Cross-talk between 4-1BB and TLR1–TLR2 Signaling in CD8⁺ T Cells Regulates TLR2′s Costimulatory Effects

Toll-like receptors (TLR) play a central role in activating immune cells and clearing infectious entities by recognizing various molecules derived from microbial pathogens (1, 2). TLRs also bind a range of molecules released from dying or stressed cells (3). Myeloid differentiation factor-88 (MyD88) is an adapter molecule used by most TLRs and necessary for TLR-induced signaling. The activation of TLR-MyD88 signaling in CD4⁺ and CD8⁺ T cells prolongs their survival, augments T-cell expansion, and can enhance effector functions against tumors and infections (4–6). MyD88 signaling in T cells plays a vital role in T-cell survival even in the absence of exogenous TLR agonists (7). The mechanisms underlying the costimulatory effects of MyD88 signaling in T cells have yet to be defined.

A T cell's ability to proliferate and persist in vivo is heavily influenced by the stimulation of various costimulatory receptors, such as the tumor necrosis factor receptor (TNFR) members 4-1BB, CD70, LTA, OX-40, and GITR (8–10). 4-1BB signaling in T cells enhances proliferation and promotes T-cell survival by increasing IL2 and by upregulating the expression of antiapoptotic proteins (11, 12). 4-1BB plays an important role in generating a responsive memory T-cell population (13, 14). Preclinical models indicate that stimulating 4-1BB signaling on T cells or natural killer (NK) cells with agonistic antibodies elicits potent antitumor responses (10, 15–17). Clinical trials are examining the antitumor activity of 4-1BB agonists alone or when administered together with other anticancer agents such as PD-1 inhibitor in patients with melanoma, colorectal cancer, head and neck cancer, or relapsed/refractory B-cell non-Hodgkin lymphoma (NCT02179918, NCT00612664, NCT01775631, NCT02110082, and NCT01307267). Preliminary data thus far show partial responses in melanoma patients and an increased frequency of activated CD8⁺ T cells in circulation (9, 18).

To better understand how TLR-MyD88 signals enhanced CD8⁺ T-cell responses, we assessed changes in gene expression profiles of the CD8⁺ T-cell receptor transgenic “pmel” mice, which recognize the epitope gp100_25–33 expressed on melanoma cells, and MyD88^−/−pmel CD8⁺ T cells stimulated with or without the TLR1–TLR2 ligand (TLR1–TLR2L) Pam3CSK4. TLR1–TLR2 engagement on T cells increased the expression of 4-1BB, OX40, OX40L, GITR, and LTA. We found that 4-1BB played a central role in regulating the costimulatory effects of TLR1–TLR2 signaling in T cells. Combination therapy using an agonistic antibody to 4-1BB and TLR1–TLR2L enhanced antitumor responses in mice with established tumors. These studies offer insights into the molecular mechanisms through which TLR–TLR2 signals costimulate CD8⁺ T cells and highlight the biological significance of exploiting these signaling pathways to augment T-cell responses.

C57BL/6 and MyD88^−/− mice were purchased from Charles River, whileTLR2^−/− and pmel (B6.Cg-Thy1/Cy Tg(TcraTcrb)8Rest/J) mice were purchased from The Jackson Laboratory. The IRAK4 kinase dead mice were a generous gift from Dr. Stefanie Vogel (University of Maryland, Baltimore, MD) and 4-1BB^−/− mice from Dr. Lieping Chen (Yale University, New Haven, CT). 4-1BB^−/−pmel and MyD88^−/−pmel mice were obtained by crossing pmel with 4-1BB^−/− and MyD88^−/− mice and crossing offspring over nine generations. All the protocols were approved by the University of Maryland Institutional Animal Care and Use Committee.

