All individuals were examined as part of a natural history study protocol approved by Institutional Review Boards of the National Institute of Allergy and Infectious Diseases, USA and the National Institute for Research in Tuberculosis, India (ClinicalTrials.gov identifiers NCT00375583, and NCT00001230), and informed written consent was obtained from all participants.

We studied a total of 74 individuals comprising 34 clinically asymptomatic, S. stercoralis-infected individuals (Inf) and 40 uninfected, healthy individuals with endemic normal status (EN) in Tamil Nadu, South India (Tables 1, 2). All individuals were recruited from a rural population by screening of individuals for helminth infection by stool microscopy and serology. Inclusion criteria were age of 18 to 65 years and willingness to give blood and stool samples for examination; exclusion criteria were past anthelmintic treatment, other helminth infections, or HIV infection. Strongyloides infection was diagnosed by the presence of IgG antibodies to the recombinant Strongyloides antigen, NIE, as described previously (Bisoffi et al., 2014; Buonfrate et al., 2015). None of the individuals had lymphatic filariasis verified with TropBio Og4C3 enzyme-linked immunosorbent assay (ELISA) (Trop Bio Pty. Ltd., Townsville, Queensland, Australia) or other intestinal helminths (based on stool microscopy). None of the tested individuals suffered from acute tuberculosis, analyzed via QuantiFERON TB Gold-in-Tube enzyme-linked immunosorbent assay (ELISA) (Cellestis).

All infected individuals were treated with single doses of ivermectin and albendazole. All uninfected individuals were anti-Strongyloides NIE negative and negative for filarial and other intestinal helminths as well as acute tuberculosis.

Hemograms were performed on all individuals using the Act-5 Diff hematology analyzer (Beckman Coulter, Brea, CA, USA).

Heparinized blood was centrifuged at 400 × g for 10 min, plasma separated and stored at 4°C. Samples were refilled with RPMI-1640 and PBMC isolated by Ficoll diatrizoate gradient centrifugation (LSM; ICN Biomedicals). Cells were then washed, resuspended in freezing medium (RPMI 1640, 10% DMSO, 45% heat inactivated FCS, Harlan Bioproducts) and stored in liquid nitrogen until usage. For in vitro culture the cryopreserved cells were thawed gently, washed twice and cultured with RPMI 1640 medium supplemented with penicillin-streptomycin (100 U and 100 μg/ml, respectively), L-glutamine (2 mM), 1% NEAA MEM (all from PAN–Biotech, Aidenbach, Germany) and 5% heat inactivated AB human serum (Biochrom, Berlin, Germany). For stimulations and intracellular staining cells were counted using trypan blue and adjusted to 1 × 10⁷ cells/ml. 2 × 10⁶ cells/well were placed on round-bottom 96 well tissue culture plates, stimulated with phorbol 12-myristate 13-acetate (PMA) and ionomycin (both Sigma-Aldrich, MO, USA) at concentrations of 25 ng/ml and 0.5 μg/ml or media alone and incubated for 4 h at 37°C. Brefeldin A solution (10 μg/ml, eBioscience, CA, USA) was added after 30 min. After 4 h cells were washed, stained and analyzed as given below.

Plasma levels of total IgE, IgG1, IgG3, and IgG4 were evaluated using Ready-Set-Go! ELISA kits (eBioscience) according to the manufacturer's instructions. All samples were run in duplicates.

Animal experiments were performed in accordance with the National Animal Protection Guidelines and approved by Federal Health Authorities of the States of Hamburg (permission number 55/13). The S. ratti life cycle was maintained in Wistar rats purchased from Charles River (Sulzfeld, Germany) as described earlier (Eschbach et al., 2010). Female C57BL/6 mice were bred in house at the Bernhard Nocht Institute Hamburg, Germany. Mice were kept in individually ventilated cages under specific pathogen-free (SPF) condition, infected by 200 or 2,000 S. ratti iL3 in the hind footpad and sacrificed by isoflurane inhalation followed by cervical dislocation at day 10 or 20 post infection at 8–10 weeks of age. Parasite burdens were examined at day 6 post infection in faces via quantification of S. ratti 28S ribosomal RNA using real-time quantitative PCR as described elsewhere (Nouir et al., 2012).

Spleens and mesenteric lymph nodes (mLN) were isolated and placed in cold RPMI 1640 wash medium containing 1% FCS, 100 U/ml penicillin and 100 μg/ml streptomycin (all from PAN – Biotech, Aidenbach, Germany) and then forced through 70 μm cell strainers (BD Bioscience, San Jose, CA) to obtain single cell suspensions. Erythrocytes in spleen cell suspension were lysed using ACK buffer containing 150 mM NH₄Cl, 0.1 mM KHCO₃ and 0.1 mM Na₂EDTA, pH 7.2 for 5 min on ice followed by two washing steps. Small intestinal lamina propria (siLP) cells were isolated as described earlier (Strandmark et al., 2016).

