More than 2 billion people worldwide are infected with helminths. Thus, it is possible for individuals to experience concomitant infection with helminth and intracellular microbes. Although the helminth-induced type 2 response can suppress type 1 proinflammatory responses required for the immunity against intracellular pathogens in the context of a coinfection, conflicting evidence suggest that helminth infection can enhance antimicrobial immunity. Using a coinfection model with the intestinal helminth Heligmosomoides polygyrus followed by infection with Toxoplasma gondii in Mus Musculus, we showed that the complex and dynamic effect of helminth infection is highly suppressive during the innate phase (days 0–3) of T. gondii infection and less stringent during the acute phase (d10). Helminth coinfection had a strong suppressive effect on the neutrophil, monocytic, and early IFN-γ/IL-12 responses. The IFN-γ response was later restored by compensatory production from T cells despite decreased effector differentiation of T. gondii–specific CD8 T cells. In accordance with the attenuated IFN-γ response, parasite loads were elevated during the acute phase (d10) of T. gondii infection but were transiently controlled by the compensatory T cell response. Unexpectedly, 40% of helminth-coinfected mice exhibited a sustained weight loss phenotype during the postacute phase (d14–18) that was not associated with T. gondii outgrowth, indicating that coinfection led to decreased disease tolerance during T. gondii infection. Our work uncovers the dynamic nature of the helminth immunomodulatory effects on concomitant infections or immune responses and unveils a loss of disease tolerance phenotype triggered by coinfection with intestinal helminth.
Helminths are ubiquitous multicellular organisms that infect one third of the world’s population (representing >2 billion people) (1). On helminth infection, the host mounts a type 2 immune response that is required for worm expulsion and effective repair of the cellular damage caused by traversal of the worm through the host’s tissue. Previous studies have shown that helminth coinfection results in decreased T cell immune responses to heterologous Ags, including pathogens (2–5) and autoantigens (6–8), which is deleterious or beneficial depending on the context. Using a mouse coinfection model, our laboratory has shown that the intestinal helminth Heligmosomoides polygyrus blocks the effector differentiation of CD8 T cells induced by a nonreplicating Toxoplasma gondii vaccine (3). Others have also shown that coinfected mice failed to establish antimicrobial immunity in various models of type 1 immunity (2, 4, 5). These effects correlate with depressed innate DC responses (3, 9, 10) and increased regulatory T (Treg) cell responses (11–13). How broad the helminth-suppressive effects are and whether innate and adaptive responses are equally affected remain open questions. Nevertheless, helminth coinfection does not always lead to decreased antimicrobial immunity and sometimes increases it. Evidence exists that helminth infection can also paradoxically enhance the adaptive immunity against antimicrobial pathogens through bystander effects of IL-4 (14, 15). Infection with helminths has also been shown to improve the immune response against respiratory syncytial virus (16), Pseudomonas aeruginosa (17), and T. gondii (18). Furthermore, it has been shown that helminth-triggered type I IFNs are required for optimal priming of IL-4–producing T cells (19). This raises the possibility that the suppressive effects of helminth infection may not be “airtight” or stable and can be overcome by either counterregulatory effects driven by the coinfecting infectious agent or by, unexpectedly, the type I responses triggered during helminth infection. Finally, although decreased T cell effector function may lead to increased microbial pathogen burdens and is often predicted to cause increased disease pathogenesis, this deleterious effect might be negated by the Treg cell and tolerance-promoting effects of helminth infection (20). Whether increased disease tolerance in the face of higher parasite burdens arises after coinfection with helminths has not been addressed before. In this study, we dissect the effects of helminth coinfection on the innate and adaptive immune responses, as well as disease tolerance, using a previously described coinfection model (2) involving live infections with the helminth H. polygyrus and a nonlethal strain of T. gondii that causes chronic infection.
Materials and Methods
Wild-type female C57BL/6 were purchased from The Jackson Laboratory (Bar Harbor, ME). All mice used for experiments were 6–8 wk old. Handling and housing of all mice was under specific pathogen-free conditions at Rutgers University (Newark, NJ; formerly University of Medicine and Dentistry of New Jersey) and according to the Rutgers Institutional Animal Care and Use Committee guidelines.
Parasite infections and antihelminth treatment
Mice were gavaged with 200 L3 larvae of H. polygyrus on day (d) 0, as previously described (7). On d9, helminth-preinfected mice and control uninfected mice were infected i.p. with 10 cysts of the type II avirulent T. gondii strain, ME49. ME49 cysts were obtained from the homogenization of brains from a chronically infected mouse colony (3–4 wk postinfection). When indicated, helminth-preinfected mice and control uninfected mice were infected i.p. with 1 × 106 tachyzoites of the type II T. gondii strain PTG (ATCC). PTG were passed in monolayers of human foreskin fibroblasts and cultured in DMEM supplemented with 1% FBS and 1% penicillin/streptomycin. For zymosan-induced peritonitis, 10 μg of zymosan (Sigma-Aldrich) was injected i.p., and 4 h later peritoneal responses were analyzed. When indicated, the commercially available antihelminth drug, pyrantel pamoate, was administered orally at d14 and d16 after H. polygyrus infection.
