Abstract
Programmed death-1 (PD-1) inhibits T and B cell function upon ligand binding. PD-1 blockade revolutionized cancer treatment, and although numerous patients respond, some develop autoimmune-like symptoms or overt autoimmunity characterized by autoantibody production. PD-1 inhibition accelerates autoimmunity in mice, but its role in regulating germinal centers (GC) is controversial. To address the role of PD-1 in the GC reaction in type 1 diabetes, we used tetramers to phenotype insulin-specific CD4+ T and B cells in NOD mice. PD-1 or PD-L1 deficiency, and PD-1 but not PD-L2 blockade, unleashed insulin-specific T follicular helper CD4+ T cells and enhanced their survival. This was concomitant with an increase in GC B cells and augmented insulin autoantibody production. The effect of PD-1 blockade on the GC was reduced when mice were treated with a mAb targeting the insulin peptide:MHC class II complex. This work provides an explanation for autoimmune side effects following PD-1 pathway inhibition and suggests that targeting the self-peptide:MHC class II complex might limit autoimmunity arising from checkpoint blockade.
Introduction
Programmed death-1 (PD-1) is an inhibitory receptor expressed mainly by activated T and B lymphocytes (1). Upon binding to ligands PD-L1 and PD-L2, PD-1 recruits SHP2 phosphatase, which then dephosphorylates molecules downstream of the TCR and CD28, leading to a block in T cell effector function (2). Chronically stimulated T cells, such as those infiltrating a tumor or fighting a persistent viral infection, express high levels of PD-1 and have an exhausted phenotype characterized by diminished capability to produce cytokines, mediate target cell killing, and proliferate (2).
Blocking the PD-1/PD-L1 signaling pathway via mAbs can reinvigorate these exhausted T cells and kick-start antitumor immunity (2). Exciting results from clinical trials testing the efficacy of PD-1/PD-L1 checkpoint blockade led to Food and Drug Administration approvals to treat a wide variety of tumor types (3). However, a significant proportion of patients do not respond, and many develop immune-related adverse events (irAE), including overt autoimmunity such as type 1 diabetes (T1D) (3, 4). Interestingly, T1D occurs in 1–3% of patients receiving checkpoint therapy. Over 70% of these individuals have HLA alleles associated with T1D risk, suggesting that PD-1 may maintain islet tolerance in that subset of individuals (4). Increased B cell clonality and an increase in plasmablasts are predictive of grade 3 and 4 irAE after combined checkpoint blockade, but there are still no reliable biomarkers that can predict the development of naturally occurring or checkpoint blockade–induced autoimmunity (4). To develop biomarkers, we must first understand the mechanism by which PD-1 maintains tolerance to self-antigens.
Autoantibody production depends on cognate interactions between CD4+ T and B cells in the germinal center (GC) region of the lymph node (5). T follicular helper (TFH) cells express PD-1, ICOS, CXCR5, and Bcl-6 and provide IL-4, IL-21, and CD40-ligand stimulation to GC B cells, thus promoting Ab affinity maturation and somatic hypermutation (5). Increases in circulating TFH-like cells have been reported in patients with autoimmunity, suggesting that these cells may contribute to disease (5). T follicular regulatory (TFR) cells express PD-1, ICOS, CXCR5, CD25, Bcl-6, and Foxp3 and suppress TFH–B cell interactions to limit autoimmunity (6).
Given that the critical cellular players involved in the GC express PD-1 (5) and PD-L1 (7), it is likely that this pathway plays an important role in regulating CD4+ T cell–B cell cross-talk. Indeed, loss of PD-1/PD-L1 in C57BL/6 mice precipitates autoantibody production against dsDNA and a lupus-like disease (8), whereas PD-1 deficiency in BALB/c mice leads to autoantibody production against cardiac troponin I and autoimmune cardiomyopathy (9). Loss or blockade of PD-1 or PD-L1 in NOD mice accelerates T1D (reflecting irAE after checkpoint therapy); however, the effect on autoantibody production is unclear (10). Several studies showed that PD-1 deficiency or blockade impairs the outcome of the GC, resulting in fewer long-lived plasma cells (7) and lower-affinity Abs (11). Others demonstrated that PD-1 blockade enhances Ab production (12). In recent work, these contradictory findings were revisited, as PD-1 blockade was shown to enhance both the TFH and TFR CD4+ T cells, but their ratio determined the final outcome of the GC during foreign Ag immunization and in experimental autoimmune encephalomyelitis (13).
In this study, we investigated the role of PD-1 in regulating an Ag-specific, endogenous, polyclonal GC response in prediabetic NOD mice to further reconcile the role of PD-1 in regulating self-antigen–specific T/B interaction and autoantibody responses. Insulin is a critical autoantigen precipitating autoimmune T1D in NOD mice (14), and most insulin-specific CD4+ T cells in NOD mice respond to the insulin B10–23 (insB10–23) peptide (15, 16). Thus, we used insulin B10–23:MHC class II (MHCII) tetramers to track insulin-reactive CD4+ T cells, and generated insulin tetramers to track cognate insulin-specific B cells. We show that these two cell populations collaborate in prediabetic mice to give rise to insulin autoantibodies (IAA), as previously shown with T and BCR transgenic cells (17). We demonstrate that PD-1 or PD-L1 deficiency, as well as PD-1 but not PD-L2 blockade, impacted both insulin-specific TFH and TFR cells and increased their survival. Using mixed bone marrow chimeric mice, we also demonstrate that PD-1–deficient insulin-specific CD4+ T cells had an advantage in the GC over wild-type (WT), suggesting that T cell–intrinsic PD-1 restrains the GC. Furthermore, using an Ab that specifically disrupts TCR interactions with insulin peptide:MHCII complex, we show that the effects of PD-1 blockade on the GC are peptide:MHCII-dependent, as insulin-specific B cell numbers were significantly decreased in Ab-treated mice. We hypothesize that blocking self-peptide:MHCII complexes during checkpoint blockade may circumvent the autoimmune side effects of immunotherapy while still allowing antitumor immunity to reboot and eradicate the tumor.
