Long pentraxin 3 (PTX3) regulates IL-17A-mediated immunity to Leishmania major infection in mice

Cutaneous leishmaniasis, caused by protozoan parasites of the Leishmania genus, remains a significant health concern in endemic regions such as the Middle-East, Asia, Latin America, and North Africa. The disease affects millions of people worldwide, with over one million new infections reported annually. Despite its health impact, there is currently no approved vaccine largely due to limited understanding of immunological mechanisms underlying protective immunity and disease pathogenesis. We previously reported that long pentraxin 3 (PTX3), a pattern recognition molecule involved in inflammation, tissue repair, and wound healing, is a negative regulator of immunity in primary Leishmania major infection. Specifically, we showed that PTX3 exacerbates disease by suppressing protective Th17 responses. Here, we extend these findings by showing that PTX3 also influences secondary (memory) immunity to L. major. PTX3-deficient (PTX3^-/-) mice which had resolved a primary infection exhibited enhanced resistance to secondary challenge compared to their wild-type (WT) controls. This enhanced resistance correlated with higher frequencies of effector memory CD4⁺ T cells in the spleens and draining lymph nodes. Upon re-infection, healed PTX3^-/- mice produced significantly more IL-17A, while levels of IFN-γ, TNF-α, and IL-10 were similar. In vivo BrdU incorporation assays further revealed increased proliferation of IL-17⁺ CD4⁺ T cells in PTX3^-/- mice. Importantly, neutralization of IL-17A during secondary challenge abolished the enhanced resistance observed in PTX3^-/- mice, confirming a central role of IL-17 in PTX3-regulated secondary immunity. Collectively, our findings identify PTX3 as a key regulator of secondary immunity in cutaneous leishmaniasis and underscores the importance of IL-17 in this process.

1 Introduction

Cutaneous leishmaniasis (CL), the most common form of leishmaniasis affecting about a million new individuals annually, causes significant morbidity and mortality, leaving patients with skin ulcers, permanent scars, disability, and social stigma (1–3). Humans and mice that recover from Leishmania major infection, one of the species that cause CL, retain a low number of parasites at the primary infection site, promoting durable immunity through IFN-γ-producing CD4⁺ T cells (4). This infection-induced immunity is considered the gold standard for effective vaccines or immunotherapies against CL (5). Therefore, understanding the mechanisms governing its generation, maintenance, and loss is critical for developing successful vaccination strategies.

PTX3 is an acute phase inflammatory protein, comprising of a signal peptide, an N-terminal domain unrelated to any known protein, and a C-terminal pentraxin domain homologous to the short pentraxins, CRP and SAP (6, 7). Expressed by immune and non-immune cells (8), it has a dual role in infection. While protective against bacterial, viral, and fungal pathogens, PTX3 has also been implicated in disease susceptibility (9, 10). Although this suppression hinders primary immunity, its role in regulating secondary immunity remains unclear.

We recently showed that PTX3 is induced during L. major infection and suppresses Th17 responses without affecting Th1 immunity (11). Thus, PTX3-deficient (PTX3^-/-) mice show enhanced resistance to primary L. major infection compared to wild-type controls (11). However, if this enhanced primary immunity does not translate to robust secondary immunity, targeting PTX3 may not be a viable therapeutic approach for individuals in endemic regions who face repeated exposures to Leishmania.

In this study, we investigated the impact of PTX3 on secondary immunity using PTX3^-/- mice. Our findings demonstrate that PTX3 suppresses optimal secondary immunity to L. major re-infection by negatively regulating IL-17A responses, thereby providing new insights into its role in immunity and pathogenesis of cutaneous leishmaniasis.

2 Materials and methods

2.1 Animals

Heterozygous female PTX3^+/- and homozygous male PTX3^-/- (originally on129SvEv/Bl/6 background and backcrossed for over 10 generations unto C57BL/6 background) mice were bred at the University of Manitoba Central Animal Care Services (CACS) breeding facility. Female homozygous PTX3 deficient (PTX3^-/-) and their age-matched female littermates control mice (6–8 weeks old) were used in the studies. All animal studies were approved by the University of Manitoba Animal Use and Care Committee in accordance with the Canadian Council for Animal Care guidelines.

2.2 Parasites

Leishmania major parasites (MHOM/IL/80/Friedlin) were grown in M199 medium (Gibco, catalog #11150059) supplemented with 20% heat inactivated fetal bovine serum (FBS) (Hyclone catalog #SH30071.03IH30-45), 2 mM L glutamine (Gibco catalog #25030081), 100 U/ml penicillin, 100 µg/ml streptomycin (Gibco catalog #15140122), 25 mM HEPES (Gibco catalog #15630080) (complete parasite medium). Seven-day stationary phase promastigotes were used for all infection.

2.3 Primary infection, secondary challenge, and measurement of DTH response

Groups of WT and PTX3^-/- mice (4–6 mice per group) were infected in the footpads with 1×10⁶ parasites suspended in 50 µl of sterile PBS. For secondary challenge, infected mice were challenged with 1×10⁶ parasites in the contralateral footpads between 12–16 weeks following primary infection when lesions were fully resolved. Lesion development and delayed type hypersensitivity response (DTH) were determined by measuring the thickness of infected footpads with Vernier caliper.

2.4 Determination of parasite burden

At 3 weeks after primary or secondary infection, mice were sacrificed and parasite burden in the footpads was determined by limiting dilution as previously described (12). Briefly, the footpads were collected and homogenized in 2 ml complete parasite medium using 15 ml tissue grinders. The suspension was then plated in 96-well plates in triplicates at 10-fold serial dilution, incubated for 7 days at 27°C and assessed for parasite growth under a microscope.

2.5 Preparation of soluble Leishmania antigen

Soluble Leishmania antigen (SLA) used for in vitro restimulation of cell cultures was prepared as previously described (13). Briefly, 10⁹ Leishmania major stationary-phase L. major promastigotes (MHOM/IL/80/Friedlin) were pelleted by centrifugation and resuspended in Tris-EDTA buffer (100 mM Tris, 1 mM EDTA, pH 8) supplemented with protease inhibitors. Parasites were lysed by sonication, and cellular debris was removed by ultracentrifugation at 15,000 rpm for 30 minutes. The resulting supernatant was clarified by ultracentrifugation at 35,000 rpm for 2–4 hours, followed by overnight dialysis against PBS. The dialyzed preparation was centrifuged at 10,000 rpm for 10 minutes and filtered using a 0.22 um syringe filters to ensure sterility. Protein concertation of the final SLA preparation was determined using the Bradford assay.

2.6 In vitro recall response and intracellular cytokine staining

At sacrifice, the draining popliteal lymph nodes and spleens from infected WT and PTX3^-/- mice were harvested and made into single-cell suspensions. Cells were washed, resuspended at 5 x 10⁶/ml in complete medium (DMEM (Gibco catalog #12430112 supplemented with 10% heat-inactivated FBS (Hyclone catalog #SH30071.03IH30-45), 2 mM glutamine (Gibco catalog #25030081), 100 U/ml penicillin, and 100 mg/ml streptomycin (Gibco catalog #15140122), and plated at 1 ml/well in 24-well tissue culture plates. Cells were stimulated with SLA, 50 mg/ml for 72 h, and the supernatant fluids were collected and stored at -20°C until assayed for cytokines by ELISA.