Quantification of parasite burden
Brain, liver, and spleen tissue were collected from either infected or uninfected control animals and stored at −80°C. DNA was extracted using DNeasy Blood and Tissue Kit (QIAGEN) following the manufacturer’s instructions. Parasite load in the indicated amount of genomic DNA per experiment was measured using QuantiTect SYBR Green PCR Kit (QIAGEN) targeting Toxoplasma B1 gene. For quantification of cysts in the brain of chronically infected mice, half of the brain was homogenized in 1 ml of 1X PBS, and the number of cysts was counted under light microscope at a 1:20 dilution.
Serum and peritoneal cytokine levels were measured using OptEIA ELISA Kit for IL-12 (p40/70) (Fisher Scientific) and DuoSet mouse GDF-15 kit (R&D Systems). For Mouse Cytokine/Chemokine 31-Plex Discovery Assay (Assay Genie) of the peritoneal lavage, samples were submitted to Eve Technologies (Calgary, AB, Canada). For the XL cytokine array from the serum at d18 after T. gondii infection, the Proteome Profiler Mouse XL Cytokine Array (R&D) was used. For isolation of proteins from the peritoneal cavity, peritoneal lavage was performed with 1 ml of ice-cold PBS, cells were spun, and supernatants were stored at −80°C.
Tissue preparation and flow cytometry
Mice were sacrificed at the specified study times after T. gondii infection. To harvest peritoneal exudate cells (PECs), we performed peritoneal lavage with 5 ml of RPMI 1640 (Life Technologies) supplemented with 1% FBS and 1% penicillin/streptomycin. Single-cell suspension of spleens was prepared by physical disruption and passage through a 70-μm nylon mesh and washed, and erythrocytes were lysed using a hypotonic Tris-buffered NH4Cl solution. Live cell numbers were counted on a hemocytometer using trypan blue exclusion. Single-cell suspension of the brain was prepared by physical disruption of the brain followed by two washes in cold 1X PBS. Leukocytes were isolated using Histopaque (Millipore Sigma) following the manufacturer’s instruction. Single-cell suspensions were incubated for 20 min in LIVE/DEAD Fixable Violet Dead Cell Stain Kit (ThermoFisher) + Fc Block (BD Pharmingen) on ice. Staining for surface markers and tetramer staining (when indicated) were performed simultaneously in FACS buffer (1X PBS, 10% FBS, 1% penicillin/streptomycin) for 1 h on ice in the dark. The cells were then fixed in BD Cytofix for 20 min at room temperature. For ex vivo staining of intracellular cytokines, fixed cells were incubated in 1X BD Perm/Wash buffer at room temperature for 15 min. Abs targeting intracellular Ags were then diluted in 1X BD Perm/Wash buffer, and samples were stained for 30 min on ice. Samples were then washed and resuspended in FACS buffer for analysis. Flow cytometry data were acquired on BD LSRII and analyzed with FlowJo software (Tree Star, Ashland, OR). Mouse-specific Abs were obtained from BD Biosciences: CD62L (MEL-14), CD45.2 (104); eBioscience (San Diego, CA): CD44 (IM7), IFN-γ (XMG1.2), CD3ε (145.2C11), CD8α (53-6.7), TCRβ (H57-597), KLRG1 (2F1), Ly6C (HK1.4); BD Horizon: CD11b (M1/70), F4/80 (T45-2342), NK1.1 (PK136); BD Pharmingen: Ly6G (1A8), IL-12 (p40/p70), CD4 (RM4-5); or Invitrogen: NKp46 (29A1.4). For PE-labeled SVLAFRRL, H-2Kb tetramers were prepared as previously described (21) and obtained from the National Institutes of Health Tetramer Core facility.