Materials and Methods
Mice
Female NOD (Taconic), BALB/cJ (The Jackson Laboratory, Bar Harbor, ME), NOD.CD45.2, NOD.PD-1KO (16), NOD.PD-L1KO (16), NOD.Tg125 (18), C57BL/6.MD4.Rag1KO (19), and C57BL/6.I-Ag7 (B6.g7) mice (20) were housed in specific pathogen–free conditions. The Institutional Animal Care and Use Committee of the University of Minnesota approved all animal experiments.
In vivo PD-1 pathway blockade
IAA ELISA
For IAA ELISA, high-bind 96-well plates were coated with 10 μg/ml human insulin (I2643; Sigma-Aldrich, St. Louis, MO) and incubated overnight at 4°C. Plates were washed and blocked for 1 h with 5% BSA in 1× TBST, washed, and incubated with diluted serum for 1 h and subsequently with HRP-conjugated anti-mouse IgG for 1 h (1:2000 dilution, 405306; BioLegend, San Diego, CA). ABTS substrate (50-66-01; Kirkegaard & Perry Laboratories, Gaithersburg, MD) was added to each well for 10 min, and the plate was analyzed at 405 nm.
Insulin peptide:I-Ag7 tetramer development and validation
B cell tetramer development and validation
To detect Ag-specific B cells, we developed insulin B cell tetramers using described methods with several modifications (19). Briefly, human insulin (I2643; Sigma-Aldrich) or hen egg lysozyme (HEL; L6876; Sigma-Aldrich) was biotinylated using the EZ-Link Sulfo-NHS-LC Biotinylation Kit (21435; Thermo Fisher Scientific, Waltham, MA) at a biotin:protein ratio of 1:1. Excess biotin was removed using 3 ml of SpinOUT GT-600 size exclusion column (786-721; G-Biosciences, St. Louis, MO). Western blot was performed to determine the molar amount of biotinylated protein. Streptavidin (SA)–PE (PJRS27; Prozyme, Agilent Technologies, Santa Clara, CA) or SA–allophycocyanin (PJ27S; Prozyme) was added at a 4-fold molar excess to yield tetramers. Free unbiotinylated protein was eliminated from the tetramer solution by overnight dialysis in PBS using 20-kDa cutoff Slide-A-Lyzers (66005; Thermo Fisher Scientific). Tetramer specificity was confirmed by staining single-cell suspensions from pooled spleen and lymph nodes of NOD.Tg125 or C57BL/6.MD4.Rag1KO mice with insulin or HEL tetramers and preincubating samples with insulin prior to tetramer stain (Supplemental Fig. 1E).
Decoy tetramer development
To exclude B cells specific for the tetramer scaffold, we developed decoy reagents that encompassed SA, PE, or allophycocyanin and biotin (19, 28). To accomplish this, SA–PE and SA–allophycocyanin were conjugated to Alexa Fluor (AF) 647 (A20173) or DyLight 755 (84538; Thermo Fisher Scientific). SA–PE–AF647 and SA–allophycocyanin–DyLight 755 concentration was adjusted to 1 μM based on PE and allophycocyanin absorbance at 565 and 650 nm, respectively. Both reagents were biotinylated at a 6-fold molar excess of d-biotin.
Mixed bone marrow chimera setup
Five-week-old female NOD.CD45.2 mice were lethally irradiated (550 rad twice, 4 h apart) using an RS 2000 x-ray irradiator (Rad Source Technologies, Buford, GA). Between irradiations, bone marrow was isolated from femora and tibiae of donor NOD.CD45.1/2 and NOD.PD-1KO.CD45.1 mice. Mature T cells were depleted from the bone marrow cell preparations using biotinylated anti-CD90.2 Ab (53-2.1; Thermo Fisher Scientific) and anti-biotin magnetic beads (130-090-485; Miltenyi Biotec, Bergisch Gladbach, Germany) and LS columns (Miltenyi Biotec). Irradiated mice were reconstituted with a 50:50 mixture of WT and PD-1KO cells (10 × 106 cells per mouse) injected i.v. Antibiotic water with polymyxin B and neomycin was provided to mice for 3 wk. Mice were harvested for CD4+ T and B cell analyses 5 wk after reconstitution.
Flow cytometry
Single-cell suspensions were stained with PE- and allophycocyanin-conjugated insB10–23:I-Ag7 tetramers (p8E and p8G) and anti-CXCR5 for 1 h at 25°C in the presence of Fc block (2.4G2; Bio X Cell) followed by 30 min of incubation at 4°C with cell surface marker Abs, fixed and permeabilized (Tonbo Biosciences, San Diego, CA), and stained intracellularly (Supplemental Table I) (28). Abs, clones, and colors listed in Supplemental Table I are from BD Biosciences (San Jose, CA), BioLegend, Tonbo, and Thermo Fisher Scientific. Insulin-specific CD4+ T cells were defined as singlets, live, lineageneg (B220, CD8, CD11b, and CD11c), CD4+, and PE- and allophycocyanin-tetramer+ cells. To detect B cells, samples were stained with 10 nM decoy reagents for 10 min at 25°C followed by 10 nM tetramers for 30 min at 4°C (19). Tetramer+ cells were magnetically enriched from spleen and other lymph node cell suspensions (EasySep; STEMCELL Technologies, Vancouver, Canada), surface-stained, fixed, permeabilized, and stained with anti-Ig(H and L chain) (Supplemental Table I). Insulin-reactive B cells were defined as singlets, live, lineageneg (CD11c, F4/80, Gr1, and CD90.2), decoyneg, and PE- and allophycocyanin-tetramer+ cells. Flow cytometry was performed on BD Biosciences LSRII or Fortessa cytometers and analyzed using FlowJo software (FlowJo, Ashland, OR; BD Biosciences).
