CD4+ T helper cells are essential for control of malaria in mouse models of the disease6. We show that the frequencies of activated T helper cells increased in mice with malaria and in a cohort of Malian children (n = 15; Supplementary Table 1) diagnosed with febrile malaria (Supplementary Figs. 1c and 2). T helper cell depletion after established infection with the normally nonlethal Plasmodium yoelii 17XNL (P. yoelii) resulted in uncontrolled parasitemia and death in infected mice (Supplementary Fig. 1a,b). Unlike CD4+ T cell responses to most other infections in mice (for example, Listeria monocytogenes), expansion of pathogen-specific T helper cells (defined as CD49d+CD11ahiCD4+)7 in response to P. yoelii infection is distinctly biphasic. Specifically, the frequencies and total numbers of pathogen-specific T helper cells increased after P. yoelii infection and then temporarily fell or plateaued before rising again prior to parasite clearance (Supplementary Fig. 1d–f). Although the mechanisms underlying this unique hiatus in T helper cell expansion were unknown, we speculated that it was due to the widely acknowledged immunosuppression that occurs in blood-stage malaria5. This notion was reinforced by our observation of increased numbers and frequencies of FOXP3+CD4+ Treg cells, a key immunosuppressive cell population, during the blood stage of malaria in humans and mice (Fig. 1a–d). We observed that higher parasite densities were associated with higher frequencies of Treg cells in humans; also, chloroquine treatment to decrease parasitemia reduced Treg cell frequencies and increased T helper cell frequencies in mice (Supplementary Fig. 3a–c). The role of Treg cells in malaria remains controversial5,8,9: some independent studies suggest that Treg cells suppress protection10,11,12, while others imply that they enhance it13,14. Importantly, these studies manipulated Treg cells (most of which express CD25) before or shortly after Plasmodium infection using anti-CD25 antibody–mediated depletion10,11,13,14,15 or the (more precise) Foxp3–diptheria toxin receptor (DTR) system16,17. Although variations in approaches may underlie these inconsistencies, the timing of Treg cell targeting could be a critical consideration for understanding malaria. Here we observed that expansion of Treg cells in P. yoelii–infected mice preceded or coincided with the hiatus in T helper cell responses, suggesting a causal relationship beginning 10 days post infection (d.p.i.). To test this proposition, we depleted both circulating and lymphoid18,19,20 Treg cells in P. yoelii–infected Foxp3-DTR (Supplementary Fig. 4a,b) and C57BL/6 (Supplementary Fig. 5a) mice with diphtheria toxin21 and anti-CD25 antibody10, respectively, beginning at 9 d.p.i. Treg depletion interrupted the hiatus in T helper cell responses, restored expansion of Plasmodium-specific T helper cells and substantially accelerated control of P. yoelii infection (Fig. 1e,f and Supplementary Fig. 5b,c). In contrast, Treg depletion at 0 and 2 d.p.i. in Foxp3-DTR mice resulted in the death of P. yoelii–infected mice (Supplementary Fig. 4c).

Figure 1: Treg cells expand and modulate T helper cell responses and immunity to malaria.
figure 1

(a) Longitudinal frequencies of FOXP3+ Treg cells among the total CD4+ T cell population in peripheral blood mononuclear cells (PBMCs) from a cohort of children in Mali (Supplementary Table 1) before, during, and after infection with acute febrile malaria. Each connected line indicates a unique subject. (b) Kinetics of activated (CD49d+CD11ahi) T helper or FOXP3+ Treg cell frequencies among the total CD4+ T cell population in circulation in P. yoelii–infected C57BL/6 mice. (c,d) Absolute numbers of FOXP3+CD4+ Treg cells in spleen (c) and lymph nodes (LN) (d) at the indicated time points in P. yoelii–infected C57BL/6 mice. (e,f) Frequencies of activated CD4+ T helper cells among the total CD4+ T cell population in circulation (e) and parasitemia (f) at various time points following infection with P. yoelii in C57BL/6 or Foxp3-DTR mice treated with diphtheria toxin (Rx DT) or PBS (Rx PBS) at 9 and 11 d.p.i., as indicated by the pink label below the x axis. All experimental data represent one of at least three separate experiments with five mice per group and are presented as mean ± s.e.m. Statistical analysis was performed by comparing the indicated groups using one-way ANOVA with Bonferroni correction in a (F3,52 = 18.89) or the groups of Foxp3-DTR mice treated with diphtheria toxin or PBS and C57BL/6 mice treated with diptheria toxin using two-way ANOVA with Tukey's correction in e (F2,60 = 28.6) and f (F2,53 = 62.17). **P ≤ 0.01.