Single-cell RNA sequencing
Splenic TCRβ+CD8α+Tetramer+ cells, which recognize the T. gondii–derived peptide Tgd057, were sort-purified and processed using the single-cell RNA sequencing (scRNAseq) 10× Genomics (Pleasanton, CA) technology. Cell number and viability were determined by a propidium iodide–based fluorescence assay using Moxi GO II System, Orflo Prod#MXG102 (ORFLO Technologies, LLC). Droplet-based single-cell partitioning and scRNAseq libraries were generated from 1000 cells using the 10× Chromium Controller and the Chromium Single-Cell 5′ Reagent v2 Kit (10× Genomics) as per the manufacturer’s protocol. In brief, live cells in single-cell suspension with a >90% viability were mixed with RT reagents and loaded onto a Single-Cell 5′ Chip along with gel beads and partitioning oil in the recommended order, and then the chip was processed through 10× Chromium Controller for the generation of gel beads-in-emulsion. Gel beads-in-emulsion generation was followed by 5′ Gene Expression Library prep protocol. Sample quantification and quality control were determined using Qubit Fluorometer (Invitrogen, Life Technologies) and TapeStation (Agilent Technologies, Santa Clara, CA) respectively. cDNA libraries were sequenced on Illumina NextSeq500 sequencer (Illumina, San Diego, CA) with a configuration of 26/8/0/98 cycles (Read1-10XBarcode+UMI/i7index/i5index/Read2-mRNA reads). The Chromium Single Cell Software was used to analyze and visualize single-cell 3′ RNA sequencing data produced by the 10× Chromium Platform. The 10× chromium software package includes Cell Ranger Pipelines and Loupe Cell Browser. Cell Ranger pipelines use raw 10× single-cell sequencing data from an Illumina sequencer and perform demultiplexing, unique barcode processing, and single-cell 3′ gene counting. Loupe Cell Browser uses data output from the Cell Ranger pipelines for fast analysis of significant genes, cell types, and substructure using single-cell data.
At least three mice per group were used for each experiment. All studies were replicated in one or two repeat experiments. Statistical analysis was carried out using Prism (GraphPad Software, San Diego, CA). Statistical significance was analyzed by two-tailed unpaired Student t test or one-way ANOVA.
Helminth-coinfected mice fail to control T. gondii growth in the acute and chronic phases of toxoplasmosis
T. gondii infection can cause severe congenital disease in developing fetuses (22); however, in immunocompetent individuals, infection is controlled by the innate and adaptive immune response. This is the case with infection with the ME49 (avirulent) strain of T. gondii in mice. Our laboratory and others have shown that mice coinfected with H. polygyrus and T. gondii have defective adaptive immune response against T. gondii (2, 3). To address whether the impaired anti–T. gondii response could have a negative effect on mouse survival, we monitored the viability of coinfected mice after T. gondii infection. Survival of coinfected mice was decreased compared with control ME49-infected mice during the postacute phase (after d12) of toxoplasmosis (Fig. 1A). Parasite load quantification by qPCR in the spleen during peak T. gondii infection (d10) showed that parasite burden was significantly elevated in coinfected mice (Fig. 1B). Coinfected mice also had elevated cyst counts in the brain during chronic toxoplasmosis (d30) (Fig. 1C). Despite the elevated T. gondii loads during the acute and chronic phases of toxoplasmosis, mortality never exceeded 50%, and mice surviving beyond the 30-d time point did not show overt signs of toxoplasmic encephalitis. The decreased survival of coinfected mice during the postacute phase of T. gondii infection and the increased parasite burdens in the spleen and the brain of coinfected mice at d10 and d30, respectively, indicate a partial deficiency in controlling parasite growth.
The helminth-induced functional impairment of the antitoxoplasma response is more evident during the early acute phase of toxoplasmosis
Lack of IFN-γ causes acute susceptibility to T. gondii because of the host’s inability to control replication of the parasite (23). Given that coinfected mice had elevated parasite burdens on d10 after T. gondii infection, we hypothesized that deficiency in IFN-γ production could be the basis for impaired containment of T. gondii growth and/or spread. To test this hypothesis, we studied the kinetics of systemic IFN-γ levels from d3 to d8 after T. gondii infection. The levels of IFN-γ were significantly decreased at d3 in coinfected mice; however, that difference vanished as the levels of IFN-γ escalated on these mice on d5 and d8 (Fig. 2A). By d8 after T. gondii infection, a trend toward higher levels of IFN-γ was evident in coinfected animals. As the levels of IFN-γ decayed in the serum of T. gondii–infected mice at d10 after T. gondii infection, coinfected mice sustained significantly higher levels of the protein. An equivalent delayed response in systemic levels of IL-12 was observed in the coinfected mice (Fig. 2B). At d5 after T. gondii infection, peak IL-12 response was evident in T. gondii–infected mice; however, this response was significantly blunted in coinfected mice. By d8, the IL-12 response in coinfected mice was normalized, mirroring the dynamics in IFN-γ response.