Anti-peptide:MHCII Ab generation
To generate a mAb that recognizes both insB10–23p8E:I-Ag7 and insB10–23p8G:I-Ag7 complexes, we modified a recently published method for generation of anti-peptide:MHCII Abs (28). Briefly, BALB/cJ mice were immunized s.c. with 25 μg of insB10–23p8E:I-Ag7 monomer in CFA (day 0) and boosted twice (days 28 and 42) with 25 μg of insB10–23p8G:I-Ag7 monomer in PBS i.v. Animals were harvested on day 45, and insB10–23p8E and insB10–23p8G:I-Ag7 tetramer+ B cells were enriched using anti-PE magnetic beads (18557; STEMCELL Technologies) and fused with SP2/0 mouse myeloma cells using the HY Hybridoma Cloning Kit method A (03800; STEMCELL Technologies). Individual colonies were hand-picked, grown, and tested for specificity as previously described (28). Supernatants reacting to both insB10–23p8E- and insB10–23p8G:I-Ag7 monomers, but not others, were considered cross-reactive to insulin peptide:I-Ag7, and corresponding hybridomas were further subcloned and isotyped. Clone Ins4G8 was considered specific for both insB10–23p8E and insB10–23p8G:I-Ag7, and the Ig V regions of the H and L chains were sequenced (U.S. Patent Application No. 15/952,965) using the SMARTer Mouse BCR Profiling Kit (634422; Clontech).
Ab staining of peptide-pulsed bone marrow–derived dendritic cells
Bone marrow was isolated from femora and humeri of donor NOD mice and cultured in complete RPMI 1640 medium supplemented with 10% FCS, 2-ME, HEPES, penicillin/streptomycin, nonessential amino acids, and recombinant murine GM-CSF (200 U/ml, 415Ml010; Biotechne) for 10 d. Nonadherent cells were collected by gentle pipetting, centrifuged at 300 × g for 5 min at 25°C, and resuspended in 3 ml of complete RPMI 1640 medium supplemented with 10 ng/ml recombinant murine GM-CSF, 1 μg/ml LPS (LPS from Escherichia coli J5(Rc); List Biological Laboratories, Campbell, CA), and 40 μM peptide. After overnight culture, cells were harvested, counted, and surface-stained with Ins4G8–AF647 and Abs against I-Ag7 (10-2.16–AF488), CD45.1 (PE Cy7, A20), CD11b (PE, M1/70; Tonbo Biosciences), CD11c (BUV395, HL3; BD Biosciences), and viability Ghost Dye (Violet 510; Tonbo Biosciences).
Immunofluorescence
Pancreata were harvested and frozen in OCT compound (Sakura Finetek, Torrance, CA) as previously described (29, 30). For insulitis quantification, 7-μm-thick pancreas sections were stained with guinea pig anti-insulin (A0564; Agilent, Santa Clara, CA) at 1:1000, anti-guinea pig IgG (H and L chain) (Cy5, 706175148; Jackson ImmunoResearch, West Grove, PA) at 1:1000, and anti-CD4 (AF488; GK1.5) and anti-CD8 (PE, 53–6.7; Thermo Fisher Scientific) at 1:100. Slides were mounted with Prolong Gold antifade reagent with DAPI (P36935; Thermo Fisher Scientific). Images were acquired on a Leica DM6000B epifluorescent microscope (Leica Microsystems, Buffalo Grove, IL). Islet scoring was performed by a blinded investigator using the following scale: 0, no insulitis; 1, peri-insulitis; 2, <25% of islet mass infiltrated; 3, <75% of islet mass infiltrated; 4, more than 75% of islet mass infiltrated (29).
In vivo Ins4G8 Ab validation
Ten-week-old NOD mice were immunized with 20 μg of peptide (p8E, p8G, or p63) and 50 μg of LPS on day 0 via i.p. route and treated with 500 μg of Ins4G8 or isotype control (IgG1; clone MOPC21; Bio X Cell) on days 0 and 4. Animals were harvested on day 7 for enumeration and phenotypic analysis of Ag-specific CD4+ T cells from the secondary lymphoid organs.
Ins4G8 and anti–PD-1 treatment
Statistics
Statistical analyses (paired or unpaired Student t test and one-way ANOVA with Tukey post hoc test) were performed in Prism 7 or 8.1 (GraphPad, La Jolla, CA). The p values lower than 0.05 were considered significant. Graphs show mean ± SEM.
Results
B cell tetramers reliably detect insulin-specific B cells
To detect insulin-binding B cells from peripheral blood of T1D patients, Smith et al. (31) used monomeric biotinylated insulin and magnetic enrichment. We reasoned that a tetrameric reagent would allow enhanced detection of lower-affinity autoreactive B cells as well as lower-avidity GC B cells, as these typically downregulate their BCR (32). In our approach, we also used decoy tetramers to exclude B cells specific for tetramer scaffold components including biotin, SA, or fluorochromes (19). Therefore, we generated fluorescent tetrameric insulin or HEL reagents and confirmed their specificity by staining single-cell suspensions from BCR transgenic C57BL/6.MD4.Rag1KO (33) and NOD.Tg125 mice (18) (Supplemental Fig. 1). Insulin but not HEL tetramers stained >92% of B cells from NOD.Tg125 mice, whereas HEL but not insulin tetramers stained >98% of B cells from C57BL/6.MD4.Rag1KO animals (Supplemental Fig. 1A–D). Next, we verified the specificity of tetramer-binding B cells by preincubating splenocytes from NOD.Tg125 with excess monomeric insulin prior to tetramer stain (Supplemental Fig. 1E). Incubation with monomeric insulin blocked insulin tetramer staining by >99% for NOD.Tg125 B cells, suggesting that the tetramer is detecting insulin-specific cells (Supplemental Fig. 1E). Priming NOD mice with insulin (25 μg) in CFA led to a significant ∼2-fold expansion of insulin tetramer+ B cells (p < 0.05) and a corresponding increase in IAA titer, further showing reagent specificity (Supplemental Fig. 1F, 1G). Having validated the tools to track insulin-specific B cells in a polyclonal setting, we proceeded to examine the effects of PD-1/PD-L1 deprivation on this cell population and on cognate CD4+ T cells in the polyclonal repertoire of prediabetic NOD mice.