For intracellular cytokine analysis, spleen cells and dLNs were assessed for IL-17A, IFN-γ, TNF-α and IL-10 secretion directly ex vivo by flow cytometry following 4 hr in vitro stimulation with Cell activation cocktail with Brefeldin A (Biolegend catalog #423304). Thereafter, the cells were routinely stained for surface marker expression, permeabilized with 0.1% saponin (Sigma-Aldrich catalog #S7900-25G) in staining buffer and then stained with specific fluorochrome-conjugated mAbs against IL-17A (clone: TC11-18H10.1), IFN-γ (clone: W18272D), TNF-α (clone: MP6-XT22) and IL-10 (clone: JES5-16E3) (BioLegend, catalog #506916, 163504, 506308, 505008). Samples were acquired on a BD FACS Cantor machine and analyzed using FlowJo software (TreeStar Inc, Ashland, OR).

2.7 Quantification of transcript levels by RT-PCR

Total RNA was extracted from murine foot pad using the Trizol (Sigma). mRNA was reverse transcribed and cDNA was amplified by RT-PCR using SYBR Green chemistry as described previously (14). Murine primers (PTX3 and β-actin) and reaction conditions were obtained from the PRIMER BANK website http://pga.mgh.harvard.edu/primerbank (Massachusetts General Hospital Primer Bank). Data were normalized to the housekeeping gene β-actin and presented as fold induction over naïve uninfected foot pad cells using the delta-delta CT method. Mouse Forward PTX3-CCTGCGATCCTGCTTTGTG (Primer Bank ID: 31982085a1), Mouse Reverse PTX3-GGTGGGATGAAGTCCATTGTC Primer Bank ID: 31982085a1, Mouse Forward β-actin-AGGAAGTCCCGAAGCAAGAGA (Primer Bank ID: 33585589a1, Mouse Reverse β-actin-CTCCAGTGATTTCTGTGGCAA Primer Bank ID: 33585589a1).

2.8 Cytokine ELISAs

IL-17A, IFN-γ, TNF-α and IL-10 concentrations in cell culture supernatant fluids were measured by sandwich ELISA using Ab pairs from BD Pharmingen (Mouse IFN-γ ELISA Kit catalog #558258) or Biolegend (ELISA MAX Standard Set mouse IL-17A catalog #432501, ELISA MAX Standard Set mouse IL-17A catalog #431411), according to manufacturer’s suggested protocols. The sensitivities of the ELISAs were15 pg/ml, 7.5 pg/ml, I0 pg/ml, and 7.5 pg/ml for IL-17A, IFN-γ, TNF-α, and IL-10, respectively.

2.9 Ex vivo assessment of memory T cell and Treg subsets

L. major infected WT and PTX3^-/- mice were sacrificed at 12 weeks post-infection and single cell suspensions of the draining lymph nodes and spleens were made. The cells were counted and adjusted to 5 × 10⁶/ml and 100 µl aliquots were stained directly ex vivo with different flourochrome-conjugated mAbs against CD3 (clone: 145-2C11), CD4 (clone:

GK1.5), CD44 (clone: NIM-R8), CD62L (clone: MEL-14), CCR7 (clone: 4B12) and CD127 (clone:SB/199) (Biolegend, catalog #100334, 100422, 156008, 104412, 120115 and 121111) and analyzed by flow cytometry. In some experiments, t-distributed Stochastic Neighbor Embedding (tSNE) analysis was performed using FlowJo v10 (FlowJo LLC, Ashland, OR) to visualize high-dimensional flow cytometry data. After gating on live single cells, CD3⁺CD4⁺ T cells were identified and exported, and 2,000 events per sample were randomly downsampled to standardize cell numbers across groups. Dimensionality reduction incorporated the surface markers CD44, CD62L, CCR7, and CD127. t-SNE was run using default settings with 1,000 iterations and a perplexity of 30, and the resulting two-dimensional embeddings were used to display population distributions. Analyses and overlays were annotated according to sample ID.

For Treg staining, spleens and dLN cells were directly stained ex vivo for surface expression of CD3 (clone: 145-2C11), CD4 (clone: GK1.5) and CD25 (clone:PC61) (Biolegend, catalog #100334, 100406, 102012). Thereafter, they cells were stained for intracellular expression of Foxp3 using Treg staining kit (eBioscience, catalog # 00-5523-00) according to the manufacturer’s suggested protocol.

2.10 Ki67 staining and in vivo BrdU labeling

Naïve and healed WT and PTX3^-/- mice were injected (ip) with 2 mg BrdU/mouse and challenged with 1 × 10⁶ L. major parasites in the contralateral footpad the next day. BrdU injection was repeated on days 4 and 6 post-infection and mice were sacrificed on day 7 post-infection to assess cytokine production (IL-17A, IFN-γ, TNF-α and IL-10) by proliferating (i.e., BrdU⁺ or Ki67⁺) CD4⁺ T cells in the dLNs. BrdU and Ki67 intracellular staining were performed with kits/antibodies from BD Pharmingen (catalog# 559619) and Biolegend (catalog# 652422).

2.11 In vivo neutralization of IL-17A

Naïve and healed mice (WT and PTX3^-/-) were treated with either control immunoglobulin (isotype Ig) or anti-IL-17A, neutralizing monoclonal antibodies (0.5 mg/mouse, i.p, all purchased from BioXcell, anti-IL-17A (clone 17F3) catalog# BE0173 and anti-IgG1 isotype (clone: MOPC-21)-catalog# BE0083) 1 day before challenge infection with L. major. Antibody treatment was continued once weekly at 0.5 mg/mouse for additional 3 weeks. Mice were sacrificed after 3 weeks post infection to determine parasite burden.

2.12 Statistics

Results are presented as means ± standard error (SE). Comparisons between groups were performed using paired Student’s t-test or one-way and two-way analysis of variance (ANOVA) with Tukey post hoc correction for multiple comparisons, as appropriate. A p value ≤0.05 was considered statistically significant.

3 Results

3.1 PTX3^-/- mice display enhanced resistance to secondary L. major infection

Recent studies by our group have shown that PTX3 negatively regulates primary immunity to L. major infection (11). To assess whether PTX3 also modulates secondary (memory) immunity L. major, we first determined changes in the levels of PTX3 gene expression following secondary L. major challenge in previously infected and healed mice. Qualitative RT-PCR analysis showed a significant upregulation of PTX3 mRNA at the site of secondary L. major infection (Figure 1A), suggesting that, as in primary infection, PTX3 may also influence immunity during secondary immunity.

Figure 1

Figure 1. Elevated PTX3 expression at the site of secondary L. major challenge is associated with reduced delayed-type hypersensitivity response and increased parasite burden. Naïve or healed (previously infected with L. major) C57BL/6 mice (n = 5) were injected in the footpads with PBS or infected with 1 x 10⁶ stationary-phase L. major promastigotes (contralateral footpads for healed mice). After 7 days, mice were sacrificed and PTX3 mRNA expression at the infection site was assessed by RT-PCR (A). WT and PTX3^-/- mice were infected with 1×10⁶ stationary-phase L. major promastigotes in the footpads and allowed to heal (> 12 weeks). Healed mice, along with their age-matched naïve controls, were then challenged with L. major (contralateral for healed mice). After 72 hours, footpad swelling (a measure of delayed-type hypersensitivity, DTH response) was measured with digital calipers (B). Three weeks after challenge, parasite burden in the infected footpads was quantified by limiting dilution (C). Data represent means ± standard error (n = 5 mice per group) and are representative of 2 independent experiments with similar results. *p < 0.05; **p < 0.01 ***p < 0.005; ****p < 0.0001; ns Not Significant, ND, No parasites detected.