To further investigate the opposing contributions of Treg and T helper cells in the control of parasitemia during the hiatus in T helper cell responses in P. yoelii–infected mice, we took advantage of the differential expression of high- and low-affinity IL-2 receptors by these cell populations. Specifically, beginning at 9 d.p.i., we treated mice with either IL-2–JES6 antibody complexes that signal through the high-affinity IL-2 receptor CD25 and amplify CD25+ Treg cells or IL-2–S4B6 antibody complexes that selectively expand pathogen-specific T helper cells by signaling through the low-affinity IL-2/IL-15 receptor, which is expressed by activated T helper cells22. Increasing the frequencies of Treg cells further dampened pathogen-specific T helper cell responses, resulting in higher parasitemia and death, whereas increasing pathogen-specific T helper cell frequencies resulted in better control of infection (Supplementary Fig. 5d–f). Together, these data suggest that Treg cells suppress T helper cell responses during a critical window of time in blood-stage malaria, compromising control of acute infection.

There are two major mechanisms through which Treg cells counter T helper cell responses in the context of infection: IL-10-mediated inhibition and CTLA-4-mediated repression of co-stimulation by antigen-presenting cells (APCs)23. In mouse malaria, Treg cells transcriptionally upregulate both IL-10 and CTLA-4 (ref. 16). In accordance with another study16, blocking IL-10 at 9 d.p.i. failed to alter the T helper cell response and the course of parasitemia (Supplementary Fig. 6). However, at 9 d.p.i. in P. yoelii–infected mice, Treg cells exhibited enhanced upregulation of CTLA-4 in comparison to Treg cells from mice with acute infection with influenza or vaccinia virus (Fig. 2a,b). Additionally, elevated amounts of soluble CTLA-4, potentially cleaved from the surface of T cells, were detectable in blood plasma and spleen lysates throughout the course of P. yoelii infection (Fig. 2c). Of note, of all the Treg cells, the percentage expressing CTLA-4 in P. yoelii–infected mice was substantially higher than the percentage of T helper cells that had detectable CTLA-4 expression (Fig. 2d)16. In accordance with these mouse data, longitudinal analyses of samples from humans without malaria obtained at the end of the dry season in Mali and samples taken subsequently at diagnosis of febrile malaria showed increased frequencies of circulating CTLA-4+ T helper cells and CTLA-4+ Treg cells in blood-stage malaria; these frequencies returned to preinfection levels after treatment with an antimalarial drug (Fig. 2e,f and Supplementary Fig. 7a). Febrile malaria in humans was associated with higher frequencies of Treg cells positive for Helios (a marker of superior suppressive function)24,25, Helios+CTLA-4+ Treg cells, and CTLA-4+ follicular Treg (TFR) cells26,27 (Supplementary Fig. 7b–d). Together, these data suggest that Treg cells may modulate T helper cells, and possibly humoral immunity to blood-stage malaria, through CTLA-4.

Figure 2: CTLA-4 expression is enhanced in malaria and is integral to TFH–B–TFR cell interactions in GCs.
figure 2

(a) Representative histograms quantifying CTLA-4 expression in FOXP3+ Treg cells in C57BL/6 mice infected with influenza A virus (IAV), vaccinia virus (VacV), or P. yoelii at 9 d.p.i. Closed histograms represent isotype controls. Inset gates and numbers correspond to the proportions of CTLA-4+FOXP3+ Treg cells. (b) Summaries of the proportions of CTLA-4+FOXP3+ Treg cells shown in a. Boxes and whiskers depict the limits of the data distribution in each group, with upper and lower whiskers representing the range of data distribution, the box perimeters representing its 25th and 75th percentiles, and the midline showing median of the distribution. (c,d) Kinetics of CTLA-4 expression in spleen or serum (c) and of the proportion of splenic FOXP3+ Treg cells or T helper cells expressing CTLA-4 relative to the total Treg cell or T helper cell population, respectively (d), during the T helper cell hiatus after P. yoelii infection in C57BL/6 mice. The dashed line in c is the threshold of detection. (e,f) Proportions of T helper (e) and Treg (f) cells expressing CTLA-4 in a cohort of children in Mali before, during, and after acute febrile malaria as compared to healthy controls (US control). (g) Representative pseudocolored images of fluorescently labeled sections from the spleen (left, middle) and lymph node (right) of a C57BL/6 mouse with a resolving P. yoelii infection (21–27 d.p.i.). TFH–B–TFR cell clusters in the nascent GCs are enclosed by dashed circles in the left panel; a magnified view of an encircled GC from the left panel is shown in the middle panel. A single B cell and CTLA-4+ TFR cell in apposition are shown in the right panel. Markers and their colors are indicated in the figure. All experimental data represent one of at least three experiments with five mice per group and are presented as mean ± s.e.m. Confocal microscopy images represent at least five regions observed in nine sections examined in three separate mice from three separate experiments. Statistical analysis was performed by comparing the indicated groups using two-way ANOVA with Bonferroni correction in b (F2,14 = 25.08), two-tailed Student's t-test in d, or one-way ANOVA with Tukey's correction in e (F3,52 = 18.70) and f (F3,52 = 18.89). *P ≤ 0.05, **P ≤ 0.01.