Loss of PD-1 or PD-L1 unleashes insulin-specific CD4+ T and B cells
NOD.PD-1KO mice had significantly more insB10–23:I-Ag7 tetramer+ CD4+ T cells in the pancreatic lymph nodes (pLN) compared with age-matched NOD mice (p = 0.0097), consistent with our previous observations (16) (Fig. 1A). Additionally, PD-1 deficiency resulted in an increase in insulin-specific B cells in the pLN, pancreas, and other secondary lymphoid organs (spleen and nondraining lymph nodes [ndLNs]: axillary, brachial, cervical, and inguinal) and an increase in IAA (Fig. 1B, 1C). Similarly, NOD.PD-L1KO mice had increased insulin-specific CD4+ T cells in the pLN (p = 0.0012; Fig. 1D). PD-L1–deficient mice also had a larger population of insulin-specific B cells in the pLN and spleen and ndLNs but not in the pancreas (Fig. 1E). The increase in insulin-specific lymphocytes was accompanied with an increase in IAA in NOD.PD-L1KO animals (Fig. 1F). Importantly, deficiency in the PD-1/PD-L1 pathway did not affect CD4+ T and B cell populations specific for foreign Ags, as NOD and NOD.PD-L1KO mice had comparable numbers of HEL-specific CD4+ T cells (16) and HEL-specific B cells in the secondary lymphoid organs (Supplemental Fig. 1H). Taken together, these results support a model in which insulin-specific CD4+ T and B cell populations are held in check by the PD-1/PD-L1 pathway in autoimmune-prone mice.
Loss of PD-1 or PD-L1 results in increased numbers of insulin-specific CD4+ T and B cells and increased IAA production. (A and D) Number of insB10–23:I-Ag7 tetramer+ CD4+ T cells in the pLN of 5–7-wk-old NOD, PD-1KO, and PD-L1KO mice (n = 9–13 per group). (B and E) Number of insulin tetramer+ B cells in the spleen plus nondraining lymph nodes (Spl+nondLNs; which include the inguinal, axillary, brachial, and cervical lymph nodes) in NOD, PD-1KO, and PD-L1KO mice (n = 9–13 per group). (C and F) IAA ELISA OD measurements at 405 nm (n = 19–22). (A–F) Data are compiled from three to four independent experiments. Statistical significance was determined by unpaired Student t test. *p < 0.05, **p < 0.01.
Loss of PD-1 or PD-L1 results in increased numbers of insulin-specific CD4+ T and B cells and increased IAA production. (A and D) Number of insB10–23:I-Ag7 tetramer+ CD4+ T cells in the pLN of 5–7-wk-old NOD, PD-1KO, and PD-L1KO mice (n = 9–13 per group). (B and E) Number of insulin tetramer+ B cells in the spleen plus nondraining lymph nodes (Spl+nondLNs; which include the inguinal, axillary, brachial, and cervical lymph nodes) in NOD, PD-1KO, and PD-L1KO mice (n = 9–13 per group). (C and F) IAA ELISA OD measurements at 405 nm (n = 19–22). (A–F) Data are compiled from three to four independent experiments. Statistical significance was determined by unpaired Student t test. *p < 0.05, **p < 0.01.
Loss of PD-1 or PD-L1 results in increased insulin-specific follicular T cells and GC B cells
Ab production was shown to positively correlate with a high TFH/TFR CD4+ T ratio after foreign Ag immunization (13). To detect bulk and insulin-specific TFH and TFR cells, we used a well-established and previously reported gating strategy (13, 34). Follicular cells were identified as ICOS+CXCR5+. Within this gate, GITR and CD25 expression was used to identify TFR cells (GITR+CD25+). As shown in Supplemental Fig. 1I, >95% of ICOS+CXCR5+GITR+CD25+ cells also expressed Foxp3, confirming their regulatory lineage. When examining bulk CD4+ T cells in the pLN, we observed >4-fold increase in the TFH and TFR cells in NOD.PD-1KO and NOD.PD-L1KO mice compared with age-matched NOD mice (Supplemental Fig. 2A, 2B, respectively). Because CXCR5+CD8+ T cells have been described and shown to participate in the GC (35–37), we tested whether these cells were impacted by PD-1 deficiency. In both NOD and PD-1KO mice, <0.3% of CD8+ T cells in the pLN expressed CXCR5 (Supplemental Fig. 2F). We next evaluated insulin-specific follicular CD4+ T cells, identified as shown in Supplemental Fig. 1I. NOD.PD-1KO mice had a 2-fold increase in insulin-specific TFH cells and a trend toward increased TFR cells in the pLN compared with age-matched NOD mice (Fig. 2A). Similarly, NOD.PD-L1KO mice had increased insulin-specific TFH cells and increased TFR cells in the pLN (Fig. 2B). The increase in insulin-specific TFH cells in NOD.PD-1KO and NOD.PD-L1KO mice was accompanied with an increase in GC (GL7+) insulin-specific B cells in the pLN (Fig. 2C, 2D; p = 0.034), a more modest increase in the spleen and ndLNs (Fig. 2E, 2F; p = 0.057 and p = 0.084, respectively), and a trend toward increased isotype-switched insulin-specific B cells. The gating strategy for GC and switched cells is shown in Supplemental Fig. 1J. In summary, enhanced insulin-specific CD4+ T and B cell interactions in the absence of PD-1/PD-L1 led to increased output of isotype-switched insulin-specific B cells and likely contributed to accelerated T1D (10, 16, 22, 38).
Loss of PD-1 unleashes a larger anti-insulin GC reaction. (A and B) Number of TFH and TFR insB10–23:I-Ag7 tetramer+ CD4+ T cells in the pLN of 5–7-wk-old NOD, PD-1KO, and PD-L1KO mice (n = 9–13). (C and D) Number of GC (GL7+) and isotype-switched (swIg; IgM−IgD−) insulin tetramer+ B cells in the pLN (n = 8–13) and (E and F) spleen and nondraining lymph nodes (Spl+nondLNs) in NOD, PD-1KO, and PD-L1KO mice (n = 9–13). (A–F) Data are compiled from three to four independent experiments. Statistical significance was determined by unpaired Student t test. *p < 0.05, **p < 0.01.
Loss of PD-1 unleashes a larger anti-insulin GC reaction. (A and B) Number of TFH and TFR insB10–23:I-Ag7 tetramer+ CD4+ T cells in the pLN of 5–7-wk-old NOD, PD-1KO, and PD-L1KO mice (n = 9–13). (C and D) Number of GC (GL7+) and isotype-switched (swIg; IgM−IgD−) insulin tetramer+ B cells in the pLN (n = 8–13) and (E and F) spleen and nondraining lymph nodes (Spl+nondLNs) in NOD, PD-1KO, and PD-L1KO mice (n = 9–13). (A–F) Data are compiled from three to four independent experiments. Statistical significance was determined by unpaired Student t test. *p < 0.05, **p < 0.01.