To determine this, we challenged healed wild-type (WT) and PTX3-deficient (PTX3^-/-) mice with L. major and compared their delayed-type hypersensitivity (DTH) responses which is a surrogate marker of memory CD4⁺ T cell-mediated immunity. At 3 days post-secondary challenge, healed PTX3^-/- mice displayed a significantly stronger DTH response compared to healed WT mice (Figure 1B). This heightened DTH response in healed PTX3^-/- mice was associated with significantly lower parasite burden at this site of infection compared to WT healed mice at 3 weeks post L. major challenge (Figure 1C).

Given that DTH responses are primarily mediated by antigen-experience CD4⁺ T cells, which are critical for infection-induced resistance in cutaneous leishmaniasis (15), we hypothesized that healed PTX3^-/- mice would have higher frequency of different subsets of memory CD4⁺ T cells in their spleens and draining lymph nodes compared to their WT counterpart mice. Flow cytometric analysis (Figure 2; Supplementary Figure S5) showed that healed PTX3^-/- mice had significantly higher frequency of splenic effector memory-like (CD44⁺CD62L^-) CD4⁺ T cells in their spleens compared to their WT controls. In contrast, the percentage of central memory-like (CD44⁺CD62L⁺) CD4⁺ T cells were comparable between the two groups (Figures 2A–C). To further characterize the distribution of memory CD4⁺ T cell populations in the spleen, we employed t-distributed stochastic neighbor embedding (tSNE) analysis, which clearly identified two distinct clusters designated P1and P2. The P1 cluster was more prominent in healed PTX3^-/- mice, whereas the P2 cluster was enriched in healed WT mice (Figure 2D). Phenotypic profiling showed that P1 cells expressed higher levels of CD44 and CD127, markers associated with effector memory and long-term survival. In contrast, CD62L and CCR7 expression levels were comparable in both P1 and P2 clusters (Figure 2E). Collectively, these findings indicate that PTX3 regulates immunity to secondary L. major infection, consistent with its previously described role in regulating immunity during primary infection.

Figure 2

Figure 2. Increased frequency of splenic effector memory-like CD4⁺ T cells in L. major infected and healed PTX3^-/- mice. Wild type (WT) and PTX3^-/- were infected with L. major and allowed to heal (>12 weeks after infection). Healed and aged matched naïve control mice were sacrificed, and the frequencies of CD44⁺CD62L^- (effector memory-like) CD4⁺ T cells were analyzed directly ex vivo by flow cytometry in their spleens (A, B) and draining lymph nodes (dLN, A, C) cells by gating on CD3⁺CD4⁺ cells. A tSNE plot of CD4⁺ CD44⁺ T cell clustering from the spleen of healed and aged matched naïve WT and PTX3^-/- mice is shown in (D) along with the expression profiles of CD44, CD62L, CCR7 and CD127 (E). Data are presented as means ± standard error and are representative of 2 independent experiments (n = 3 mice per group) with similar results. *p < 0.05. ns, Not significant.

3.2 Enhanced secondary immunity in PTX3 deficient mice is not associated with increased IFN-γ production

Secondary immunity to L. major, (also known as infection-induced resistance) is primarily mediated by IFN-γ produced by CD4⁺ T cells (4, 16, 17). Consistent with this, we observed both healed WT and PTX3^-/- mice upon secondary L. major challenge exhibited increased frequencies of IFN-γ⁺CD4⁺ T cells in their spleen and draining lymph nodes (dLNs) and their cultures produced significantly higher levels of this cytokine compared to their primary-infected control mice (Figures 3A–E; Supplementary Figure S6). However, there were no significant differences in the frequency of IFN-γ⁺CD4⁺ T cells and levels of IFN-γ in spleen and dLN culture supernatant fluids of WT and PTX3^-/- mice following secondary L. major challenge (Figures 3A–E; Supplementary Figure S6). These findings suggest that the enhanced resistance observed in healed PTX3^-/- mice following secondary challenge was not due to enhanced Th1 response, implicating alternative immune mechanisms in PTX3-regulated secondary protection.

Figure 3

Figure 3. PTX3 deficiency does not affect IFN-γ response during secondary L. major infection. Healed and age-matched naïve wild-type (WT) and PTX3^-/- mice were infected in the footpad (contralateral for healed mice) with 1x10⁶ stationary-phase L. major promastigotes. Three weeks post-infection, mice were sacrificed and the frequency of IFN-γ-producing CD4⁺ T cells in the spleen (A, B) and dLN (A, C) were analyzed directly ex vivo by flow cytometry by gating on CD3⁺CD4⁺ T cells. Additionally, single cell suspensions from spleen (D) and dLN (E) cells were restimulated in vitro with soluble leishmania antigen (SLA, 50 μg/ml) for 72 hr and IFN-γ levels in the culture supernatant fluids were measured by ELISA. Data are presented as means ± standard error and are representative of 2 independent experiments (n = 5 mice per group) with similar results. *p < 0.05; **p < 0.01 ***p < 0.005; ns, Not significant.

3.3 Enhanced secondary immunity in PTX3 deficient mice is not associated with increased TNF-α and IL-10 production

Both TNF-α (18) and IL-10 (4, 19) have also been implicated in regulating secondary immunity in CL. To assess whether differential production of these cytokines contribute to the enhanced protection observed in healed PTX3^-/- mice, we compared their expression in healed WT and PTX3^-/- mice following secondary L. major challenge. Flow cytometric analysis revealed no significant differences in the frequencies of TNF-α⁺CD4⁺ T cells (Supplementary Figures S1A–C, S6) and IL-10⁺CD4⁺ T cells (Supplementary Figures S2A–C, S6) between the two groups. Similarly, levels of TNF-α and IL-10 in the supernatants of spleen and draining lymph node cell cultures were comparable in WT and PTX3^-/- mice (Supplementary Figures S1D, E, 2D, E, respectively).

Given the established role of IL-10-producing Foxp3⁺CD4⁺ regulatory T cells (Tregs) in suppressing secondary immunity to Leishmania major (19–21), we also assessed Treg frequencies. However, the frequency of Tregs in the spleens (Supplementary Figures S3A, B, S6) and dLN (Supplementary Figures S3A–C, S6) were similar between WT and PTX3^-/- mice following secondary L. major challenge. Taken together, these findings suggest that the enhanced resistance observed in healed PTX3^-/- mice following secondary L. major challenge is unlikely to be due to altered production of TNF-α or IL-10 in healed, or due to differences in the frequency of regulatory T cells. This supports the notion that other immune pathways, such as IL-17A responses, may underlie the improved secondary immunity in the absence of PTX3.

3.4 Enhanced secondary immunity in PTX3 deficient mice is associated with increased IL-17A production

We previously reported that PTX3 impairs optimal primary immunity to L. major by suppressing IL-17A production by CD4⁺ T cells (11). Based on these findings, we hypothesized that the enhanced secondary immunity observed in PTX3^-/- mice would be similarly associated with increased IL-17A production. Consistent with this hypothesis, flow cytometric analysis revealed that healed PTX3^-/- mice had significantly higher frequency of IL-17A⁺ CD4⁺ T cells in both the spleen and dLN (Figures 4A–C; Supplementary Figure S6) compared to secondary challenged WT mice. In addition, culture supernatants from spleen (Figure 4D) and dLN (Figure 4E) cells showed markedly elevated levels of IL-17A in PTX^-/- mice relative to controls WT controls. Interestingly, IL-17A levels were also higher in spleen and dLN cell culture supernatants of secondary-infected healed PTX3^-/- mice compared to primary infected PTX3^-/- mice. However, this difference was not observed in direct ex vivo flow cytometric analysis, likely due to stimulation with soluble leishmania antigen during in vitro cultures which could amplify antigen-specific IL-17A response.