Mouse T cells were cultured in RPMI 1640 (Invitrogen) medium with fetal bovine serum (Gemini), NEAA, penicillin, streptomycin, and gentamycin (Invitrogen). CD8⁺ T cells were initially sorted using the negative enrichment kit followed by positive selection (Invitrogen). In some experiments, pmel T cells were stimulated with MyD88^−/− splenocytes pulsed with mouse gp-100 peptide (10 ng/mL; EGSRNQDWL, GenScript Corp) at 37°C in 7% CO₂ at 1:5 T cell:antigen presentation cells (APC) ratio, whereas WT (C57BL/6) CD8⁺ T cells were stimulated with plate-bound anti-CD3ϵ (BD Biosciences) at 0.5 μg/mL, with or without the TLR1–TLR2 agonist Pam3CSK4 (1.5 μg/mL; InvivoGen). T-cell proliferation was determined by ³H-thymidine (1 μCi/well) uptake. For in vivo T-cell survival/expansion studies, CD8⁺ pmel T cells were purified by negative selection (Invitrogen) from CD90.1⁻CD45.2⁺ pmel and CD90.1⁺CD45.2⁺MyD88^−/− pmel mice and activated in vitro with mgp100 peptide-pulsed wild-type (WT) splenocytes and, 1 day later, were enriched by negative selection, mixed at a 1:1 ratio and i.v. injected into CD45.1⁺ mice. The number of transferred T cells was determined in different organs at different time points by staining cells with antibodies against CD8, CD45.2, and CD90.1. T cells were restimulated by vaccinating mice s.c. with 100 μg of hgp100 peptide admixed in Incomplete Freund's Adjuvant (IFA) and 10 μg of CpG-ODN on day 20 after T-cell transfer. IL1-agonist (IL1α, 10 ng/mL) was from Biolegend and the cIAP1/2 inhibitor GDC-0152 was purchased from Selleck Chemicals.

Purified CD8⁺ T cells from WT, MyD88^−/−, 4-1BB^−/−, TLR2^−/−, and IRAK4 KD mice were activated with anti-CD3ϵ (0.5 μg/mL), with or without Pam3CSK4 (0.5 μg/mL), 3H3 (1 μg/mL; rat IgG2a agonistic mAb to mouse 4-1BB, kindly provided by Dr. R.S. Mittler, Emory University, Atlanta, GA), IgG2a isotype control (1 μg/mL, 2A3; BioXcell).

In some experiments, the expression of various molecules on pmel T cells was determined by flow cytometry after activation and analyzed using FlowJo software (TreeStar) at the indicated time points. Antibodies against 4-1BB, OX-40, OX-40L, CD25, CD44, CD62L, CD132, CD127, and GITR used in flow cytometry were purchased from E-biosciences or BD Biosciences. Blocking antibodies to CD28 and GITR were purchased from BD Pharmingen, OX-40L from R&D Systems and 4-1BBL (19H3) from Dr. R.S. Mittler.

MyD88^−/− splenocytes underwent two rounds of CD8 T-cell depletion by CD8-positive selection. Splenocytes were pulsed with 10 ng/mL of mgp100 for 2 hours at 37°C followed by addition of purified pmel or MyD88^−/− pmel CD8 T cells with or without 10 μg/mL of Pam3CSK4. Seventy-two hours after stimulation, pmel or MyD88^−/− pmel CD8 T cells were selected by two rounds of negative enriched (Invitrogen, Dynal AS). Purity of CD3⁺CD8⁺ was found to exceed 97% in all three experiments conducted as determined by flow cytometry. Double-stranded cDNA (dsDNA) is made from 200 ng of total RNA using oligo-dT, reverse transcriptase, and DNA polymerase as recommended by the manufacturer (Ambion). In addition, the remaining RNA is digested with RNAse H. The dsDNA is purified trough columns and used as a template to generate biotin-labeled RNA (cRNA). The GEO Submission is GSE79475, and the NCBI tracking system number is 17812033.

For whole-genome gene expression cRNA (1.25 μg in 10 μL) is mixed with hybridization buffer and processed and analyzed, as recommended by Illumina, Inc. For data analysis, the samples were normalized using the cubic spline algorithm, assuming that the distribution of transcripts is similar, and the net expression was determined by subtracting the expression levels in the reference group (control samples) from the condition group (treatment samples). A differential expression is determined by comparison between the condition group and the reference group using an algorithm that assumes that target signal intensity (l) is normally distributed among replicates corresponding to some biological condition. This experiment was conducted three times, and RNA samples from each group were used for the gene array. Changes in mRNA transcript levels observed between groups were confirmed by real-time PCR.