Humoral immunity depends on efficient TFH–B cell cooperation in secondary lymphoid organs and is perhaps the most important component of the acquired immunity that controls blood-stage malaria28,29. To examine the potential of Treg cells and CTLA-4 to interfere with humoral immunity against malaria, we examined their relationships with the TFH–B cell interactions in germinal centers (GCs) by confocal and intravital microscopy in mice. Within the GCs in secondary lymphoid organs, CD4+ T helper cells and GL-7+B220+ B cells formed discrete clusters of interaction after P. yoelii infection (Supplementary Video 1). These clusters were composed of CTLA-4+ TFR and TFH cells30 in close apposition with GC plasmablasts or B cells (Fig. 2g and Supplementary Video 2). Expression of neuropilin-1 (NRP-1) or FOXP3 distinguished TFR cells from TFH cells31. In the context of infection, CTLA-4 receptors expressed on T cells bind to B7 ligands on APCs and limit immune responses by competitive inhibition of B7–CD28 co-stimulatory interactions32. As B cells are the primary APCs that sustain TFH cell responses33 and dictate protective antibody responses in malaria, we investigated whether GC B cells interacted with CTLA-4 during the course of P. yoelii infection. In accordance with this notion, CTLA-4+ TFR cells directly associated with GC B cells (Fig. 2g and Supplementary Video 3) during P. yoelii infection, and CTLA-4 was detectable at the TFR–B cell interface (Supplementary Video 4). Moreover, intravital imaging showed that individual TFR cells transiently interacted with multiple B cells in GCs (Supplementary Video 5), suggesting an explanation for how the relatively few TFR cells could effectively modulate the GC reaction. We failed to detect other potential CTLA-4-driven suppressive mechanisms of Treg cells32, including induced idoleamine 2,3-dioxygenase (IDO) production (in vitro) in B cells or discernable transendocytosis of B7 molecules (in vitro or in vivo) from B cells after P. yoelii infection (data not shown). These observations suggest that the CTLA-4 expressed on or released from TFR cells might directly bind B7 ligands on the surface of GC B cells, restricting productive co-stimulation of TFH cells and limiting production of antibody-secreting plasma cells and memory B cells. Thus, blocking CTLA-4–B7 interactions might augment humoral immunity and clearance of blood-stage malaria.