Figure 4

Figure 4. PTX3 deficiency enhances Th17 and IL-17 responses during secondary L. major infection. Healed and age-matched naïve wild-type (WT) and PTX3^-/- mice were infected in the footpad (contralateral for healed mice) with 1x10⁶ stationary-phase L. major promastigotes. Three weeks post-infection mice were sacrificed and the frequency of IL-17⁺ -producing CD4⁺ T cells in the spleen (A, B) and dLN (A, C) were analyzed directly ex vivo by flow cytometry by gating on CD3⁺CD4⁺ T cells. Single cell suspension from spleen (D) and dLN (E) were also restimulated in vitro with SLA (50 μg/ml) for 72 hr and the levels of IL-17 in the culture supernatant fluids were determined by ELISA. Data are presented as means ± standard error and are representative of 2 independent experiments (n = 5 mice per group) with similar results. *p < 0.05; **p < 0.01. ns, Not significant.

3.5 Higher frequency of CD4⁺ T cells in healed PTX3^-/- following secondary L. major challenge

Next, we assessed and compared the in vivo proliferative and cytokine producing capacities of CD4⁺ T cells from healed WT and PTX3^-/- mice following L. major challenge. Analysis of dLN showed that both healed WT and PTX3^-/- mice showed increased but comparable frequency of BrdU⁺CD4⁺ T cells (Figures 5A, B; Supplementary Figure S7A) and IFN-γ⁺BrdU⁺CD4⁺ T cells (Figures 5C, D; Supplementary Figure S7A), relative to their primary infection controls. In contrast and consistent with direct ex vivo and in vitro antigen stimulation, healed PTX3^-/- mice showed significantly higher frequency of IL-17⁺BrdU⁺CD4⁺ T cells in the dLN compared to their WT healed mice (Figures 5E, F; Supplementary Figure S7A). This was further supported by Ki67 staining, which showed an increased frequency of IL-17⁺Ki67⁺CD4⁺ T cells in the dLN of PTX3^-/- mice compared to their WT controls (Supplementary Figure S4, S7B). Together, these findings show that antigen-experienced CD4⁺ T cells in healed PTX3^-/- mice proliferate into IL-17A-producing effector cells upon secondary L. major challenge, further implicating IL-17A as a key mediator of enhanced secondary (memory) immunity in the absence of PTX3.

Figure 5

Figure 5. PTX3 deficiency enhances in vivo recall of antigen experienced IL-17⁺CD4⁺ T cells during secondary L. major infection. WT and PTX3^-/- were infected in the footpad with 1×10⁶ stationary-phase L. major promastigotes and allowed to heal (> 12 weeks). Healed mice and their aged matched naïve control mice were injected intraperitoneally with BrdU (2 mg/100 μl) one day prior to challenge with L. major (contralateral footpad for healed mice). A second BrdU injection was administered on Day 4 post challenge, and mice were sacrificed 3 days later. The frequency of proliferating (BrdU⁺) CD4⁺ cells in the dLN (A, B) was analyzed by ex vivo by flow cytometry. The frequency of proliferating and cytokine producing cells (IFN-γ⁺BrdU⁺ and IL-17A⁺BrdU⁺) in the dLN was also analyzed (C-F). Data are presented as means ± standard error and are representative of 2 independent experiments (n = 3 mice per group) with similar results. ***p < 0.005; ns, Not significant.

3.6 Enhanced resistance to secondary L. major infection in PTX3^-/- mice is due to increased IL-17A response

Given that the enhanced resistance of healed PTX3^-/- mice to secondary L. major infection was not associated with higher IFN-γ and TNF-α responses compared to WT controls, we postulated that the enhanced resistance was instead mediated by increased IL-17A production. To test this, we performed in vivo neutralization of IL-17A in both healed WT and PTX3^-/- mice following secondary L. major challenge and assessed parasite burden (Figure 6). Neutralization of IL-17A in healed PTX3^-/- mice abrogated their enhanced secondary immunity as evidenced by significantly higher parasite burden compared to isotype-treated controls (Figure 6). Collectively, these findings show that the enhanced secondary immunity observed PTX3^-/- mice is dependent, at least in part, on increased IL-17A production that likely act in concert with IFN-γ to promote more effective parasite control (Figure 7).

Figure 6

Figure 6. Neutralization of IL-17A abolished enhanced immunity in healed PTX3^-/- mice during secondary L. major infection. Healed and age-matched naïve WT and PTX3^-/- mice were treated intraperitoneally with either an isotype control Ig or anti-IL-17A neutralizing monoclonal antibody (mAb) one day prior to secondary challenge with 1x10⁶ stationary phase L. major promastigotes. Anti-IL17 mAb treatment was continued once weekly for additional 3 weeks. Mice were sacrificed 4 weeks post-infection, and parasite burden in the infected footpads was quantified. Data are presented as means ± standard error and are representative of 2 independent experiments (n = 5 mice per group) with similar results. ****p < 0.0001; ns, Not significant.

Figure 7

Figure 7. Proposed mechanistic model of PTX3 regulation of secondary immunity to Leishmania major. In a PTX3 sufficient mice, healing from Leishmania major infection results in relatively low frequency of effector memory CD4⁺ T cells, suggesting that PTX3 may negatively regulate expansion of these subsets of memory cells. Upon secondary L. major challenge, these memory cells predominantly differentiate into IFN-γ producing CD4⁺ T cells. However, the presence of PTX3 limits the development of IL-17A-producing effector CD4⁺ T cells resulting in lower IL-17A production and reduced IL-17A-mediated effector activity. In contrast, PTX3 deficient mice exhibit increased frequencies of effector memory-like CD4⁺ T cells following primary infection. Upon secondary L. major challenge, these cells proliferate robustly and differentiates into higher frequency of IL-17A-producing CD4⁺ T cells. This elevated IL-17A together with IFN-γ synergistically enhances parasite killing leading to increased host resistance. Thus, in the absence of PTX3, the augmented IL-17A response contributes to improved resistance during secondary infection.

4 Discussion

Our study identifies PTX3 as a key negative regulator of secondary immunity to Leishmania major. We demonstrate that PTX3 expression is elevated during secondary Leishmania major infection, and that its absence (by genetic deletion) results in enhanced protective immunity as evidenced by reduced parasite burden, heightened delayed-type hypersensitivity and increased frequencies of effector memory CD4⁺ T cells. Mechanistically, this enhanced protection was associated with increased production of IL-17A, which acts synergistically with IFN-γ to promote more effective parasite control. This is the first study showing the importance of IL-17A in mediating enhanced secondary immunity to Leishmania infection (Figure 7).

It is well established that IFN-γ-producing CD4⁺ T cells are essential for mediating infection-indued immunity in cutaneous leishmaniasis (16). Depletion of CD4⁺ T cells or neutralization of IFN-γ abolish resistance to infection (22, 23). In the current study, both WT and PTX3-deficient mice mounted strong and comparable IFN-γ response in the spleen and dLN following secondary L. major challenge. In addition, there were no significant differences in the frequency of regulatory T cells or IL-10-producing CD4⁺ T cells between healed WT and PTX3^-/- mice. These observations suggest that the enhanced resistance observed in healed PTX3^-/- mice following secondary L. major challenge is not attributable to enhanced production of IFN-γ by CD4⁺ T cells or reduction in the production of deactivating cytokines such as IL-10 or frequency of regulatory T cells, both of which have been shown to modulate protective immune response in cutaneous leishmaniasis (18, 19).