Cell-intrinsic loss of PD-1 favors CD4+ T cell skewing to the follicular phenotype
We and others have previously shown that 5-wk-old NOD.PD-1KO and NOD.PD-L1KO mice have more severe islet infiltration compared with age-matched NOD mice and develop T1D more quickly (10, 16, 22, 38). We next performed mixed bone marrow chimeras to determine if cell-intrinsic PD-1/PD-L1 inhibition contributed to the enhanced GC reaction. This was also done to exclude Ag availability and systemic inflammation as potential factors contributing to increased insulin-specific CD4+ T and B cells. WT 5-wk-old NOD.CD45.2 mice were lethally irradiated and reconstituted with a 50:50 mixture of CD45.1+ PD-1KO and CD45.1/CD45.2+ WT cells (10 × 106 cells total). Five weeks after reconstitution, we assessed the number of insulin-specific CD4+ T cells in the thymus and the pLN. We did not detect differences in the number of donor-derived insulin-specific CD4+ T cells in the thymus; however, in the pLN, PD-1KO cells vastly outnumbered WT cells (Fig. 3A). This was also true for the bulk CD4+ T cells; however, the bias toward PD-1KO cells was higher among tetramer+ cells (Supplemental Fig. 2D). In the pLN, PD-1KO insulin-specific TFH and TFR cells outcompeted WT counterparts, as most detectable follicular cells originated from the PD-1KO donor (Fig. 3B). Similarly, PD-1KO insulin-specific B cells significantly outnumbered WT cells in the pLN and other secondary lymphoid organs but did not exhibit a strong bias toward GC or isotype-switched phenotype (Fig. 3C, 3D). Importantly, we did not observe a difference in the number of PD-1KO and WT HEL-specific B cells in the pLN (Supplemental Fig. 2E). These results indicate that PD-1 has a cell-intrinsic effect on insulin-specific CD4+ TFH and TFR cells and, to a lesser extent, on insulin-specific B cells but not on foreign Ag HEL-specific B cells.
PD-1–deficient insulin-specific CD4+ T cells outcompete WT cells in the GC. Five-week-old female NOD mice were irradiated and reconstituted with a 50:50 mixture (10 × 106 cells) of congenically disparate WT and PD-1KO bone marrow (depleted of mature T cells) (n = 8). Mice were harvested 5 wk after reconstitution. (A) Number of WT and PD-1KO insB10–23:I-Ag7 tetramer+ CD4+ T cells in the thymus and pLN of chimeric mice (n = 8). (B) Number of WT and PD-1KO TFH and TFR insB10–23:I-Ag7 CD4+ T cells in the pLN (n = 8). (C) Number of WT and PD-1KO insulin tetramer+ B cells in the pLN and spleen and nondraining lymph nodes (Spl+nondLNs) (n = 8). (D) Number of WT and PD-1KO GC (GL7+) and isotype-switched (swIg; IgM−IgD−) insulin-tetramer+ B cells in the pLN (n = 8). (A–D) Data are compiled from two independent experiments. Statistical significance was determined by paired Student t test. *p < 0.05, **p < 0.01, ***p < 0.0001.
PD-1–deficient insulin-specific CD4+ T cells outcompete WT cells in the GC. Five-week-old female NOD mice were irradiated and reconstituted with a 50:50 mixture (10 × 106 cells) of congenically disparate WT and PD-1KO bone marrow (depleted of mature T cells) (n = 8). Mice were harvested 5 wk after reconstitution. (A) Number of WT and PD-1KO insB10–23:I-Ag7 tetramer+ CD4+ T cells in the thymus and pLN of chimeric mice (n = 8). (B) Number of WT and PD-1KO TFH and TFR insB10–23:I-Ag7 CD4+ T cells in the pLN (n = 8). (C) Number of WT and PD-1KO insulin tetramer+ B cells in the pLN and spleen and nondraining lymph nodes (Spl+nondLNs) (n = 8). (D) Number of WT and PD-1KO GC (GL7+) and isotype-switched (swIg; IgM−IgD−) insulin-tetramer+ B cells in the pLN (n = 8). (A–D) Data are compiled from two independent experiments. Statistical significance was determined by paired Student t test. *p < 0.05, **p < 0.01, ***p < 0.0001.
PD-1 blockade increases insulin-specific TFH cells and insulin-specific GC B cell numbers and enhances IAA production in NOD mice
To better model the clinical scenario of irAE associated with checkpoint blockade, we examined the effects of PD-1 blockade on the GC reaction. Five-week-old NOD mice were treated with vehicle or anti–PD-1 (J43 clone) (10, 21) and were harvested on day 4, 10, 15, or 22 after treatment initiation. Anti–PD-1–treated mice had more insulin-specific CD4+ T cells in the pLN at day 10 but not at earlier or later time points (Fig. 4A). We analyzed the phenotype of insulin-specific CD4+ T cells and found that anti–PD-1–treated mice had significantly higher total (Supplemental Fig. 2C) and insulin-specific TFH cells in the pLN at day 10 (Fig. 4B). Previous work has shown that the relative ratio of TFH/TFR subsets determines the outcome of the GC reaction (13, 34, 39). Indeed, anti–PD-1 treatment led to an increase in the TFH/TFR ratio among tetramer+ cells on day 10 (Fig. 4C). To determine whether PD-1 blockade promoted increased TFH cell proliferation or survival, we measured Ki67 expression (Fig. 4D) and caspase-3 activation in follicular CD4+ T cells on day 10 post–treatment initiation. As shown in Fig. 4E, we detected significantly fewer caspase-3+ CD4+ TFH cells in mice treated with anti–PD-1 (p = 0.0218).