CD8⁺T cells were activated in an anti-CD3ϵ (0.5 μg/mL) coated plate for 3 days, in the presence or absence of Pam3CSK4. mRNA was collected at 0, 2, 4, 12, and 16 hours after actinomycin D (Sigma) treatment (10 μg/mL). Reverse transcription was performed using the high-capacity reverse transcription kit from Invitrogen. 4-1BB and β-actin mRNA levels were measured by qPCR using the SYBR green mix from Biorad and the following primer sets synthesized by Integrated DNA Technologies (IDT): 4-1BB, CCTTCCTGAAATTCAGGTGCTGCAG and GCAGCACAATGACCACCACGTTG; b-actin, GAAAAGATGACCCAGATCATG and ATCTTCATGAGGTAGTCCGTC.

CD8 T cells (10⁶) were cultured in a 24-well plate coated with 1 μg/mL anti-CD3ϵ in RPMI, 10% FBS, 1% penicillin–streptomycin, 1% NEAA, and 0.1% gentamycin, for 24, 48, 72, and 96 hours in the presence or absence of Pam3CSK4 (1 μg/mL). At each time point, cells were collected and chromatin immunoprecipitation was done using the Magnetic ChIP kit from Thermo Scientific. Cross-linking of protein and DNA was performed by adding 37% formaldehyde to the culture solution to attain a final concentration of 1% formaldehyde. The cells were then lysed using lysis buffer provided. Chromatin was digested using the MNase enzyme, and the nuclear membrane was disrupted using short pulses of sonication. The resulting chromatin was incubated overnight with antibodies to p65, c-Jun, RNA polymerase, and Isotype control from Cell Signaling Technologies and to H3K4me3 from SABiosciences EpiTech ChIP antibody kit. The immune complexes were isolated using Protein A/G-coated magnetic beads and magnetic stand. The cross-links were reversed and protein digested using Proteinase K. The purified DNA was then used in PCR to detect the promoter regions of 4-1BB. The following primer sets were used: P1, ACGTCCTAATGGGCAACAGCTG, GTGAGGTTCTGCCGCTCCAC; P2, TTGGCCACCACACCATGC, CAAGGGTTTCAAGGTCCCC. Densitometry data were obtained using Image J.

For T-cell survival/expansion studies, CD8⁺ pmel T cells were purified by negative selection (InvivoGen) from CD90.1⁻CD45.2⁺ pmel and CD90.1⁺CD45.2⁺MyD88^−/− pmel mice and activated in vitro with mgp100 peptide-pulsed WT splenocytes and, 1 day later, were enriched by negative selection, mixed at a 1:1 ratio and i.v. injected into CD45.1⁺ mice. The number of transferred T cells was determined in different organs at different time points by staining cells with antibodies against CD8 (553033), CD45.2 (553772), and CD90.1 (557266; BD Biosciences). T cells were restimulated by vaccinating mice s.c. with 100 μg of hgp100 peptide admixed in IFA and 10 μg of CpG-ODN (tlrl-1826 InvivoGen) on day 20 after T-cell transfer. CD90.1⁻CD45.2⁺ pmel and CD90.1⁺CD45.2⁺ MyD88^−/− pmel CD8 T cells were activated using 1 μg/mL of mgp100_25–33 for 5 days and then mixed at 1:1 and injected i.v. into CD45.1⁺ mice. For tumor growth experiments, 10⁵ B16 F1 melanoma cells were injected s.c. into the flanks of C57B6 mice on day 0. B16 F1 melanoma cells were purchased from ATCC in 2013 and were used within 3 years of receiving them. We authenticated that our B16 cells expressed the melanin pigment by visual inspection and microscopic inspection and that B16 cells were recognized as target by pmel CD8⁺ T cells. On day 9, when tumors were detected, the mice were sublethally irradiated (400 rads) using a Cesium irradiator. After 24 hours, 5 × 10⁶–7 × 10⁶ pmel T cells, which were activated 3 days prior with 2.5 μg/mL of hgp100 and 100 U/mL of IL2 (589106 Biolegend), were injected i.v. into all groups. On days 10, 14, 17, and 20 after tumor injection, the mice were administered TLR1–TLR2 ligand (10 μg), 3H3 (100 μg, BE0239 BioXcell), TLR1–TLR2 ligand (10 μg), and 3H3 (100 μg), i.p. or i.v. Tumors were measured regularly with a caliper and mice were euthanized when tumor reached the set size limit or if the mice appeared moribund.