Previously we showed that PTX3 negatively regulates primary immunity to L. major by suppressing Th17 responses (11). Mechanistically, PTX3 suppresses the production of IL-6 and TGF-β by dendritic cells and expression of Th17 transcription factors such as RORγt, STAT3 and AhR in CD4⁺ T cells (11). Thus, in PTX3^-/- mice, the absence of this regulation leads to increased production of IL-6 and TGF-β by infected DCs (11), favoring optimal differentiation of CD4⁺ T cells into Th17 cells leading to the enhanced production of IL-17A. Several cell types, including γδ T lymphocytes and group 3 innate lymphoid cells (ILC3s), are established sources of IL-17A and play critical roles in shaping immune responses during cutaneous leishmaniasis (24, 25). Although the present study did not directly assess the effect of PTX3 deficiency on IL-17A production from these populations, we observed IL-17A expression within the CD3⁺CD4⁻ compartment, most likely corresponding to CD8⁺ T cells. Notably, PTX3⁻/⁻ mice exhibited significantly higher IL-17A levels in this subset (data not shown), suggesting that PTX3 may exert broader regulatory control over IL-17A responses beyond CD4⁺ T cells and influence additional lymphoid lineages.

In healed mice, the development of a delayed-type hypersensitivity (DTH) response and resistance to reinfection depends on the presence of antigen-specific memory CD4⁺ T cells (22, 23), which can rapidly migrate to peripheral tissues at the site of antigen re-exposure (24). DTH is often used as a surrogate marker of vaccine-induced or infection-acquired immunity in cutaneous leishmaniasis (25). In line with this, we observed that healed PTX3^-/- mice exhibited significantly greater DTH response, a higher frequency of splenic effector memory CD4⁺ T cells (CD44⁺CD62L^-), and reduced parasite burdens following secondary L. major challenge.

Development of memory T cells involves initial activation of naive T cells upon antigen encounter, followed by clonal expansion and acquisition of effector phenotype (26). Following antigen clearance and subsequent immune contraction, a small subset of effector T cells transition into memory phenotype and remain in circulation as either central and effector memory cells while some remain in tissues as tissue resident memory cells (27–30). Central memory T cells preferentially express CCR7 and CD127 and produce IL-2, whereas effector memory cells are poised for rapid IFN-γ production upon antigen restimulation (31–34) Since the quality and magnitude of memory T cell response is directly proportional to the magnitude of primary response, a primary infection must generate sufficient clonal T cell expansion to ensure the generation of optimal memory pool that mediates effective memory T cell response (26). We do not have any evidence suggesting that PTX3 influences initial CD4⁺ T cell clonal expansion following L. major infection, although we did find that healed PTX3 deficient mice contain significantly higher frequency of effector memory-like T cells in their spleens.

In our study, healed PTX3 deficient mice exhibited higher frequencies of effector memory CD4⁺ T cells in the spleen than their WT controls. This likely contributes to their enhanced recall responses manifested as increased DTH, proliferation and cytokine production upon secondary challenge. Flow cytometric analysis further supported this, showing that the P1 cluster (enriched for CD44⁺CD127⁺ cells) was expanded in PTX3 deficient mice. While the exact mechanism by which PTX3 modulates the memory CD4⁺ T cell pool remains to be defined, it is plausible that PTX3 inhibits key aspects of antigen presentation, such as costimulatory molecule expression or cytokine production, that are critical for memory T cell differentiation (28). Although the role of IL-17 in primary resistance to leishmaniasis is equivocal and may be dependent on parasite strain and species, numerous studies support a protective role for IL-17A in L. infantum, L. major and human post kala-azar dermal leishmaniasis (35–37). We previously showed that in PTX3 deficient animals, enhanced IL-17 response synergizes with IFN-γ to promote more efficient parasite clearance in macrophages during primary L. major infection (11). However, its role in secondary immunity has not been elucidated. Our current in vivo IL-17 neutralization studies demonstrated that akin to primary infection, PTX3 deficient mice lose their enhanced resistance to L. major challenge following IL-17A blockade, thereby confirming a critical role for IL-17A in mediating effective secondary immunity in cutaneous leishmaniasis.

In the current study, we observed that healed PTX3⁻/⁻ mice produced significantly higher levels of IL-10 than wild-type controls following secondary L. major challenge. This finding differs from our previous observations during primary infection, in which PTX3 deficiency was associated with markedly reduced IL-10 production (11). We propose that early attenuation of IL-10 responses in PTX3⁻/⁻ mice during primary infection may promote the development of a stronger effector T-cell pool and support more efficient parasite clearance (38, 39). Conversely, the elevated IL-10 detected in healed PTX3⁻/⁻ mice during secondary challenge may act as a counter-regulatory mechanism that tempers the accelerated Th1-driven recall response, thereby reducing the risk of immune-mediated tissue damage and limiting potential immunopathology (37).

The contribution of PTX3 to the pathogenesis of infectious diseases is context dependent and may vary with the nature of the pathogen and the affected tissue. PTX3 confers protection against several bacterial (40–43) and fungal (44) infections through modulation of host inflammatory and innate immune pathways. In contrast, our studies demonstrate that PTX3 promotes susceptibility to cutaneous leishmaniasis caused by L. major, largely by downregulating IL-17A production, which we found to enhance the development of a more effective CD4⁺ Th1 response (11). The role of PTX3 in visceral leishmaniasis caused by L. donovani or L. infantum has not yet been evaluated in our models. Because the immunopathological mechanisms underlying cutaneous and visceral disease are distinct, it is conceivable that PTX3 may play divergent, and potentially protective, functions in VL. Dedicated studies in experimental visceral leishmaniasis are therefore required to define how PTX3 regulates immunity across the different clinical forms of leishmaniasis.

We acknowledge several limitations of this work. First, the study was performed exclusively in a murine model of cutaneous leishmaniasis, and it remains uncertain whether these findings can be fully recapitulated in humans. Second, only female mice were used, which may introduce sex-related bias because sex hormones are known to influence the outcome of CL (45) and also regulate PTX3 expression, including modulation by estrogen and progesterone (46). Consequently, the observed immunological phenotypes may be specific to a particular biological sex and hormonal milieu. Finally, the present study did not define the mechanisms by which PTX3 shapes the function of antigen-specific memory effector CD4⁺ T cells. We did not employ T-cell receptor transgenic models in which CD4⁺ lymphocytes exclusively recognize a defined Leishmania antigen and therefore could not directly assess parasite-specific recall responses at the clonal level. Further studies using such reductionist systems, together with human validation, are required to delineate how PTX3 regulates protective memory immunity.

In summary, our findings clearly highlight the importance of PTX3 and IL-17A in regulating secondary immunity to cutaneous leishmaniasis. We identify PTX3 as a negative regulator of IL-17A-mediated secondary immunity to L. major. In its absence, CD4⁺ T cells expand into effector memory pool capable of mounting robust IL-17A responses leading to improved protection upon reinfection. These findings have important implications for vaccine development and suggest that targeting PTX3 or enhancing IL-17A responses could serve as a strategy to boost long-term protective immunity against cutaneous leishmaniasis.

Data availability statement

The original contributions presented in the study are included in the article/Supplementary Material. Further inquiries can be directed to the corresponding author.

Ethics statement

The animal study was approved by University of Manitoba Animal Care Committee. The study was conducted in accordance with the local legislation and institutional requirements.