We next quantified the levels of IFN-γ at d3 in the peritoneal cavity, the initial site of T. gondii infection in our model. The levels of IFN-γ in the peritoneal fluid of coinfected mice were decreased 3-fold compared with T. gondii–infected control (Fig. 2C). The numbers of IFN-γ–producing cells in the PECs of coinfected mice were also significantly reduced compared with T. gondii alone–infected mice (Fig. 2D). Neutrophils and NK cells have been shown to be an early source of IFN-γ during T. gondii infection (24, 25). For this reason, we determined the cell-type-specific contribution of IFN-γ in the peritoneal cavity at d3 after T. gondii infection. Neutrophils represented 76% of the IFN-γ–producing cells, followed by T cells (14%) and NK cells (10%) (data not shown). We next assessed the recruitment of neutrophils in the peritoneal cavity at d3 after T. gondii infection and observed that neutrophil numbers were not elevated in coinfected mice, unlike in T. gondii single-infected mice (Fig. 3A). To rule out a helminth-driven disruption of neutrophil formation in the bone marrow, we compared the number of neutrophils in the bone marrow of coinfected, T. gondii single-infected, or H. polygyrus single-infected mice. The number of neutrophils in the bone marrow of coinfected and H. polygyrus–infected mice was elevated compared with T. gondii–infected mice, suggesting that impaired generation of neutrophils in the bone marrow is not the cause for their diminished number in the peritoneum (Fig. 3B). Despite the paucity of neutrophils in the peritoneal cavity of coinfected mice, the number of neutrophils in the blood was significantly increased when helminth-preinfected mice were subsequently infected with T. gondii (Fig. 3C). Peritoneal recruitment of inflammatory monocytes (CD11b+Ly6Chi), but not NK cells, was also impaired in coinfected mice (Fig. 3D and Supplemental Fig. 1A, respectively). We also compared the number of CD8 and CD4 T cells (4 d after T. gondii infection); however, we did not observe a statistically significant reduction in these immune cells (Supplemental Fig. 1B). Altogether, our data suggest that preinfection with an intestinal helminth impairs the innate responses against T. gondii by blocking the recruitment of neutrophils and inflammatory monocytes into the site of infection.
As early as 4 h upon peritoneal injection with T. gondii, neutrophils (26, 27) and, later on (d2–3), inflammatory monocytes (28) are recruited. Gradients of inflammatory chemoattractants are involved in this recruitment (29). Given the observed reduction in the number of infiltrating neutrophils and inflammatory monocytes into the peritoneal cavity of coinfected mice, we tested whether chemoattractant deficiencies could explain the impaired recruitment. Thus, the levels of various chemoattractants were quantified from the peritoneal fluid at 3 d after T. gondii infection. On T. gondii infection, control mice had increased levels of KC, MCP-1, Eotaxin, and MIP-1β; however, this induction was absent in coinfected mice (Fig. 4A–E). Taken together, these results suggest that coinfection with H. polygyrus impairs the T. gondii–induced innate chemokine and cellular responses, resulting in a blunted production of early IFN-γ.
Impaired peritoneal recruitment of neutrophils and monocytes in helminth-coinfected mice is not T. gondii specific
Next, we aimed to determine whether the blunted innate response in helminth-coinfected mice was T. gondii specific. Thus, we first studied the early dynamics of innate cell infiltration into the peritoneum at 4 h after T. gondii (PTG strain) or zymosan challenge. Naive mice challenged with PTG or zymosan exhibited an increase in neutrophils (Fig. 5A, 5C) and inflammatory monocytes (Fig. 5B, 5D) in the peritoneal cavity. In contrast, H. polygyrus–preinfected mice challenged with PTG or zymosan exhibited a decreased infiltration of neutrophils (Fig. 5A, 5C, respectively) and inflammatory monocytes (Fig. 5B, 5D, respectively). These results indicate that the suppressive effect over neutrophil and inflammatory monocyte recruitment is evident as early as 4 h postperitoneal challenge. Given that T. gondii is recognized by TLR2 (30), TLR11 (31), and TLR12 (32), whereas zymosan is recognized by TLR2 (33), TLR6 (34) and Dectin-1, our results suggest that the helminth impairment of peritoneal responses is not T. gondii–specific and is more global in nature. To determine whether helminth blockade of innate responses occurs at the level of innate recognition or more downstream of responses of neutrophils and other innate cells, we used recombinant TNF-α to induce peritonitis. TNF-α treatment in naive mice induced neutrophil recruitment into the peritoneal cavity (Fig. 5E). Interestingly, TNF-α treatment in H. polygyrus–preinfected mice resulted in even a greater increase in neutrophils, approaching levels seen in zymosan-treated naive mice (Fig. 5C). Our results suggest that helminth impairment of innate responses occurs at the level of innate sensing (TLR and Dectin signaling) rather than blocking the cytokine and chemokine responsiveness of effector cells.