We examined the biological significance of MyD88 signaling in T-cell survival. An equal number of antigen-activated pmel (CD45.2⁺CD90.1⁻) and MyD88^−/−pmel (CD45.2⁺ CD90.1⁺) CD8⁺ T cells were coinjected into CD45.1⁺ mice, and T-cell numbers in various organs were compared at different time points after cell transfer. Figure 1A shows a representative dot plot of pmel and MyD88^−/−pmel CD8⁺ T cells 7 and 45 days after transfer. The number of pmel T cells and MyD88^−/−pmel T cells was similar on day 7 in the spleen, lymph node, and bone marrow (Fig. 1B). However, more pmel T cells were recovered from each of the different organs starting on day 14. T-cell expansion and contraction kinetics in response to antigen restimulation were also assessed 20 days after T-cell transfer. Pmel cells exhibited a greater potential to expand and persist than did MyD88^−/−pmel cells (Fig. 1C). These data indicate that MyD88 signaling in T cells provides a distinct survival and/or proliferative advantage over MyD88-deficient pmel T cells.

Microarray gene expression profiles were compared between TLR1–TLR2-stimulated and unstimulated pmel and MyD88^−/−pmel CD8⁺ T cells. Gene expression in the pmel, MyD88^−/−pmel, and MyD88^−/−pmel + TLR1–TLR2L groups was related, whereas TLR1–TLR2-stimulated pmel cells were only distantly related (Fig. 2A). TLR stimulation enhanced the expression of immune response genes and genes regulating apoptosis and survival, signal transduction pathways, and metabolism (Fig. 2B). Under the classification of T-cell costimulation, expression of the TNF family members Tnfsf9/OX40l, Tnfrsf9/4-1bb, Tnfrsf4/OX-40, Tnfrsf25/Dr3, Lta, and Tnfrsf18/Gitr was most prominently enhanced after TLR1–TLR2 stimulation. Expression of the genes was confirmed by quantitative RT-PCR (Fig. 2C). Surface expression of OX40L, 4-1BB, and GITR was increased in response to TLR1–TLR2 stimulation and correlated with the RNA transcript data (Fig. 2D). OX40 surface expression was moderately increased. In contrast, the expression of each of these proteins remained similar in TLR1–TLR2-stimulated and unstimulated MyD88^−/− T cells (Fig. 2D, bottom). These data highlight a previously unappreciated association between TLR–MyD88 signaling and TNFR family member expression in CD8⁺ T cells.

We examined the effect that blocking 4-1BB, OX-40, and GITR had on the costimulatory effects of TLR1–TLR2 stimulation. Pmel T cells were activated with mgp100-pulsed MyD88^−/− splenocytes in the presence or absence of TLR1–TLR2L, with or without blocking antibodies to OX-40L, 4-1BBL, GITR, or CD28. Blocking 4-1BB signaling reduced the costimulatory effects of TLR1–TLR2 ligand (Fig. 3A). Blocking GITR also modestly reduced TLR2 signals. In contrast, blocking OX40L or CD28 did not impair TLR1–TLR2 signaling, suggesting that although upregulated after TLR stimulation, these receptors do not appear to modulate the costimulatory effects of TLR1–TLR2 signaling in CD8⁺ T cells. Because the costimulatory effect of TLR1–TLR2 stimulation on T cells was influenced by 4-1BB expression, we assessed TLR1–TLR2L's proliferative effects on WT and 4-1BB^−/− CD8⁺ T cells. TLR1–TLR2L increased WT CD8⁺ T-cell proliferation but did not alter 4-1BB^−/− CD8⁺ T-cell proliferation (Fig. 3B). Likewise, TLR1–TLR2 engagement increased antigen-driven pmel T-cell expansion but did not augment 4-1BB^−/−pmel T-cell proliferation (Fig. 3C).