Author contributions

GG: Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Project administration, Visualization, Writing – original draft, Writing – review & editing. ZM: Data curation, Formal analysis, Methodology, Investigation, Writing – review & editing. PJ: Methodology, Investigation, Writing – review & editing. CO: Methodology, Investigation, Writing – review & editing. LS: Resources, Writing – review & editing. AS-G: Resources, Writing – review & editing. JU: Conceptualization, Funding acquisition, Project administration, Supervision, Writing – review & editing, Resources.

Funding

The author(s) declared that financial support was received for this work and/or its publication. Our research is supported by financial grants from the Canadian Institutes of Health Research (CIHR) and Research Manitoba.

Acknowledgments

We thank members of the Uzonna lab for their insightful comments.

Conflict of interest

The author(s) declared that this work was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

The author GG declared that they were an editorial board member of Frontiers, at the time of submission. This had no impact on the peer review process and the final decision.

Generative AI statement

The author(s) declared that generative AI was not used in the creation of this manuscript.

Any alternative text (alt text) provided alongside figures in this article has been generated by Frontiers with the support of artificial intelligence and reasonable efforts have been made to ensure accuracy, including review by the authors wherever possible. If you identify any issues, please contact us.

Publisher’s note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

Supplementary material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fimmu.2026.1740323/full#supplementary-material

Supplementary Figure 1 | PTX3 deficiency does not affect TNF-α production during secondary L. major infection. Healed and age-matched naïve wild-type (WT) and PTX3⁻/⁻ mice were infected in the footpad (contralateral for healed mice) with 1×10⁶ stationary-phase L. major promastigotes Three weeks post-infection, mice were sacrificed, and the frequency of TNF-α⁺ -producing CD4⁺ T cells in the spleen (A, B) and dLNs (A, C) was assessed ex vivo by flow cytometry, gating on CD3⁺CD4⁺ cells. Additionally, spleen (D) and dLN (E) cells were restimulated in vitro with soluble Leishmania antigen (SLA; 50 μg/mL) for 72 hours, and TNF-α levels in the culture supernatants were measured by ELISA. Data are presented as means ± standard error and are representative of 2 independent experiments (n = 5 mice per group) with similar results. ns, Not significant.

Supplementary Figure 2 | PTX3 deficiency does not affect IL-10 production during secondary L. major infection. Healed and age-matched naïve wild-type (WT) and PTX3⁻/⁻ mice were infected in the footpad (contralateral for healed mice) with 1×10⁶ stationary-phase L. major promastigotes Three weeks post-infection, mice were sacrificed and the frequency of IL-10⁺ -producing CD4⁺ T cells in the spleen (A, B) and dLNs (A, C) was analyzed directly ex vivo by flow cytometry by gating on CD3⁺CD4⁺ T cells. Additionally, spleen (D) and dLN (E) cells were restimulated in vitro with SLA (50 μg/mL) for 72 hours, and IL-10 levels in the culture supernatants were measured by ELISA. Data are presented as means ± standard error and are representative of 2 independent experiments (n = 5 mice per group) with similar results. *p < 0.05; ****p < 0.0001; ns, Not significant.

Supplementary Figure 3 | PTX3 deficiency does not affect the frequency of CD4⁺CD25⁺FoxP3⁺ (regulatory T cells during secondary L. major infection. Healed and age-matched naïve wild-type (WT) and PTX3⁻/⁻ mice were infected in the footpad (contralateral for healed mice) with 1×10⁶ stationary-phase L. major promastigotes. Three weeks postinfection, mice were sacrificed and single cell suspensions were prepared from spleen (A, B) and dLN (A, C). The frequency of FoxP3⁺CD25⁺ cells (regulatory T cells) was analyzed directly ex vivo by flow cytometry, gating on CD3⁺CD4⁺ T cells. Data are presented as means ± standard error and are representative of 2 independent experiments (n = 5 mice per group) with similar results. ns, Not significant.

Supplementary Figure 4 | PTX3 deficiency enhances in vivo recall of antigen experienced Ki67⁺ IL-17⁺ CD4⁺ T cells during secondary L. major infection. WT and PTX3^-/- were infected in the footpad with 1x10⁶ stationary-phase L. major promastigotes and allowed to heal (> 12 weeks). Healed mice and age-matched naïve controls were challenged with L. major (contralateral footpad for healed mice), and sacrificed on Day 7 post-challenge. Draining lymph node cells were analyzed directly ex vivo by flow cytometry. Proliferating CD4⁺ T cells (Ki67⁺CD4⁺) were assessed (A, B), and the frequency of proliferating, cytokine-producing CD4⁺ T cells was determined by co-expression of Ki67 and IFN-γ (C, D) or IL-17A (E, F), gating first on CD3⁺CD4⁺ cells followed by analysis of Ki67⁺ subsets. Data are presented as means ± standard error and are representative of 2 independent experiments (n = 3 mice per group) with similar results. *, p < 0.05; ns, Not significant.

Supplementary Figure 5 | Gating strategy for Memory CD4⁺ T cells. Splenic or draining lymph node cells were first gated on FSCA (x-axis) and SSCA (y-axis) to select lymphocytes. These lymphocytes were gated on FSCA (x-axis) and FSCH (y-axis) to select singlets. These singlets were then gated on CD3 (x-axis) and CD4 (y-axis) to select double positive CD3⁺ CD4⁺ T cells. These were then gated on CD62L (x-axis) and CD44 (y-axis) to resolve them into effector memory (CD44⁺CD62L^-) CD4⁺ T cells.

Supplementary Figure 6 | Gating strategy for cytokine producing CD4⁺ T cells and T regulatory cells. Splenic or draining lymph node cells were first gated on FSCA (x-axis) and SSCA (y-axis) to select lymphocytes. These lymphocytes were gated on FSCA (x-axis) and FSCH (y-axis) to select singlets. These singlets were then gated on CD3 (x-axis) and CD4 (y-axis) to select double positive CD3⁺ CD4⁺ T cells. These were then gated on IFN-γ⁺ CD4⁺ T cells, IL-10⁺ CD4⁺ T cells, TNF-α⁺ CD4⁺ T cells, IL-17⁺ CD4⁺ T cells, FOXP3⁺ CD25⁺ CD4 ⁺ T cells.

Supplementary Figure 7 | Gating strategy for assessing ex vivo proliferation and cytokine producing CD4⁺ T cells in BrdU and Ki67 stained cells. Splenic or draining lymph node cells were first gated on FSCA (x-axis) and SSCA (y-axis) to select lymphocytes. These lymphocytes were gated on FSCA (x-axis) and FSCH (y-axis) to select singlets. These singlets were then gated on CD3 (x-axis) and CD4 (y-axis) to select double positive CD3⁺ CD4⁺ T cells. These were then gated on proliferating BrdU⁺ CD4⁺ T cells (A) or Ki67⁺ CD4⁺ T cells (B). These proliferating CD4+ T cells were further gated on IFN-γ secreting cells (IFN-γ⁺ BrdU⁺ or IFN-γ⁺ Ki67⁺) and IL-17A secreting cells (IL-17A⁺ BrdU⁺ or IL-17A⁺ Ki67⁺).