Helminth coinfection results in defective generation of toxoplasma-specific effector CD8 T cells
The delayed response of the IFN-γ/IL-12 axis in coinfected mice supported previously published data from our laboratory, which showed that helminth-preinfected mice that were subsequently vaccinated with a nonreplicative T. gondii strain (CPS) had decreased frequency of effector differentiated CD44hiCD62L−KLRG1+ CD8 T cells (3). Thus, we next wanted to investigate the effect of H. polygyrus infection on effector CD8 T cell differentiation in the context of a live T. gondii infection. Helminth-preinfected mice were infected with the T. gondii avirulent strain ME49, and the generation of T. gondii–specific (tetramer+) effector CD8 T cells was measured in the spleen and peritoneal cavity at d10 after T. gondii. We observed decreased numbers of T. gondii–specific effector CD8 T cells in the spleen and peritoneal cavity of helminth-coinfected mice (Fig. 6A). The frequency of IFN-γ–producing T. gondii–specific CD8 T cells was significantly elevated in the spleen and the PECs of coinfected mice (Fig. 6B). This pattern of elevated response was mirrored in the number of IFN-γ–producing T. gondii–specific CD8 T cells only in the spleen and not in the PECs (Fig. 6C). These results agree with the elevated levels of IFN-γ in the serum of coinfected mice at d10 (Fig. 2A). Our results suggest that the helminth-associated impaired effector differentiation of T. gondii–specific CD8 T cells is also evident during live T. gondii infection. Our results also indicate that delayed IL-12 response in coinfected mice could represent a vehicle for the impaired effector differentiation of T. gondii–specific CD8 T cells.
Helminth coinfection does not alter the transcriptional programming of T. gondii–specific CD8 T cells
The decreased representation of T. gondii–specific effector CD8 T cells in helminth-coinfected mice observed in (Fig. 6 prompted us to investigate whether helminth coinfection led to a deviation in the CD8 T cell effector differentiation that is associated with a distinct transcriptional profile. Thus, we sort-purified T. gondii–specific CD8 T cells from the spleen of coinfected or control mice at d10 after T. gondii infection and subjected them to scRNAseq using the 10× genomics. Unbiased clustering of T. gondii–specific CD8 T cells from ME49-infected mice revealed nine distinct clusters that differed in their expression of canonical metabolic, memory, and effector gene markers. The same nine clusters were also present in CD8 T cells isolated from coinfected mice (Fig. 7A). Notably, helminth coinfection did not give rise to a new or unique cluster of T. gondii–specific CD8 T cells.
To functionally define the clusters, we assessed the expression of key transcription factors and T cell effector and adhesion molecule transcripts associated with effector cell and memory precursor cells. Clusters 1 and 2 expressed a low level of the effector-associated transcription factors Tbx21 (35) and Id2 (36) (Fig. 7B) and low expression of Klrg1 and Gzmb (Fig. 7C), but high expression of mitochondrial genes (data not shown), suggesting these cells are metabolically active that have yet to undergo differentiation. Clusters 3 and 4 resembled proliferative cells in the early stages of effector differentiation because these clusters expressed high levels of Mki67 and Birc5 (Supplemental Fig. 2A) and moderate levels of Tbx21 and Id2 (Fig. 7B). Cluster 5 uniquely expressed Tcf7 and the highest levels of Sell (CD62L), Il7r, and Ccr7 (Supplemental Fig. 2B), suggesting these cells are memory precursors. Clusters 6–9 highly expressed the transcription factors Tbx21, Prdm1 (37), and Id2, all of which are essential for differentiation of effector CD8 T cells (Fig. 7B). Consistent with effector differentiation, clusters 6–9 also expressed the canonical effector markers Klrg1, Ifng, and Gzmb (Fig. 7C), and adhesion and chemokine receptors required T cell migration of Itga4, Selplg, and Cxcr3 (Fig. 7D). Restricted expression of transcripts for the fractalkine receptor Cx3cr1 in clusters 7–9 suggested that these clusters represented a more terminally differentiated cell.
Unexpectedly, the variation in the expression patterns of both memory and effector markers across clusters was not statistically different between CD8 T cells derived from ME49 and coinfected mice. Although the number of clusters identified between control and coinfected mice was remarkably similar, the frequency distribution of the CD8 T cells in coinfected mice showed a diminishment for clusters 1 and 4 and enrichment of clusters 6 and 8 relative to ME49-infected mice (Fig. 7E). Thus, our results suggest that rather than causing a deviation of effector differentiation, coinfection with H. polygyrus led to only an attenuation of effector cell differentiation.