PD-1 blockade increases the number of insulin-specific CD4+ T and B cells and IAA production. (A) Number of insB10–23:I-Ag7 tetramer+ CD4+ T cells in the pLN of control and anti–PD-1–treated mice at day 10 post–treatment initiation (n = 6 per group). (B) Number of TFH and TFR insB10–23:I-Ag7 tetramer+ CD4+ T cells in the pLN of control and anti–PD-1–treated mice at day 10 post–treatment initiation (n = 6 per group). (C) Ratio of TFH and TFR insulin tetramer+ CD4+ T cells in the pLN of control and anti–PD-1–treated mice (n = 6–12 per group combined from two to three independent experiments). (D) Percent of Ki67+ TFH and TFR cells from pLN of control and anti–PD-1–treated mice at day 10 post–treatment start (n = 5 per group). (E) Percentage of cleaved caspase-3+ TFH and TFR cells from pLN of control and anti–PD-1–treated mice at day 10 post–treatment start (n = 5 per group). (F) Number of insulin tetramer+ B cells in the pLN at days 0, 10, 15, and 22 following treatment (n = 5–6 per time point). (G) Number of GC (GL7+) and isotype-switched (swIg; IgM−IgD−) insulin tetramer+ B cells in the pLN at day 10 posttreatment (n = 6 per group). (H) IAA ELISA OD measurements at 405 nm at days 0, 10, 15, and 20 following treatment (n = 5 per time point). (I) Insulitis score at days 0, 10, 15, and 20 postinjection (n = 3–6 per group). (J) Number of insulin tetramer+ B cells in the spleen and nondraining lymph nodes (Spl+nondLNs) in control and treated mice at day 10 posttreatment (n = 6). (K) Number of GC (GL7+) and swIg (IgM−IgD−) insulin tetramer+ B cells in Spl+nondLNs in control and treated mice at day 10 posttreatment (n = 6). Data are representative of two independent experiments. Statistical significance was determined by unpaired Student t test. *p < 0.05, **p < 0.01, ***p < 0.001.
PD-1 blockade increases the number of insulin-specific CD4+ T and B cells and IAA production. (A) Number of insB10–23:I-Ag7 tetramer+ CD4+ T cells in the pLN of control and anti–PD-1–treated mice at day 10 post–treatment initiation (n = 6 per group). (B) Number of TFH and TFR insB10–23:I-Ag7 tetramer+ CD4+ T cells in the pLN of control and anti–PD-1–treated mice at day 10 post–treatment initiation (n = 6 per group). (C) Ratio of TFH and TFR insulin tetramer+ CD4+ T cells in the pLN of control and anti–PD-1–treated mice (n = 6–12 per group combined from two to three independent experiments). (D) Percent of Ki67+ TFH and TFR cells from pLN of control and anti–PD-1–treated mice at day 10 post–treatment start (n = 5 per group). (E) Percentage of cleaved caspase-3+ TFH and TFR cells from pLN of control and anti–PD-1–treated mice at day 10 post–treatment start (n = 5 per group). (F) Number of insulin tetramer+ B cells in the pLN at days 0, 10, 15, and 22 following treatment (n = 5–6 per time point). (G) Number of GC (GL7+) and isotype-switched (swIg; IgM−IgD−) insulin tetramer+ B cells in the pLN at day 10 posttreatment (n = 6 per group). (H) IAA ELISA OD measurements at 405 nm at days 0, 10, 15, and 20 following treatment (n = 5 per time point). (I) Insulitis score at days 0, 10, 15, and 20 postinjection (n = 3–6 per group). (J) Number of insulin tetramer+ B cells in the spleen and nondraining lymph nodes (Spl+nondLNs) in control and treated mice at day 10 posttreatment (n = 6). (K) Number of GC (GL7+) and swIg (IgM−IgD−) insulin tetramer+ B cells in Spl+nondLNs in control and treated mice at day 10 posttreatment (n = 6). Data are representative of two independent experiments. Statistical significance was determined by unpaired Student t test. *p < 0.05, **p < 0.01, ***p < 0.001.
We next determined if the increased TFH/TFR ratio was associated with enhanced insulin-specific B cell frequency, IAA, and insulitis at day 10, day 15, and day 20/22 after anti–PD-1 injection. We detected significantly higher numbers of insulin-specific B cells in the pLN at all tested time points (Fig. 4F). This was concomitant with an increase in GC B cells (pLN, p = 0.0003; Fig. 4G). Importantly, PD-1 blockade led to de novo generation of IAA, as treated mice had significantly higher titers after anti–PD-1 at day 10, day 15, and day 20 compared to baseline and compared with vehicle-treated mice (Fig. 4H). Increased IAA production was accompanied with enhanced early islet infiltration at days 10 and 15 after anti–PD-1 administration (Fig. 4I). This increased frequency of insulin-specific B cells was mirrored in the spleen and ndLNs (Fig. 4J) and isotype-switched B cells (Fig. 4K), mirroring findings with 5-wk-old PD-1– and PD-L1–deficient mice. Collectively, these findings reflect clinical observations, in which some patients treated with checkpoint blockade rapidly developed autoantibodies and, in some cases, T1D (40).
Because PD-1 can also bind to PD-L2 (38) and GC B cells express PD-L2 (7), we examined whether PD-L2 blockade would have the same effect on insulin-specific CD4+ T and B cells. Five-week-old NOD mice were treated with vehicle or anti–PD-L2 (TY25 clone) five times and harvested 1 d after the last injection (41). PD-L2 blockade did not increase the number of total or follicular insulin-specific CD4+ T cells (Supplemental Fig. 3A). We also did not detect a change in insulin-specific B cells or in serum IAA (Supplemental Fig. 3B–D).
We next examined the effects of PD-1 pathway blockade on the GC formation in diabetes-resistant B6.g7 mice (41). Five-week-old B6.g7 mice were treated with vehicle or anti–PD-1 (J43 clone) five times and harvested 1 d after the last injection (day 10 post–treatment initiation). As shown in Supplemental Fig. 3E–H, PD-1 blockade did not promote a significant increase in insulin-specific TFH or TFR cells or lead to an increase in IAA production, and treated mice remained diabetes-free (16). These results suggest that the pre-existence of effector CD4+ T cells is required for PD-1 blockade to cause IAA and insulitis (16).
Effects of PD-1 blockade are reduced by treatment with a mAb targeting insB10–23:I-Ag7 complexes
IAA production in NOD mice depends on CD4+ T cell help (42), and specifically on CD4+ T cell recognition of tyrosine at position 16 in the insB10–23 peptide (14, 17, 43). Blocking TCR recognition of insB10–23 p8E and p8G peptide variants in the context of I-Ag7 with a mAb reduced IAA titers and significantly delayed T1D onset in NOD mice (43). We therefore reasoned that a similar strategy could be used to dampen the effects of PD-1 blockade on the GC reaction.
