. L. Lu,
    5. H. Cantor
    . 2010. Inhibition of follicular T-helper cells by CD8(+) regulatory T cells is essential for self tolerance. Nature 467: 328–332.
    OpenUrlCrossRefPubMed
    1. Kim H. J.,
    2. X. Wang,
    3. S. Radfar,
    4. T. J. Sproule,
    5. D. C. Roopenian,
    6. H. Cantor
    . 2011. CD8+ T regulatory cells express the Ly49 Class I MHC receptor and are defective in autoimmune prone B6-Yaa mice. Proc. Natl. Acad. Sci. USA 108: 2010–2015.
    OpenUrlAbstract/FREE Full Text
  63. ↵
    1. Zhang L.,
    2. A. M. Bertucci,
    3. R. Ramsey-Goldman,
    4. R. K. Burt,
    5. S. K. Datta
    . 2009. Regulatory T cell (Treg) subsets return in patients with refractory lupus following stem cell transplantation, and TGF-beta-producing CD8+ Treg cells are associated with immunological remission of lupus. J. Immunol. 183: 6346–6358.
    OpenUrlAbstract/FREE Full Text
  64. ↵
    1. Kollias G.,
    2. D. Kontoyiannis
    . 2002. Role of TNF/TNFR in autoimmunity: specific TNF receptor blockade may be advantageous to anti-TNF treatments. Cytokine Growth Factor Rev. 13: 315–321.
    OpenUrlCrossRefPubMed
  65. ↵
    1. Shibata H.,
    2. Y. Yoshioka,
    3. A. Ohkawa,
    4. K. Minowa,
    5. Y. Mukai,
    6. Y. Abe,
    7. M. Taniai,
    8. T. Nomura,
    9. H. Kayamuro,
    10. H. Nabeshi,
    11. et al
    . 2008. Creation and X-ray structure analysis of the tumor necrosis factor receptor-1-selective mutant of a tumor necrosis factor-alpha antagonist. J. Biol. Chem. 283: 998–1007.
    OpenUrlAbstract/FREE Full Text
  66. ↵
    1. Cabal-Hierro L.,
    2. P. S. Lazo
    . 2012. Signal transduction by tumor necrosis factor receptors. Cell. Signal. 24: 1297–1305.
    OpenUrlCrossRefPubMed
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