Mortality in helminth-coinfected mice is associated with sustained weight-loss phenotype during the postacute phase (d13–18) of toxoplasmosis
The decreased survival in coinfected mice at d10 after T. gondii infection (Fig. 1A) prompted us to characterize in depth the disease state of these mice during the postacute phase of T. gondii infection. To achieve this, we monitored weight, parasite burden, and systemic levels of proinflammatory cytokines during the postacute phase of toxoplasmosis. Published data (38) and unpublished data from our laboratory indicate that T. gondii infection in rodents triggers an IFN-γ– and GDF-15–driven weight loss that peaks between d10 and d12. This weight-loss phase is followed by a recovery phase (d13–18) in which the mice regain weight. We found that during the recovery phase, 45% of coinfected mice did not recover (unrecovered coinfected) from the toxoplasma-induced weight loss, whereas the other 55% exhibited a weight regain (recovered coinfected) pattern similar to T. gondii alone–infected mice (Fig. 8A). When comparing the percentage of weight lost on d18 after T. gondii infection, the unrecovered coinfected cohort sustained a significantly higher percentage of weight loss compared with the recovered coinfected or ME49-infected mice (Fig. 8B). Given the observed increased parasite burden at d10 (Fig. 1B), we hypothesized that these unrecovered coinfected mice were undergoing a phase of uncontrolled parasite growth that resulted in a disease state. For this reason, parasite burden and systemic levels of the proinflammatory cytokine IFN-γ were assessed at d18 after T. gondii infection. No differences were found in parasite burden on the brain and liver between unrecovered coinfected, recovered coinfected, and T. gondii alone–infected mice (Fig. 8C, 8D). Examination of CD8 and CD4 T cell responses from the spleen and mesenteric lymph node on d18 after T. gondii infection revealed that the immune responses between recovered and unrecovered coinfected mice were indistinguishable (Supplemental Fig. 3A–D), suggesting that the heightened weight loss in the unrecovered coinfected mice was not caused by impaired immune responses. There was also no elevation in serum levels of IFN-γ in unrecovered relative to recovered coinfected or ME49-infected mice (Fig. 8E). Altogether, these data led us to conclude that the sustained weight loss in unrecovered coinfected mice is not caused by sporadic loss of parasite growth control or impaired T cell responses.
The earlier data suggested that weight loss in coinfected mice may represent a loss of disease tolerance, rather than impaired immune resistance. Recently, we reported that CD36-deficient mice have impaired disease tolerance during the postacute phase of infection, with elevated serum levels of tissue stress hormones GDF-15 and fibroblast growth factor 21 (FGF-21), both of which cause weight loss (39). Thus, we used a cytokine array to interrogate what cytokines are selectively elevated in unrecovered coinfected mice. Regardless of their weight-loss pattern, coinfected mice did not exhibit any elevation in systemic serum levels of GDF-15, FGF-21, or leptin (Fig. 8E). Among the >100 serum proteins measured in the cytokine array, only three markers were selectively elevated in the unrecovered coinfected mice relative to recovered coinfected and ME49-infected mice. IGFBP-1, but not any other IGFBP, was selectively increased and may signify the presence of endoplasmic reticulum stress in certain tissues and organs (Fig. 8F). Serpin-E1/PAI-1 and pentraxin 3, both of which are acute-phase proteins, were also elevated in the unrecovered coinfected group (Fig. 8F). Thus, rather than presenting with a systemic inflammatory and stress response, the unrecovered coinfected cohort may be undergoing tissue stress and proinflammatory responses in certain organs.
Interestingly, these data also revealed that despite the elevated T. gondii burden observed during acute toxoplasmosis (Fig. 1B), immune control of parasite spread was transiently achieved in helminth-coinfected mice between d13 and d18 after T. gondii infection. The subsequent elevation of cyst counts in the brain of coinfected mice at d30 (Fig. 1C) suggests that immune control of parasite growth falters despite the apparent control observed during the postacute phase. Indeed, examination of brain immune cells on d30 indicated that coinfected mice have a lower frequency of T. gondii–specific CD8 T cells with compensatory elevated frequency of KLRG1+ of T. gondii–specific and total CD8 T cells (Supplemental Fig. 4A–C).
Helminth infection is often associated with increased disease tolerance. Given that we found no differences in the systemic proinflammatory response that could potentially explain the weight loss observed in 44% of the coinfected mice, we next aimed to determine the role of active H. polygyrus infection in decreasing disease tolerance and exerting a sustained weight loss. For this experiment, helminth-coinfected mice were treated with the antihelminth drug, pyrantel pamoate, at d14 and d18 after H. polygyrus infection (d5 and d9 after T. gondii infection) to promote worm expulsion. Worm clearance was visually confirmed by the absence of granuloma-containing H. polygyrus in the duodenum, the region of the small intestine where H. polygyrus attaches once in the lumen (40, 41). Regardless of antihelminth treatment, ∼40–44% of coinfected mice exhibited the high weight-loss phenotype at d18 after T. gondii infection (Fig. 9A). Cyst counts of the brain confirmed no difference in the T. gondii burden is observed at this time point between coinfected and control mice (Fig. 9B).