Our precise definitions of Treg cell kinetics, CTLA-4 expression dynamics, and the timing of GC reactions during the course of P. yoelii infection suggest that CTLA-4–B7 interactions may be meaningfully targeted during the hiatus in T helper cell expansion to improve immune responses and parasite clearance. Hence, P. yoelii–infected C57BL/6 mice were treated with CTLA-4-blocking (anti-CTLA-4) or IgG control antibodies at the onset of the hiatus in the expansion of pathogen-specific T helper cells (Fig. 3a). Similar to Treg cell depletion, therapeutic blockade of CTLA-4 truncated the hiatus and enhanced the total numbers of CD4+ T cells, pathogen-specific T helper cells, follicular CD4+ T cells, and TFH cells in the spleen as compared to treatment with IgG control antibodies (Fig. 3b–e). Hypothetically, CTLA-4 blockade could target Treg cells, T helper cells, or both (Fig. 2d). However, CTLA-4 blockade failed to further improve the T helper cell response in Treg cell–depleted mice, suggesting only a minor role for CTLA-4 expressed on T helper cells (Supplementary Fig. 8). Unlike in tumor models34, anti-CTLA-4 treatment during the course of malaria did not deplete Treg cells or TFR cells in the spleen (Fig. 3f and Supplementary Fig. 9). We observed a corresponding increase in the total numbers of splenic B cells, plasmablasts, GC B cells, GC plasmablasts, and P. yoelii–specific, protective antibody titers in serum35 of anti-CTLA-4-treated mice (Fig. 3g–k). Inhibiting GC B cell formation with anti-CD40L treatment33 prevented the revival of T helper cell responses after CTLA-4 blockade, indicating that Treg cell interaction with GC B cells likely mediated the repression of T helper cell responses (Supplementary Fig. 10), although effects on other APCs remain possible. CTLA-4 blockade resulted in considerably reduced splenomegaly (Fig. 3l), a hallmark of P. yoelii infection, and improved splenic architecture, with distinct T cell zones, B cell follicles, and GC reactions visible shortly after treatment (Fig. 3m). Therapeutic blockade of CTLA-4 dramatically accelerated control of P. yoelii infection in C57BL/6 and BALB/c mice compared to control IgG treated mice and partially rescued BALB/c mice (40% survival) from lethal Plasmodium berghei ANKA infection (Fig. 4a–c). However, CTLA-4 blockade before or after the critical window of expansion of Treg cells did not resolve the T helper cell hiatus or accelerate control of P. yoelii infection (Supplementary Fig. 11), in accordance with some previous results17,36. We previously showed that blockade of programmed cell death 1 protein (PD-1) and lymphocyte-activation gene 3–encoded protein (LAG-3) signaling beginning at 14 d.p.i., during the post-hiatus revival of T helper cell responses, resulted in accelerated clearance of P. yoelii infection in mice7. In contrast, blocking PD-1 and LAG-3 signaling starting at 9 d.p.i., during the Treg cell–mediated interruption in T helper cell responses, did not improve immunity or parasite clearance, indicating a minimal contribution of these pathways to the dampening of immune responses during this critical interval (Supplementary Fig. 12). Of note, stimulation of OX40 signaling starting at 7 d.p.i. can also improve immunity against blood-stage P. yoelii by enhancing T helper cell responses37. Together, these results reinforce the notion that immunomodulation in blood-stage malaria is based on multiple molecular pathways that may be dominant during discrete time windows over the course of the infection.

Figure 3: CTLA-4 blockade enhances CD4+ T cell and B cell responses, GC reaction, and antibody titers after P. yoelii infection.
figure 3

(a) A schematic of therapeutic blockade in P. yoelii–infected C57BL/6 mice; i.v., intravenous injection; i.p., intraperitoneal injection. (bk) Total numbers of CD4+ T helper (b), Plasmodium-specific CD49d+CD11ahiCD4+ T helper (c), CXCR5+ICOS+PD-1+CD4+ follicular T (d), CXCR5+ICOS+PD-1+FOXP3CD4+ TFH (e), and CXCR5+ICOS+PD-1+FOXP3+CD4+ TFR (f) cells, CD19+B220+ B cells (g), CD138+IgDCD19+B220+ plasmablasts (h), CD95+GL7+CD19+B220+ GC B cells (i), and CD95+GL7+CD138+IgDCD19+B220+ GC plasmablasts (j) in spleen at various time points and the relative titers of P. yoelii merozoite surface protein (MSP1–19)-specific antibody in serum at 18 d (k) following P. yoelii infection in C57BL/6 mice with or without CTLA-4 blockade as depicted in a. Data are presented as mean ± s.e.m. at each time point or serum dilution and represent one of three separate experiments, each with at least five mice per group. Statistical analysis was performed by comparing the treatment and control groups at the indicated time points in bj or serum dilutions in k with two-tailed Student's t-tests. *P ≤ 0.05, **P ≤ 0.01. (l,m) Representative images of the gross appearance at 18 d.p.i. of spleens from P. yoelii–infected C57BL/6 mice with or without CTLA-4 blockade (l) and H&E-stained or pseudocolored fluorescently labeled sections of those spleens at the indicated time points (m). Markers and their colors are indicated in the figure in m. All microscopy images represent at least five regions observed in nine sections examined from three separate mice from three separate experiments.