We examined whether differences in the expression of cytokine receptors could help explain the changes in T-cell expansion between 4-1BB signaling–competent and 4-1BB–deficient T cells. TLR1–TLR2 stimulation increased the expression of CD25 (the α-chain of the IL2 receptor), CD132 (common-γ chain), and CD127 (IL7 receptor subunit) on WT and 4-1BB^−/− T cells (Supplementary Fig. S1). WT and 4-1BB^−/−T cells also expressed more of the activation markers CD69, CD44, and CD62L and the costimulatory molecule CD28 after TLR stimulation. However, TLR1–TLR2L did not alter the expression of these molecules in MyD88^−/−CD8⁺ T cells. Thus, the costimulatory effect of TLR1–TLR2 engagement likely occurs via mechanisms that do not involve these proteins.

To further understand the association between 4-1BB and TLR1–TLR2 signaling, we investigated how the absence of MyD88 or TLR2 altered the costimulatory effects of 4-1BB signaling. Purified WT, MyD88^−/−, or TLR2^−/− CD8⁺ T cells were activated with -CD3ϵ antibody and agonistic 4-1BB antibody. Whereas wild-type and TLR2^−/− CD8⁺ T cells proliferated more in response to 4-1BB stimulation, MyD88^−/− T cells did not respond (Fig. 3D). These results suggest a potential role for MyD88 in 4-1BB–mediated costimulation. We tested whether the lack of 4-1BB on T cells might have a global impact and prevent T cells from responding to other common costimulatory signals. We observed that although 4-1BB^−/− T cells did not respond to TLR1–TLR2 L costimulation, they did respond to both CD28 and OX40 (Supplementary Fig. S2), indicating that 4-1BB deficiency does not globally affect all costimulatory signals. Instead, the contribution of 4-1BB to modulating TLR1–TLR2 costimulation is somewhat specific.

We assessed 4-1BB expression kinetics in WT and MyD88^−/− CD8⁺ T cells over a period of 5 days with or without TLR1–TLR2L. As reported by others (19), 4-1BB expression on CD8⁺ T cells increased upon T-cell activation, peaking between 2 and 3 days after activation, and returning to basal levels by day 4 in non-TLR–stimulated T cells. TLR1–TLR2L increased 4-1BB expression over non–TLR-stimulated WT T cells between days 2 and 5. However, 4-1BB expression on MyD88^−/− CD8⁺ T cells was not affected by TLR1–TLR2L (Fig. 4A). The increase in 4-1BB surface expression in response to TLR1–TLR2 stimulation was correlated with increased transcripts, as assessed by quantitative real-time PCR (Fig. 4B).

MyD88 signaling can enhance IFNγ mRNA stability (20), and we thus assessed whether TLR engagement increased 4-1BB transcripts by increasing mRNA stability. The decay rate of 4-1BB transcripts was the same in TLR-stimulated and unstimulated T cells (Fig. 4C, left), indicating that the increase in 4-1BB transcripts in TLR-stimulated T cells was not a result of enhanced mRNA stability, with both TLR-stimulated or unstimulated T cells maintaining more than 80% of the starting amount of 4-1BB mRNA. In contrast, the total mRNA dropped over 75% in both TLR-stimulated and unstimulated T cells (Fig. 4C, right).

We investigated whether TLR stimulation altered the amount of transcription factors bound to the 4-1BB promoter. The 4-1bb gene has three distinct promoter regions (PI, PII, and PIII), of which PI and PII each have an AP-1 and NF-κB binding site (21). We used a chromatin immunoprecipitation assay to assess NF-κB (p65) and AP-1(cJun) binding to each the PI and PII promoter regions at different time points after T-cell activation. Both p65 and c-Jun bound to both the PI and PII regions as early as 24 hours after T-cell activation (Fig. 4D). However, binding of p65 to the PI and PII regions in TLR-stimulated cells was increased 48 hours after T-cell activation (Fig. 4D). In TLR-stimulated cells c-Jun bound to the PI but not PII region. Because histone 3 lysine 4 trimethylation (H3K4me3) is associated with transcriptionally active genes, we also assessed whether TLR1–TLR2L regulated 4-1BB transcription by increasing H3K4me3 binding (Fig. 4E). H3K4me3 was undetectable on PI and PII in naïve T cells and was similar in TLR-stimulated and unstimulated CD8⁺ T cells. Thus, increased 4-1BB transcripts were primarily regulated by enhanced binding of p65 and c-Jun to the PI and PII.