On visual inspection of the peritoneal cavity, we observed that within the untreated group, unrecovered coinfected mice had loss of epididymal white adipose tissue (eWAT) compared with ME49-infected mice (Fig. 9C). Interestingly, a loss of eWAT was not observed in antihelminth-treated mice (Fig. 9C). These results indicate that the sustained weight loss in unrecovered coinfected mice cannot be entirely explained by dynamic changes in eWAT, given that the sustained weight loss in the antihelminth-treated coinfected mice occurred in the absence of changes in eWAT. These results also suggest the sustained weight loss in unrecovered coinfected mice is not dependent on an active H. polygyrus infection but may be a sequela of the type 2 immunity elicited during helminth infection. Lastly, given that mice with a single infection of H. polygyrus or T. gondii did not exhibit persistent weight loss (Fig. 9), this indicates that the combined effects of T. gondii infection and a concurrent type 2 immune response may play a role in the weight-loss phenotype.
Concurrent helminth infection has been shown to modify the host’s ability to properly respond to infection with unicellular microbes (i.e., virus, bacteria, and protozoa) (4, 5, 42). Despite the well-recognized immunomodulatory effects of the helminth-elicited type 2 immune response over the host’s immunity, conflicting evidence showing enhancement (14, 15, 17) or inhibitory (2, 3, 5) effects on immune resistance to concomitant infection with unrelated pathogens has been reported. Whether helminth infection boosts or dampens host immunity in the context of coinfection is still unclear. Our results highlight the variable and dynamic nature of the helminth immunomodulatory effects on concomitant infections or immune responses. We show that the strength of helminth suppression varies temporally with dominant suppression of innate immunity and only partially affects adaptive immune responses to T. gondii infection. We also described for the first time, to our knowledge, a weight-loss phenotype induced by helminth coinfection during the postacute phase of toxoplasmosis. This phenomenon was only partially penetrant and occurred despite control of T. gondii infection and lack of a systemic cytokine storm, interestingly suggesting that helminth infection resulted in loss of disease tolerance against T. gondii. Our data overall illustrate the complex and often noncanonical effects of helminth infection.
The ability of helminths to suppress the host immune system and negatively affect the response to concomitant microbial infection has been often credited to their ability to induce Treg cells (43–45) and modulation of APC functions (46–49). Evidence now suggests that helminth infection can also alter the responses and migratory patterns of other innate cells (17, 50, 51). Our results highlight a model of innate suppression induced by helminth infection that places the peritoneal neutrophils and inflammatory monocytes as one of the principal targets negatively affected. We showed that neutrophils were the main source of IFN-γ in the peritoneum during the early acute phase of toxoplasmosis (d3), and that helminth coinfection altered the recruitment of IFN-γ–producing neutrophils toward the peritoneum. This resulted in a transient blunting of IFN-γ and a similar delay observed in IL-12, which coincided with the impaired differentiation of effector CD8 T cells observed at d10 after T. gondii infection. These data are in partial agreement with previously published data from neutrophil-depleted mice, which showed decreased systemic levels of IFN-γ and IL-12, a reduction of splenic T cells, and greater parasite burden during the acute phase of toxoplasmosis (52). Neutrophil responses may be critical given the early antitoxoplasmic response is dependent on the IFN-γ–driven activation of monocyte-derived macrophages, which mediate killing and stasis of T. gondii (53–55). Multiple publications show that neutrophil (26, 52, 56) and monocyte depletion (57, 58) during T. gondii infection resulted in uncontrolled growth of this parasite and decreased survival during the postacute phase. Given that early involvement of neutrophils in the initial response against T. gondii sets the grounds for a proper immunological response, the disrupted neutrophil responses we describe in coinfected mice could explain the dysregulated antitoxoplasma responses we observed during the acute and postacute phases of toxoplasmosis, including the elevated T. gondii parasite burdens in the spleen of helminth-coinfected mice. It should be noted that previous studies have shown differential activation of neutrophils during helminth infection, with increased expression of IL-13 and other type 2 markers (59). Future studies should examine whether this altered activation state may also be compromising the neutrophil’s capacity to be effectively recruited into the site of T. gondii infection. Interestingly, our data suggest that the innate deficiency in helminth-coinfected mice was later overcome by compensatory effector functions of IFN-γ from CD8 and CD4 T cells, which coincided with the normalization of IL-12 cytokine responses. Despite the latter compensatory response, parasite burdens on d10 were elevated, illustrating the key import of the early IFN-γ response. Our previous studies in Tyk2-deficient mice have demonstrated that parasite overgrowth resulting from an early IL-12 signaling defect can be difficult to control even when compensatory IFN-γ responses arise later during acute infection (60, 61).