Figure 4: Therapeutic blockade of CTLA-4 enhances immunity to malaria.
figure 4

(ac) Percentages of parasitemia at the indicated time points in C57BL/6 (a) and BALB/c (b,c) mice infected with P. yoelii (a,b) or P. berghei (c) with or without CTLA-4 blockade as depicted in Figure 3a. The time of treatment is indicated by the pink label under the x-axis. (d) Parasitemia at the indicated time points in P. yoelii–infected C57BL/6 mice that received serum (100 μl) at 0 d.p.i., obtained from donor mice treated as in a at 56 d.p.i. (e,f) Parasitemia at the indicated time points (e) and survival (f) in mice from a heterologously challenged with P. berghei at 56 d.p.i. All data are representative of one of at least three separate experiments; each experiment began with five mice per group. Data are presented as mean ± s.e.m. Statistical analysis was performed by comparing the corresponding treatment and control groups at the indicated time points with two-tailed Student's t-tests in ae or a chi-squared test in f. *P ≤ 0.05, **P ≤ 0.01. †, death of one mouse in the corresponding group.

CTLA-4 blockade might be a less realistic independent treatment option for malaria in endemic areas because of its requirements for frequent dosages, parenteral administration, and precise timing; its potential for toxicity; and its currently prohibitive costs. Nevertheless, why humans fail to generate potent adaptive immunity to subsequent Plasmodium infections, which may also involve multiple species of the parasite4,38, is a major unresolved issue in malaria pathogenesis. To investigate the role of CTLA-4 in limiting the generation of long-term immunity, we transferred sera obtained at 56 d.p.i. from C57BL/6 mice that cleared P. yoelii infection with or without CTLA-4 blockade into C57BL/6 recipients with fresh (0 d.p.i.) or established (10 d.p.i.) P. yoelii infections. Recipients of sera from anti-CTLA-4-treated mice, which contained elevated amounts of P. yoelii–specific antibodies (Fig. 3k) but no detectable residual anti-CTLA-4 antibodies (Supplementary Fig. 13a), exhibited significantly lower parasitemia than recipients of sera from mice without anti-CTLA-4 treatment (Fig. 4d and Supplementary Fig. 14). To test whether anti-CTLA-4 therapy aided long-term and perhaps species-transcending immunity, we rechallenged the C57BL/6 mice originally infected with P. yoelii and cured with or without CTLA-4 blockade, using the lethal P. berghei ANKA strain at 56 or 100 d.p.i. CTLA-4 blockade during P. yoelii infection resulted in durable, CD4+ T cell–driven immunity against P. berghei ANKA and significantly improved long-term survival of the mice (Fig. 4e,f and Supplementary Fig. 13b–d). Taken together, these findings suggest that CTLA-4 expression by Treg cells is potentially a major mechanism in the limitation of acquired, cross-species immunity to malaria. Direct evidence of a role for Treg cells and CTLA-4 in limiting protection from Plasmodium reinfection in humans can only be obtained through clinical trials and, like the translation of checkpoint blockade from animal models to human cancer immunotherapy39,40, the precise pathway forward for research in malaria treatment must be carefully defined. However, our findings provide important mechanistic insights to consider while evaluating evidence-based interventions to target host immunity for improved control of malaria.

Malaria is a global health threat, with close to 200 million clinical cases and >500,000 deaths reported annually1. Therefore, it is critically important to understand how Plasmodium protozoa circumvent effective immune responses in humans2. Here we build on detailed studies of immune-cell dynamics during the blood stage of malaria in humans and mice to show how Treg cells can act in a discrete temporal window through CTLA-4 to suppress T helper cell and humoral immune responses. Thus, Treg cells may function as an essential component of the immunoregulation observed in blood-stage malaria to inhibit clearance of acute infection and development of long-term sterilizing immunity to future infections.


Malian and US blood donors.

The Ethics Committee of the Faculty of Medicine, Pharmacy and Dentistry at the University of Sciences, Technique, and Technology of Bamako and the National Institute of Allergy and Infectious Disease of the National Institutes of Health (NIAID/NIH) Institutional Review Board approved the study in Mali (protocol no. 11-I-N126). Informed consent was obtained from the parents or guardians of participating children. The field study, described in detail elsewhere2, was conducted in Mali, where malaria transmission is seasonal. Blood samples were obtained from children at their healthy, uninfected baseline before the malaria season, during their first febrile malaria episode of the ensuing malaria season, and 7 d after treatment of this initial episode. Blood samples of healthy US adults were obtained from the NIH blood bank for research use after written informed consent was obtained from all study participants enrolled in a protocol approved by the NIH Institutional Review Board (protocol no. 99-CC-0168).