References

1. Bilgic-Temel A, Murrell DF, and Uzun S. Cutaneous leishmaniasis: A neglected disfiguring disease for women. Int J Womens Dermatol. (2019) 5:158–65. doi: 10.1016/j.ijwd.2019.01.002

PubMed Abstract | Crossref Full Text | Google Scholar

2. Bailey F, Mondragon-Shem K, Haines LR, Olabi A, Alorfi A, Ruiz-Postigo JA, et al. Cutaneous leishmaniasis and co-morbid major depressive disorder: A systematic review with burden estimates. PloS Negl Trop Dis. (2019) 13:e0007092. doi: 10.1371/journal.pntd.0007092

PubMed Abstract | Crossref Full Text | Google Scholar

3. Lemieux A, Lagacé F, Billick K, Ndao M, Yansouni CP, Semret M, et al. Cutaneous leishmaniasis in travellers and migrants: a 10-year case series in a Canadian reference centre for tropical diseases. CMAJ Open. (2022) 10:E546–E53. doi: 10.9778/cmajo.20210238

PubMed Abstract | Crossref Full Text | Google Scholar

4. Okwor I, Liu D, Beverley SM, and Uzonna JE. Inoculation of killed Leishmania major into immune mice rapidly disrupts immunity to a secondary challenge via IL-10-mediated process. Proc Natl Acad Sci U S A. (2009) 106:13951–6. doi: 10.1073/pnas.0905184106

PubMed Abstract | Crossref Full Text | Google Scholar

5. Okwor I and Uzonna J. Vaccines and vaccination strategies against human cutaneous leishmaniasis. Hum Vaccin. (2009) 5:291–301. doi: 10.4161/hv.5.5.7607

PubMed Abstract | Crossref Full Text | Google Scholar

6. Inforzato A, Rivieccio V, Morreale AP, Bastone A, Salustri A, Scarchilli L, et al. Structural characterization of PTX3 disulfide bond network and its multimeric status in cumulus matrix organization. J Biol Chem. (2008) 283:10147–61. doi: 10.1074/jbc.M708535200

PubMed Abstract | Crossref Full Text | Google Scholar

7. Massimino AM, Colella FE, Bottazzi B, and Inforzato A. Structural insights into the biological functions of the long pentraxin PTX3. Front Immunol. (2023) 14:1274634. doi: 10.3389/fimmu.2023.1274634

PubMed Abstract | Crossref Full Text | Google Scholar

8. Doni A, Stravalaci M, Inforzato A, Magrini E, Mantovani A, Garlanda C, et al. The long pentraxin PTX3 as a link between innate immunity, tissue remodeling, and cancer. Front Immunol. (2019) 10:712. doi: 10.3389/fimmu.2019.00712

PubMed Abstract | Crossref Full Text | Google Scholar

9. Foo SS, Reading PC, Jaillon S, Mantovani A, and Mahalingam S. Pentraxins and collectins: friend or foe during pathogen invasion? Trends Microbiol. (2015) 23:799–811. doi: 10.1016/j.tim.2015.09.006

PubMed Abstract | Crossref Full Text | Google Scholar

10. Daigo K, Inforzato A, Barajon I, Garlanda C, Bottazzi B, Meri S, et al. Pentraxins in the activation and regulation of innate immunity. Immunol Rev. (2016) 274:202–17. doi: 10.1111/imr.12476

PubMed Abstract | Crossref Full Text | Google Scholar

11. Gupta G, Mou Z, Jia P, Sharma R, Zayats R, Viana SM, et al. The long pentraxin 3 (PTX3) suppresses immunity to cutaneous leishmaniasis by regulating CD4. Cell Rep. (2020) 33:108513. doi: 10.1016/j.celrep.2020.108513

PubMed Abstract | Crossref Full Text | Google Scholar

12. Titus RG, Marchand M, Boon T, and Louis JA. A limiting dilution assay for quantifying Leishmania major in tissues of infected mice. Parasite Immunol. (1985) 7:545–55. doi: 10.1111/j.1365-3024.1985.tb00098.x

PubMed Abstract | Crossref Full Text | Google Scholar

13. Scott P, Pearce E, Natovitz P, and Sher A. Vaccination against cutaneous leishmaniasis in a murine model. I. Induction of protective immunity with a soluble extract of promastigotes. J Immunol. (1987) 139:221–7. doi: 10.4049/jimmunol.139.1.221

PubMed Abstract | Crossref Full Text | Google Scholar

14. Oghumu S, Gupta G, Snider HM, Varikuti S, Terrazas CA, Papenfuss TL, et al. STAT4 is critical for immunity but not for antileishmanial activity of antimonials in experimental visceral leishmaniasis. Eur J Immunol. (2014) 44:450–9. doi: 10.1002/eji.201343477

PubMed Abstract | Crossref Full Text | Google Scholar

15. Scott P and Novais FO. Cutaneous leishmaniasis: immune responses in protection and pathogenesis. Nat Rev Immunol. (2016) 16:581–92. doi: 10.1038/nri.2016.72

PubMed Abstract | Crossref Full Text | Google Scholar

16. Okwor I and Uzonna J. Persistent parasites and immunologic memory in cutaneous leishmaniasis: implications for vaccine designs and vaccination strategies. Immunol Res. (2008) 41:123–36. doi: 10.1007/s12026-008-8016-2

PubMed Abstract | Crossref Full Text | Google Scholar

17. Mou Z, Muleme HM, Liu D, Jia P, Okwor IB, Kuriakose SM, et al. Parasite-derived arginase influences secondary anti-Leishmania immunity by regulating programmed cell death-1-mediated CD4+ T cell exhaustion. J Immunol. (2013) 190:3380–9. doi: 10.4049/jimmunol.1202537

PubMed Abstract | Crossref Full Text | Google Scholar

18. Bosch-Nicolau P, Ubals M, Salvador F, Sánchez-Montalvá A, Aparicio G, Erra A, et al. Leishmaniasis and tumor necrosis factor alpha antagonists in the Mediterranean basin. A switch in clinical expression. PloS Negl Trop Dis. (2019) 13:e0007708. doi: 10.1371/journal.pntd.0007708

PubMed Abstract | Crossref Full Text | Google Scholar

19. Zayats R, Mou Z, Yazdanpanah A, Gupta G, Lopez P, Nayar D, et al. Antigen recognition reinforces regulatory T cell mediated Leishmania major persistence. Nat Commun. (2023) 14:8449. doi: 10.1038/s41467-023-44297-6

PubMed Abstract | Crossref Full Text | Google Scholar

20. Belkaid Y, Piccirillo CA, Mendez S, Shevach EM, and Sacks DL. CD4+CD25+ regulatory T cells control Leishmania major persistence and immunity. Nature. (2002) 420:502–7. doi: 10.1038/nature01152

PubMed Abstract | Crossref Full Text | Google Scholar

21. Mendez S, Reckling SK, Piccirillo CA, Sacks D, and Belkaid Y. Role for CD4(+) CD25(+) regulatory T cells in reactivation of persistent leishmaniasis and control of concomitant immunity. J Exp Med. (2004) 200:201–10. doi: 10.1084/jem.20040298

PubMed Abstract | Crossref Full Text | Google Scholar

22. Müller I. Role of T cell subsets during the recall of immunologic memory to Leishmania major. Eur J Immunol. (1992) 22:3063–9. doi: 10.1002/eji.1830221206

PubMed Abstract | Crossref Full Text | Google Scholar

23. Müller I, Kropf P, Etges RJ, and Louis JA. Gamma interferon response in secondary Leishmania major infection: role of CD8+ T cells. Infect Immun. (1993) 61:3730–8. doi: 10.1128/iai.61.9.3730-3738.1993