We assessed whether combined stimulation of 4-1BB and TLR1–TLR2 signals enhanced CD8⁺ T-cell responses above each of these signals alone. Purified WT or 4-1BB^−/− CD8⁺ T cells were activated by plate bound CD3ϵ antibody and treated with agonistic 4-1BB antibody (3H3), TLR1–TLR2L, or both. We also treated cells with IL1α to rule out that the activation of MyD88 occurred via engagement of the IL1R. WT CD8⁺ T cells stimulated with 3H3 or TLR1–TLR2L proliferated more than untreated T cells or those treated with an isotype control antibody (Fig. 5A). Combining 3H3 with TLR1–TLR2L further enhanced T-cell proliferation over 3H3 or TLR1–TLR2 L (Fig. 5A). IL1α did not affect T-cell proliferation. Costimulatory effects of TLR1–TLR2L or 3H3 were not observed in 4-1BB^−/− CD8⁺ T cells (Fig. 5A). Engagement of TLR1–TLR2 on both WT and 4-1BB^−/− CD8⁺ T cells increased IFNγ production (Fig. 5B), suggesting that although the ability of TLR1–TLR2 stimulation to augment T proliferation is dependent on 4-1BB, it is not required to enhance IFNγ production.

We assessed the effects of the above-mentioned treatments on TLR2^−/−, MyD88^−/−, and IRAK4 kinase dead (IRAK4-KD) CD8⁺ T cells (Fig. 5C). As expected, none of these T cells responded to TLR2 stimulation. However, 4-1BB stimulation augmented TLR2^−/− CD8⁺ T-cell proliferation, but did not affect MyD88^−/− or IRAK4-KD CD8⁺ T-cell proliferation. These data highlight that the proliferative effects of 4-1BB signaling in CD8⁺ T cells depend to some degree on both MyD88 and IRAK-4. However, although the costimulatory effects of TLR1–TLR2 ligand depend on 4-1BB, the costimulatory effects of 4-1BB do not rely on the expression of TLR2.

To further explore a link between TLR2 and 4-1BB signaling, we evaluated the effects of inhibiting the cellular inhibitors of apoptosis 1 and 2 (c-IAP1/2). c-IAP1/2 function as positive regulators of the canonical NF-κB signaling pathway and are essential in both TLR2 and 4-1BB signaling (22, 23). Treating T cells with a small molecule inhibitor of c-IAP1 and c-IAP2, GDC-0152, impeded the effects of combination TLR1–TLR2 and 4-1BB stimulation and decreased the costimulatory effects of each separately (Supplementary Fig. S3). Thus, the costimulatory effects of TLR1–TLR2 and 4-1BB required TRAF signaling, and NF-κB activation was required for the costimulatory effects of combination TLR1–TLR2 and 4-1BB signaling.

Combined treatment of mice with TLR1–TLR2L and agonistic 4-1BB antibody provided greater antitumor activity than did pmel T cells or TLR1–TLR2L alone and to a smaller extent over mice treated with 4-1BB antibody alone (Fig. 5D, left). Transient tumor regression was observed over the course of 2 weeks in mice receiving TLR1–TLR2L plus anti–4-1BB or anti–4-1BB alone. Combinatorial TLR1–TLR2L and anti–4-1BB treatment induced stronger antitumor responses than did 3H3 alone. The tumors in 4 of 10 mice treated with TLR1–TLR2L plus anti–4-1BB extensively regressed, and three mice remained tumor free (Supplementary Fig. S4). Combination of TLR1–TLR2L with anti–4-1BB treatment also significantly enhanced survival compared with that of mice treated with 3H3 (P < 0.05) or TLR1–TLR2 L (P < 0.001). These results indicate that the costimulatory effects of TLR1–TLR2 signaling in CD8⁺ T cells are in part mediated by 4-1BB and can be exploited to augment antitumor immune response.