Mice and pathogens.

Female C57BL/6, BALB/c and Foxp3-DTR (B6.129(Cg)-Foxp3tm3(DTR/GFP)Ayr/J) mice aged 6–8 weeks were obtained from the Jackson Laboratories. Foxp3-GFP reporter mice41 (a gift from S. Perlman, University of Iowa) and Bcl6-RFP reporter mice42 (a gift from S. Crotty, La Jolla Institute for Allergy and Immunology) were crossbred to generate Foxp3-GFP×Bcl6-RFP.B6 mice. All mice were housed at the University of Iowa Animal Facilities at the appropriate biosafety levels and were subjected to studies approved by the University of Iowa Animal Care and Use Committee. Mice were inoculated with 0.8–1.2 × 106P. yoelii 17XNL– or 8 × 105P. berghei ANKA–infected erythrocytes (originally obtained from the Insectary Core Facility at New York University or clone 234 obtained from Imperial College, London, respectively); 1 × 105 colony-forming units (CFU) of actin-assembly-inducing protein (ActA) from L. monocytogenes (strain DP-L1942)43 i.v.; 2 × 104 tissue culture infectious dose (TCID)50 of influenza A virus (PR8) intranasally; or 5 × 106 plaque-forming units (PFU) of vaccinia virus (Western Reserve) epicutaneously on the ear.

Flow cytometry.

In human studies, PBMCs were isolated and stained, and FACS analysis was performed as described previously44. FACS reagents were purchased from BioLegend, BD Biosciences or eBiosciences and included antibodies to human CD3 (catalog no. UCHT1), CD4 (catalog no. SK3), CD45RO (catalog no. UCHL1), CD45RA (catalog no. HI100), CXCR5 (catalog no. MU5UBEE), CXCR3 (catalog no. G025H7), PD-1 (catalog no. EH12.2H7), HLA-DR (catalog no. LN3), CD38 (catalog no. HIT2), Foxp3 (catalog no. 259D), Helios (catalog no. 22F6), CTLA-4 (catalog no. 14D3), and Ki-67 (catalog no. 20Raj1).

In mouse studies, parasitemia frequencies were determined by flow cytometry as described45. To phenotype lymphocytes from spleen, lymph nodes, or blood, single-cell suspensions were stained on the surface with mouse antibodies to CD16/32 (catalog no. 2.4G2), CD4 (catalog no. RM4-5), CD49d (catalog no. R1-2), CD11a (catalog no. M17/4), PD-1 (catalog no. RMP1-30), CXCR5 (catalog no. L138D7), ICOS (catalog no. C98.4A), CD19 (catalog no. 6D5), GL-7 (catalog no. GL7), CD95 (catalog no. 15A7), CD138 (catalog no. 281-2), IgD (catalog no. -26c), B220 (catalog no. RA3-6B2), CD80 (catalog no. 16-10A1), or CD86 (catalog no. GL-1), or intracellularly with antibodies to CTLA-4 (catalog no. U10-4B9), IDO1 (catalog no. 2E2/IDO1), or FOXP3 (catalog no. FJK-16s) obtained from BioLegend, eBioscience, or BD Biosciences. FOXP3 staining kit (eBiosciences) was used for intracellular staining with the manufacturer's instructions. Multicolor flow cytometry was performed on a BD LSRFortessa and results were analyzed with FlowJo software (Tree Star).

Transendocytosis assay.

As a modification of the assay described elsewhere46,47, Foxp3-eGFP+ Treg cells and Foxp3-eGFP T helper cells (as controls) were sorted using FACS from P. yoelii–infected (9 or 12 d.p.i.) Foxp3-eGFP donor mice and plated in a 1:2 ratio with LPS-matured dendritic cells (DCs) for 3 h in the presence of bafilomycin A. CD80/86 loss by DCs or gain by Treg and T helper cells were assessed by surface or intracellular staining, respectively, and flow cytometry.

Therapeutic regimens.