To generate a mAb against insB10–23p8E and insB10–23p8G:I-Ag7, we modified our recently developed approach (28). We immunized BALB/c mice with insB10–23p8E:I-Ag7 in CFA (day 0) and boosted them twice (days 28 and 42) with insB10–23p8G:I-Ag7 i.v. Ag-specific B cells were enriched using insB10–23p8E and insB10–23p8G:I-Ag7 tetramers and fused with SP2/0 mouse myeloma cells to create hybridomas. Out of 31 prescreened hybridomas, 3 were specific for insB10–23p8E:I-Ag7 and insB10–23p8G:I-Ag7, as determined by tetramer binding. We further purified a monoclonal IgG1 Ab from one hybridoma, termed Ins4G8.
To validate Ins4G8 specificity, we pulsed bone marrow–derived dendritic cells with various peptides and evaluated the ability of fluorescently labeled Ins4G8 to specifically bind to insB10–23p8E and insB10–23p8G:I-Ag7 complexes. As shown in Supplemental Fig. 3I, mean fluorescent intensity of Ins4G8 staining was at least 2-fold higher when bone marrow–derived dendritic cells were pulsed with insB10–23p8E or insB10–23p8G versus OVA323–339, BDC2.5 mimetope 1040-p63, or no peptide. To validate this Ab in vivo, we immunized NOD mice with 20 μg of 1040-p63, insB10–23p8E or insB10–23p8G peptide, and 50 μg of LPS, and treated them twice on days 0 and 4 with 500 μg of Ins4G8 or isotype (IgG1; MOPC21 clone) Ab. Seven days postimmunization, we enumerated cognate CD4+ T cells and found that Ins4G8 treatment did not impact the activation or proliferation of p63:I-Ag7–restricted CD4+ T cells but did reduce the activation, expansion, and expression of CD44 and PD-1 of both subsets of insulin-specific CD4+ T cells (Supplemental Fig. 3J, 3K).
Given the significant effect anti–PD-1 elicited on insulin-specific B cells and IAA (Fig. 4F, 4H), we next tested the effects of Ins4G8 and anti–PD-1 combination treatment on B cells. In the pLN, anti–PD-1 treatment led to a significant increase in total and insulin-reactive B cells, whereas Ins4G8 alone had no effect (Fig. 5A, 5B). Importantly, mice treated with both Ins4G8 and anti–PD-1 had fewer total and insulin-reactive B cells in the pLN compared with anti–PD-1–treated mice (Fig. 5A, 5B). Mice that received combination treatment also had a reduced degree of severely infiltrated islets compared with anti–PD-1–treated mice at day 10 post–treatment initiation (Fig. 5C, Supplemental Fig. 3L). However, combination treatment did not prevent the development of PD-1 blockade–induced T1D in NOD mice.
Blocking insB10–23:I-Ag7 complexes reduces the effects of anti–PD-1. (A) Number of total and (B) insulin tetramer+ B cells in the pLN of treated mice (n = 5 per group). (C) Frequency of infiltrated islets in mice treated five times with anti–PD-1 alone or anti–PD-1 with Ins4G8 Ab (n = 4–5 per group). Islet scoring: 0, no insulitis; 1, peri-insulitis; 2, <25% of islet mass infiltrated; 3, <75% of islet mass infiltrated; and 4, more than 75% of islet mass infiltrated (29). Data are representative of two independent experiments. One-way ANOVA with Tukey post hoc analysis was performed to determine statistical significance. **p < 0.01, ***p < 0.0005, ****p < 0.0001.
Blocking insB10–23:I-Ag7 complexes reduces the effects of anti–PD-1. (A) Number of total and (B) insulin tetramer+ B cells in the pLN of treated mice (n = 5 per group). (C) Frequency of infiltrated islets in mice treated five times with anti–PD-1 alone or anti–PD-1 with Ins4G8 Ab (n = 4–5 per group). Islet scoring: 0, no insulitis; 1, peri-insulitis; 2, <25% of islet mass infiltrated; 3, <75% of islet mass infiltrated; and 4, more than 75% of islet mass infiltrated (29). Data are representative of two independent experiments. One-way ANOVA with Tukey post hoc analysis was performed to determine statistical significance. **p < 0.01, ***p < 0.0005, ****p < 0.0001.
Discussion
The role of PD-1 in regulating the GC reaction has been a matter of intense debate. Whereas several studies reported lower Ab production, poor Ab affinity, and diversity after PD-1 blockade, others found that PD-1 deficiency or blockade augmented Ab titers in the context of immunization or infection (12, 13, 44). With reports of rapid autoantibody development after PD-1 inhibition in the clinic, there is an urgent need to determine how PD-1 regulates self-antigen–specific TFH and TFR CD4+ T cells and self-reactive cognate B cells. In this study, we used insulin peptide:MHCII tetramers to track insulin-reactive CD4+ T cells and developed a novel insulin tetramer reagent to track endogenous, polyclonal B cells in PD-1–sufficient and –deficient NOD mice.
In the current report, we determined that PD-1/PD-L1 deficiency or blockade increased the number of total and insulin-specific TFH and TFR cells in the pLN. Because we did not detect changes in foreign Ag-specific CD4+ T or B cells, we hypothesize that the overall increase in cellularity in the pLN of PD-1/PD-L1KO mice is due to other islet Ag-specific cell populations. PD-1/PD-L1 deficiency or blockade also promoted an increase in GC B cells and de novo IAA production, reflecting clinical observations (4, 40). We also observed an increase in insulin-specific TFH/TFR cell ratio after PD-1 blockade and speculate that this accounts for the increased IAA production, similarly to what has been described previously for bulk TFH/TFR ratio (13). Treating mice with anti–PD-1 allowed us to investigate the kinetics of the GC response. The GC reaction peaked at day 10 post–treatment initiation, correlating with increased insulitis and accelerated T1D onset (10). Importantly, we did not observe changes in insulin-specific CD4+ T or B cells or an increase in IAA following treatment with anti–PD-L2, suggesting that PD-1/PD-L1 but not PD-1/PD-L2 interactions control the GC. Furthermore, using mixed bone marrow chimeras, we showed that PD-1 on CD4+ T cells and, to a much lesser extent, on B cells regulates the GC and appears to limit CD4+ T cell survival. CD4+ T cell–intrinsic PD-1 might bind to PD-L1 on dendritic cells (34) and/or on B cells (7) and limit T cell dwell time (45) and, by extension, limit TCR signaling, leading to decreased cytokine production, survival, and reduced T cell help to B cells. In turn, CD4+ T cell–intrinsic PD-1 deficiency results in augmented T cell help and perhaps reduces the selective pressure for affinity maturation, thus leading to greater IAA production but lower overall IAA affinity (11, 44, 46). It is also possible that increased insulin-specific CD4+ T cell help licenses a small population of B cells to mutate toward autoreactivity (47, 48).