During the acute phase of toxoplasmosis, we found that despite their ability to produce IFN-γ, helminth-coinfected mice had a reduced number of CD62L−KLRG1+ T. gondii–specific effector CD8 T cells. The scRNAseq analysis showed an unaltered trajectory of T. gondii–specific CD8 T cell effector differentiation and the absence of any new cell subpopulation in helminth-preinfected mice, which suggests that the path toward effector differentiation is not different in coinfected mice. These data are supported by the transcriptional similarities in the respective CD8 T cell subpopulations generated in coinfected and singly infected mice, which indicates that the cues required for CD8 T cell differentiation are present in coinfected mice at the time of T. gondii challenge. These data are in agreement with previously published data that use a coinfection model of H. polygyrus and West Nile virus (WNV). In that study, expression of Tbet in WNV-specific CD8 T cells was not different between WNV and coinfected mice (4). We show that in addition to Tbet, other canonical transcription factors associated with an effector phenotype are unaltered by helminth coinfection. Desai et al. (4) showed the increased frequency of WNV-specific CD8 T cells in coinfected mice undergoing apoptosis. Altogether, these data indicate that the decrease in effector CD8 T cells in coinfected mice is not caused by impaired effector differentiation but rather increased turnover of effector CD8 T cells. Despite the decreased number of effector T. gondii–specific CD8 T cells in helminth-coinfected mice during acute toxoplasmosis, there were instances where we paradoxically observed enhanced IFN-γ responses by CD8 T cells to levels even above the control infected mice. We interpret this result as the inevitable response of the CD8 T cells to the overloaded T. gondii burden as a result of the poor innate response observed in coinfected mice. This enhancement of type 1 immune response has been previously observed in the brain of coinfected mice during the chronic phase of T. gondii infection (18).
An alternative and nonmutually exclusive hypothesis to the enhanced CD8 T cell responses in coinfected mice is the bystander role of IL-4 in mediating expansion of virtual memory CD8 T cells. Using the H. polygyrus infection model, Lin et al. (14) showed that virtual memory cells expanded by helminth infection were sufficient to transfer noncognate protection against bacteria to naive mice.
During the postacute phase of toxoplasmosis (d15–18), we showed that 40% of helminth-coinfected mice exhibited sustained weight loss. We initially hypothesized that this weight loss was being caused by uncontrolled parasite growth and elevated levels of IFN-γ and GDF-15. Interestingly, this was not the case because we observed that T. gondii burden in the liver and cyst counts in the brain were controlled at this time point in coinfected mice. It is currently unclear what mediates the weight-loss phenotype, because when we assayed serum from helminth-coinfected mice at d15–18 after T. gondii, there were no elevations in serum IFN-γ or GDF-15 in unrecovered coinfected mice. The observation that T cell responses are not different between recovered and unrecovered coinfected mice at d18 after T. gondii infection strengthens our claims that the sustained weight loss is not driven by the antitoxoplasma response.
The systemic acute-phase response to tissue injury is characterized by elevation of a large number of proteins in the serum. The elevated levels of endoplasmic reticulum stress and tissue damage-associated proteins IGFBP-1, serpin E1/PAI-1, and pentraxin 3 in unrecovered coinfected mice suggest that a more localized trauma might be occurring. The human hepatoma cell line HepG2 is known to produce the acute-phase response protein, serpin E1/PAI-1, in response to IL-1, IL-6, and dexamethasone (62). Pentraxin 3 is known to regulate innate immunity and tissue remodeling in response to liver damage (63). Shimada and Mitchison (64) have proposed a model of drug-induced weight loss in which tissue injury caused by toxins promotes the elevation of kidney-derived GDF-15 and other secreting factors that results in elevation of IGFBP-1 and subsequent increase in tissue catabolism. Although we observed no systemic elevations in GDF-15 in unrecovered coinfected mice, the increase in the stress-associated proteins IGFBP-1, serpin E1/PAI-1 and pentraxin 3 suggests that localized tissue damage could drive the sustained weight loss through alternative pathways that remain to be defined. A direct impact of helminth infection with the intestinal helminth can provide an alternative explanation for the weight loss in coinfected mice. For example, IL-4 has previously been shown to inhibit adipogenesis and promote lipolysis (65). Infection with H. polygyrus and N. brasiliensis, another intestinal helminth, has also been shown to reduce glucose absorption by the small intestine in a process dependent on STAT6 signaling (66, 67). Thus, increased utilization of the stored fat and biomass may represent a maladaptive host response to the metabolic demands of dual (H. polygyrus and T. gondii) infection. Future studies should carefully investigate the relative contributions of decreased food intake, poor nutrient absorption, or increased energy expenditure to the secondary weight loss in coinfected mice.
This work was supported by the National Institutes of Health, National Institute of Allergy and Infectious Diseases (Grant RO1 AI134040 to G.S.Y. and W.C.G.).
The online version of this article contains supplemental material.
The authors have no financial conflicts of interest.