The following are the dosages of various reagents used in mice, along with the appropriate IgG controls or the diluent: (i) anti-CTLA-4 monoclonal antibody (mAb) (catalog no. UC10-4F10-11, BioXcell) at 500 μg per mouse, i.p., at 9, 11, 13, 15 and 17 d.p.i., (ii) anti-IL-10 mAb (catalog no. JES5-2A5, BioXcell) at 100 μg per mouse, i.p., at 9, 11, 13, 15 and 17 d.p.i., (iii) anti-CD25 mAb48 (catalog no. PC61.5; a gift from S. Varga, University of Iowa) at 500 μg per mouse, i.p., at 9, 11, 13, 15 and 17 d.p.i., (iv) IL-2–anti-IL-2 (catalog no. S4B6, mAb, PeproTech/ATCC) or IL-2–anti-IL-2 (catalog no. JES6-1A12, mAb, PeproTech/ATCC) complexes, made as previously described46 at 1.5 μg per mouse, i.p., at 9, 11, 13, 15 and 17 d.p.i., (v) anti-CD40L mAb33 (MR-1; a gift from T. Waldschmidt, University of Iowa) at 1 mg per mouse, i.v., at 9 and 11 d.p.i., (vi) anti-CD4 mAb (GK1.5, BioXcell), i.p., at 400 μg per mouse at 9 and 11 d.p.i., (vii) DT (Sigma-Aldrich) at 1 μg per mouse, i.p., at 9 and 11 d.p.i., (viii) anti-PDL-1 (catalog no. 10F.9G2, BioXcell) and anti-LAG-3 (prepared from hybridoma clone C9B7W (tested negative for mycoplasma), a gift from D.A.A. Vignali) at 100 μg per mouse each, i.p., at 9, 11, 13, 15 and 17 d.p.i. and (ix) chloroquine (CQ) at 10 mg per kg bodyweight, i.p., at 7, 9, 11, 13, 15 and 17 d.p.i.


Spleen or lymph node sections collected from mice were fixed, stained, and imaged using the Zeiss LSM 710 laser scanning confocal microscope as described in detail previously7. Direct fluorochrome-conjugated antibodies to CD4 (catalog no. GK1.5), B220 (catalog no. RA3-6B2), GL-7 (catalog no. GL7), CD138 (catalog no. 281-2), NRP-1 (catalog no. 761705), CTLA-4 (catalog no. U10-4B9) or FOXP3 (catalog no. 150D) from BioLegend or R&D Systems were used to stain the sections. The cryosections were permeabilized with 1% Triton X-100 (Fischer Bioscience) in intracellular staining for CTLA-4 and FOXP3.

For intravital confocal microscopy, Foxp3-GFP × Bcl6-RFP.B6 mice were injected with B220–Alexa Fluor 647 (Biolegend, 50 μg, i.v.) 14 h before imaging. Mice were anesthetized with ketamine and xylazine (87.5 and 12.5 mg per kg body weight, respectively) and placed with an exposed spleen in dorsal recumbency on the microscope base in a continuously heated (37 °C) enclosed chamber (Leica). A custom suction tissue window apparatus (VueBio) was placed on the spleen with 20–25 mm Hg of negative pressure to immobilize the tissue against a fixed coverslip. Images were acquired on a Leica SP8 Microscope (Leica) using a 25×, 0.95 NA water-immersion objective with coverslip correction. High-resolution confocal stacks of 30–54 xy sections sampled with 1-μm z spacing were acquired at an acquisition rate of 40 frames per second to provide image volumes of 170/388 × 170/388 × 30–54 μm3. Sequences of image stacks were transformed into volume-rendered, 4D time-lapse videos with Imaris software (Bitplane).


MSP1-19-specific antibodies in sera were detected as described previously45. Results are presented as the average endpoint titer, with absorbance readings at 450 nm. To quantify CTLA-4 in tissues, homogenates of whole, weighed spleens or sera were tested with the DuoSet mouse CTLA-4 ELISA kit (R&D Systems) according to the manufacturer's protocol. Residual anti-CTLA-4 antibody in mouse serum was detected using anti–hamster IgG (Poly4055, BioLegend) and quantified using anti-CTLA-4 mAb (UC10-4F10-11, BioXcell) as standard.

Statistical analyses.

For data from human subjects and mice, data were compared using paired or unpaired Student's t-tests, chi-squared tests or ANOVA as appropriate. Bonferroni adjustments (t-tests) and Tukey's corrections (ANOVA) were applied to give a more precise confidence interval (of at least 95%) for differences among the groups in single or multiple comparisons, respectively. All analyses were performed in Prism 6.0h (GraphPad Software).

Data availability.

Data are available from the authors on reasonable request. A Life Sciences Reporting Summary is available.

Additional information

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