Stimulation of CD8⁺ T cells with TLR ligands leads to enhanced proliferation and effector functions (24). Studies from our group have shown that MyD88-deficient CD8⁺ T cells have an impaired ability to survive for a long time in vivo (25). How MyD88 within CD8⁺ T cells contributes to survival is undefined. The current studies demonstrate that TLR stimulation altered the expression of about 200 genes, including 4-1BB, OX-40, OX-40L, GITR, and DR3, which are known to costimulate activated CD8⁺ T cells (26–29). 4-1BB plays a crucial role in enhancing the function (30) and survival of CD8⁺ T cells (13, 14, 29, 31, 32). We present another vital role of 4-1BB as a mediator of the costimulatory effects of TLR1–TLR2 signals on CD8 T cells. TLR1–TLR2 engagement on T cells failed to costimulate T-cell expansion in the absence of 4-1BB or when blocking 4-1BB using antibodies. Although both TLR and 4-1BB signals promote CD8⁺ T-cell expansion and survival, combining these signals increased T-cell expansion over each individual treatment alone. The costimulatory effects of TLR1–TLR2L and the ability for 4-1BB blockade to inhibit the costimulatory properties of TLR1–TLR2 were heavily influenced by the amount of T-cell receptor (TCR) signal. Too high a concentration of CD3ϵ antibody or peptide-pulsed APCs bypassed the costimulatory effects of TLR1–TLR2 agonist or 4-1BB agonistic antibodies. Our analyses focused on CD8⁺ T-cells; however, the effects observed in CD8⁺ T cells might also occur in CD4⁺ T cells.

We demonstrated here that the combination of both TLR ligand and 4-1BB signals enhanced T-cell proliferation and IFNγ production in vitro and augmented antitumor responses in mice to a greater extent than either treatment alone. However, in vivo 4-1BB stimulation on different cells types can generate varied responses. For example, activating 4-1BB signals on DCs in a mouse model of HSV-1 (Herpes simplex virus-1) infection leads to the IFNγ-dependent accumulation of indoleamine-pyrrole 2,3-dioxygenase (IDO) and subsequent suppression of the immune response. Additionally, 4-1BB signals enhanced immunity against influenza in a CD8⁺ T cell–dependent manner (33). Depending on the in vivo model, 4-1BB can either promote or block CD4⁺CD25⁺ regulatory T-cell activity (34, 35). The transfer of 4-1BB⁺ CD8⁺ T cells generates an effective antitumor response when administered with agonistic 4-1BB antibody therapy. That 4-1BB stimulation can elicit varied immune responses highlights a potential advantage to the targeting of therapy to specific cell subsets.

These studies reveal that the costimulatory effects of TLR1–TLR2 signaling in CD8⁺ T-cell expansion are in part mediated by 4-1BB, that 4-1BB signaling in T cells depends in part on the presence of MyD88, and combination therapy using TLR ligand and agonistic 41BB antibodies can be exploited to enhance antitumor immune response. The proposed model through which the T-cell receptor, TLR2, and 4-1BB signaling are believed to interact is shown in Supplementary Fig. S5.

No potential conflicts of interest were disclosed.

Conception and design: A.M. Joseph, E. Davila

Development of methodology: R. Srivastava, J. Zabaleta, E. Davila

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): A.M. Joseph, J. Zabaleta

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): A.M. Joseph, R. Srivastava, J. Zabaleta

Writing, review, and/or revision of the manuscript: A.M. Joseph, R. Srivastava, J. Zabaleta, E. Davila

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): R. Srivastava

This study was supported by the R01CA140917 (E. Davila), University of Maryland, Marlene and Stewart Greenebaum Cancer (E. Davila), P30CA134274 (E. Davila), and the T32 AI007540 (A.M. Joseph). J. Zabaleta has been partially supported by grants from the National Institute of General Medical Sciences (NIGMS P20GM103501, P30GM114732, and U54GM104940-01), and the National Institute on Minority Health and Health Disparities (NIMHD P20MD004817 and U54MD008176-01).

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.