Recent work has challenged CD25 expression on TFR cells. When compared with nonfollicular regulatory T cells, TFR cells were shown to express lower CD25 (49, 50). However, when compared with TFH cells, which are CD25null, TFR cells are considered CD25+ (51). Further adding to this complexity, there appear to be two populations of TFR, one that is CD25+ and one that is CD25− (51). Our study is one of few that tracks endogenous, polyclonal, self-specific TFH and TFR cells using peptide:MHCII tetramers (39). To identify TFR cells, we used ICOS, CXCR5, GITR, and CD25 expression, as has been well established and previously reported (13, 34, 52, 53). By enumerating ICOS+CXCR5+GITR+CD25+ cells, we likely underestimated the number of TFR cells. However, in a subset of confirmatory experiments, we used Foxp3 to delineate TFH versus TFR cells and also observed increased ICOS+CXCR5+Foxp3− and ICOS+CXCR5+Foxp3+ cells (Supplemental Fig. 2G), thus validating our approach and the use of CD25 to delineate TFR cells.
Previous work has shown that CXCR5+CD8+ T cells infiltrate the GC and contribute to its regulation (35–37). This was largely documented in the context of chronic viral infection, namely HIV/SIV and EBV, in which the infected cells are CD4+ T cells and B cells, respectively, and as many as 20% of CD8+ T cells in peripheral lymph nodes express CXCR5. In this study, <0.3% of CD8+ T cells in the pLNs expressed CXCR5 (in NOD and PD-1KO mice; Supplemental Fig. 2F), suggesting that CD8+ T cells likely do not participate in the GC but rather contribute to β cell death in the pancreas (54). Autoreactive B cells, including those responding to insulin, have been shown to be polyreactive (31). Although we observed a significant expansion of insulin tetramer+ B cells and a corresponding increase in IAA titer after insulin immunization and after PD-1 inhibition, it is still possible that a fraction of insulin tetramer-binding cells may be polyreactive and not solely bind insulin under physiological conditions. Whether polyreactive insulin tetramer+ B cells are more or less pathogenic remains to be determined.
In this study, treatment with an Ab that blocks insulin peptide:MHCII complexes reduced the effects of PD-1 blockade on insulin-reactive B cell expansion but had subtle effects on islet inflammation in the short term and, ultimately, did not impact T1D incidence. This is likely due to other islet-specific CD4+ T cells being unaffected by Ins4G8 treatment or due to islet-specific CD8+ T cells, which would be unaffected by Ins4G8 as they are not MHCII-restricted. Recent work has shown that methyldopa has the ability to block insB10–23 but not viral peptide docking in the context of the DQ8 molecule in T1D patients (55). We speculate that Abs targeting multiple self-peptide:MHC class I and II complexes, compounds like methyldopa, Ag-coupled nanoparticles (56), or Ag-coupled apoptotic splenocytes (57) could limit the autoimmune side effects of checkpoint blockade in at-risk patients while still allowing antitumor immunity to reboot and eradicate tumors.
We have previously shown that PD-1 blockade in mice largely impacts effector but not anergic or naive insulin-specific CD4+ T cells (16, 30). The majority of insulin-specific CD4+ T cells in B6.g7 mice are naive; hence, it is not surprising that PD-1 blockade did not augment the GC reaction or promote T1D in this strain. Because most nondiabetic individuals also harbor naive insulin-specific CD4+ T cells (58), we hypothesize that only people with pre-existing effector self-specific CD4+ T cells are at risk for developing autoimmunity after PD-1 blockade. Phenotyping self-specific CD4+ T cells prior to checkpoint blockade and screening for HLA II or anti-islet autoantibodies might be a useful prognostic tool for the development of autoimmunity, and it might identify a window of opportunity for therapeutic interventions.
Acknowledgements
We thank Drs. J. W. Thomas, B. A. Binstadt, and M. K. Jenkins for kindly providing NOD.Tg125, C57BL/6.IAg7, and C57BL/6.MD4.Rag1KO mice, respectively.
Footnotes
This work was supported by National Institutes of Health (NIH) R01 AI106791 (to B.T.F.), U24 AI118635 (to B.T.F.), and P01 AI35296 (to B.T.F. and K.A.H.), Helmsley Charitable Trust 2018PG-T1D058 (to B.T.F.), Minnesota Partnership for Biotechnology and Medical Genomics MNP#18.01 (to B.T.F.), Juvenile Diabetes Research Foundation 3-2014-215 (to J.A.S.), Center for Autoimmune Disease Research Pilot Award UMF0020624 (to J.A.S.), a Frieda Martha Kunze Fellowship (to T.M.), NIH T35 AI118620 (to L.A.S.), R37 AI039560 (to K.A.H.), F30 AI131483 (to E.R.B.), and T32 AI007313 (to E.R.B. and C.G.T.), and University of Minnesota Foundation Fund 11724 (Diabetes Cure Research Using Immune Regulation and Tolerance).
The online version of this article contains supplemental material.
Abbreviations used in this article:
- AF
Alexa Fluor
- B6.g7
C57BL/6.I-Ag7
- GC
germinal center
- HEL
hen egg lysozyme
- IAA
insulin autoantibody
- insB10–23
insulinB10–23
- irAE
immune-related adverse event
- MHCII
MHC class II
- ndLN
nondraining lymph node
- PD-1
programmed death-1
- pLN
pancreatic lymph node
- SA
streptavidin
- T1D
type 1 diabetes
- TFH
T follicular helper
- TFR
T follicular regulatory
- WT
wild-type.
References
Disclosures
The authors have no financial conflicts of interest.