PubMed Abstract | Crossref Full Text | Google Scholar

24. Dos-Santos JS, Firmino-Cruz L, Oliveira-Maciel D, da Fonseca-Martins AM, Ramos TD, Nunes-Sousa L, et al. IL-17A/IFN-γ producing γδ T cell functional dichotomy impacts cutaneous leishmaniasis in mice. J Leukoc Biol. (2025) 117. doi: 10.1093/jleuko/qiae251

PubMed Abstract | Crossref Full Text | Google Scholar

25. Singh TP, Carvalho AM, Sacramento LA, Grice EA, and Scott P. Microbiota instruct IL-17A-producing innate lymphoid cells to promote skin inflammation in cutaneous leishmaniasis. PloS Pathog. (2021) 17:e1009693. doi: 10.1371/journal.ppat.1009693

PubMed Abstract | Crossref Full Text | Google Scholar

26. Kaech SM and Wherry EJ. Heterogeneity and cell-fate decisions in effector and memory CD8+ T cell differentiation during viral infection. Immunity. (2007) 27:393–405. doi: 10.1016/j.immuni.2007.08.007

PubMed Abstract | Crossref Full Text | Google Scholar

27. Kaech SM, Wherry EJ, and Ahmed R. Effector and memory T-cell differentiation: implications for vaccine development. Nat Rev Immunol. (2002) 2:251–62. doi: 10.1038/nri778

PubMed Abstract | Crossref Full Text | Google Scholar

28. Opata MM and Stephens R. Early decision: effector and effector memory T cell differentiation in chronic infection. Curr Immunol Rev. (2013) 9:190–206. doi: 10.2174/1573395509666131126231209

PubMed Abstract | Crossref Full Text | Google Scholar

29. Jameson SC and Masopust D. Diversity in T cell memory: an embarrassment of riches. Immunity. (2009) 31:859–71. doi: 10.1016/j.immuni.2009.11.007

PubMed Abstract | Crossref Full Text | Google Scholar

30. Sallusto F, Lenig D, Förster R, Lipp M, and Lanzavecchia A. Two subsets of memory T lymphocytes with distinct homing potentials and effector functions. Nature. (1999) 401:708–12. doi: 10.1038/44385

PubMed Abstract | Crossref Full Text | Google Scholar

31. Masopust D, Vezys V, Marzo AL, and Lefrançois L. Preferential localization of effector memory cells in nonlymphoid tissue. Science. (2001) 291:2413–7. doi: 10.1126/science.1058867

PubMed Abstract | Crossref Full Text | Google Scholar

32. Bjorkdahl O, Barber KA, Brett SJ, Daly MG, Plumpton C, Elshourbagy NA, et al. Characterization of CC-chemokine receptor 7 expression on murine T cells in lymphoid tissues. Immunology. (2003) 110:170–9. doi: 10.1046/j.1365-2567.2003.01727.x

PubMed Abstract | Crossref Full Text | Google Scholar

33. Huster KM, Busch V, Schiemann M, Linkemann K, Kerksiek KM, Wagner H, et al. Selective expression of IL-7 receptor on memory T cells identifies early CD40L-dependent generation of distinct CD8+ memory T cell subsets. Proc Natl Acad Sci U S A. (2004) 101:5610–5. doi: 10.1073/pnas.0308054101

PubMed Abstract | Crossref Full Text | Google Scholar

34. Reinhardt RL, Khoruts A, Merica R, Zell T, and Jenkins MK. Visualizing the generation of memory CD4 T cells in the whole body. Nature. (2001) 410:101–5. doi: 10.1038/35065111

PubMed Abstract | Crossref Full Text | Google Scholar

35. Nascimento MS, Carregaro V, Lima-Júnior DS, Costa DL, Ryffel B, Duthie MS, et al. Interleukin 17A acts synergistically with interferon γ to promote protection against Leishmania infantum infection. J Infect Dis. (2015) 211:1015–26. doi: 10.1093/infdis/jiu531

PubMed Abstract | Crossref Full Text | Google Scholar

36. Pitta MG, Romano A, Cabantous S, Henri S, Hammad A, Kouriba B, et al. IL-17 and IL-22 are associated with protection against human kala azar caused by Leishmania donovani. J Clin Invest. (2009) 119:2379–87. doi: 10.1172/JCI38813

PubMed Abstract | Crossref Full Text | Google Scholar

37. Gonzalez-Lombana C, Gimblet C, Bacellar O, Oliveira WW, Passos S, Carvalho LP, et al. IL-17 mediates immunopathology in the absence of IL-10 following Leishmania major infection. PloS Pathog. (2013) 9:e1003243. doi: 10.1371/journal.ppat.1003243

PubMed Abstract | Crossref Full Text | Google Scholar

38. Belkaid Y, Hoffmann KF, Mendez S, Kamhawi S, Udey MC, Wynn TA, et al. The role of interleukin (IL)-10 in the persistence of Leishmania major in the skin after healing and the therapeutic potential of anti-IL-10 receptor antibody for sterile cure. J Exp Med. (2001) 194:1497–506. doi: 10.1084/jem.194.10.1497

PubMed Abstract | Crossref Full Text | Google Scholar

39. Kane MM and Mosser DM. The role of IL-10 in promoting disease progression in leishmaniasis. J Immunol. (2001) 166:1141–7. doi: 10.4049/jimmunol.166.2.1141

PubMed Abstract | Crossref Full Text | Google Scholar

40. Moalli F, Paroni M, Véliz Rodriguez T, Riva F, Polentarutti N, Bottazzi B, et al. The therapeutic potential of the humoral pattern recognition molecule PTX3 in chronic lung infection caused by Pseudomonas aeruginosa. J Immunol. (2011) 186:5425–34. doi: 10.4049/jimmunol.1002035

PubMed Abstract | Crossref Full Text | Google Scholar

41. Paczosa MK and Mecsas J. Klebsiella pneumoniae: going on the offense with a strong defense. Microbiol Mol Biol Rev. (2016) 80:629–61. doi: 10.1128/MMBR.00078-15

PubMed Abstract | Crossref Full Text | Google Scholar

42. Asgari F, Supino D, Parente R, Polentarutti N, Stravalaci M, Porte R, et al. The long pentraxin PTX3 controls. Front Immunol. (2021) 12:666198. doi: 10.3389/fimmu.2021.666198

PubMed Abstract | Crossref Full Text | Google Scholar

43. Soares AC, Souza DG, Pinho V, Vieira AT, Nicoli JR, Cunha FQ, et al. Dual function of the long pentraxin PTX3 in resistance against pulmonary infection with Klebsiella pneumoniae in transgenic mice. Microbes Infect. (2006) 8:1321–9. doi: 10.1016/j.micinf.2005.12.017

PubMed Abstract | Crossref Full Text | Google Scholar

44. Garlanda C, Hirsch E, Bozza S, Salustri A, De Acetis M, Nota R, et al. Non-redundant role of the long pentraxin PTX3 in anti-fungal innate immune response. Nature. (2002) 420:182–6. doi: 10.1038/nature01195

PubMed Abstract | Crossref Full Text | Google Scholar

45. Snider H, Lezama-Davila C, Alexander J, and Satoskar AR. Sex hormones and modulation of immunity against leishmaniasis. Neuroimmunomodulation. (2009) 16:106–13. doi: 10.1159/000180265

PubMed Abstract | Crossref Full Text | Google Scholar

46. Camaioni A, Klinger FG, Campagnolo L, and Salustri A. The influence of pentraxin 3 on the ovarian function and its impact on fertility. Front Immunol. (2018) 9:2808. doi: 10.3389/fimmu.2018.02808

PubMed Abstract | Crossref Full Text | Google Scholar