Lungs were perfused and rinsed with 0.9% NaCl until the tissue turned white before isolation of single cells. The lung was rinsed in ice-cold RPMI 1640 containing 10% FCS. Tissue was then transferred onto petri dishes, cut in small pieces (~ 2 mm) and subsequently added to 10 ml PBS containing 0.1265 U/ml collagenase D and DNase (0.15 mg/ml, both Sigma-Aldrich, Steinheim, Germany) followed by incubation in a tube shaker water bath (250 rpm, 37°C) for 1 h. Digested lung tissues were forced through 40 μm cell strainers, washed twice and erythrocytes lysed using ACK buffer followed by two washing steps.

For the isolation of PBMC, whole blood was diluted 2:1 with RPMI 1640 containing 10% FCS and layered onto Pancoll (density of 1.077 g/ml, PAN-Biotech, Aidenbach, Germany), centrifuged and washed twice with PBS.

Cells of all tissues were resuspended in RPMI 1640 containing 10% FCS, counted using a CASY automated cell counter (Roche-Innovatis, Reutlingen, Germany) and were adjusted to 1 × 10⁷ cells/ml. Cells were plated on round-bottom 96 well tissue culture plates (Costar, Corning Inc., NY, USA), stimulated with PMA and ionomycin at concentrations of 50 ng/ml and 1 μg/ml and incubated for 30 min at 37°C, followed by addition of Brefeldin A (10 μg/ml) and further stimulation for 2.5 h.

Human cells were stained with fixable viability dyes (eFluor780 or eFluor506 eBioscience), fixed and permeabilized using the eBioscience Foxp3 staining kit and stained intracellularly for the following markers: CD3 (BV510, clone OKT3), CD4 (PerCP, clone SK3), CD45RO (FITC, clone UCHL1), IL-4 (PE, MP4-25D2), IL-5 (PE, clones MP4-25D2, TRFK5), IL-13 (FITC, clone PVM13-1), IFN-γ, (eF450, clone 4S.B3), GATA-3 (eF660, clone TWAJ) and T-bet (PE-Cy7, clone eBio4B10). Murine cells were stained with fixable viability dye (eFluor780, eBioscience) and subsequently fixed and permeabilized using the Foxp3 staining kit and stained intracellularly for the following markers: CD4 (PerCP, clone RM4-5), GATA-3 (eFluor660, clone TWAJ), T-bet (PE-Cy7, clone eBio4B10), IL-4 (PE, clone 11B11), IL-13 (Alexa488, clone eBio13A), IL-5 (PE, clone TRFK5), IFN-γ (eFluor450, clone XMG1.2). All antibodies were from eBioscience, BioLegend and BD Biosciences.

Cells from spleens and peripheral lymph nodes of naïve C57BL/6 mice were stained for CD4 (BrilliantViolet 510, clone RM4-5), CD62-L (APC-eFluor780, clone MEL-14), CD44 (PE, clone IM7) and CD25 (APC, clone 61.5) (all antibodies from eBioscience and BioLegend) and naïve CD4⁺CD25⁻CD62-L^highCD44^neg cells were isolated on a FACS Aria cell sorter (BD Bioscience). Cells were plated on 48 well plates (Costar) coated with anti-CD3/anti-CD28 antibodies (1 μg/ml each, clones 145-2C11 and 37.51, from BD Biosciences). Th1 differentiation was induced by addition of recombinant murine IFN-γ and IL-12 (10 ng/ml each, PeproTech) and 10 μg/ml anti-IL-4 (clone 11B11, BioLegend, San Diego, CA, USA). Th2 development was induced by addition of 30 ng/ml recombinant murine IL-4 (PeproTech) and anti-IFN-γ/anti-IL12/23 p40 antibodies (clones AN18, C17.8, BioLegend, 10 μg/ml each). Th2/1 hybrids were generated by addition of IFN-γ, IL-12 and IL-4 in the concentrations given above. Recombinant human IL-2 (10 ng/ml, PeproTech) was added to all cultures and replaced in fresh medium on day 3. Cells were analyzed for expression of T-bet, GATA-3, IFN-γ IL-4 and IL-13 after 5 to 6 days by intracellular antibody staining.

Cells isolated from mice infected with S. ratti for 10 days were stimulated with the following recombinant murine cytokines (all from PeproTech, Hamburg) or monoclonal antibodies (from BD Biosciences) for 6 h: IL-33 (100 ng/ml) and IL-7 (10 ng/ml), IL-18 and IL-12 (both 10 ng/ml), anti-CD3 (clone 145-2C11) and anti CD28 antibodies (clone 37.51, both 2 μg/ml). Controls were left untreated. Brefeldin A (10 μg/ml) was added after 30 min. Cells were then stained for CD4, GATA-3, T-bet, IL-13 and IFN-γ.

Peritoneal cells were isolated from naïve C57BL/6 mice by flushing the peritoneal cavity with ice-cold PBS (20 mM EDTA, 0.2% BSA) followed by washing in RPMI. 3 × 10⁵ cells were plated on 96 well round-bottom plates (Costar) in 200 μl in RPMI and incubated at 37°C for 1 h. Non-adhering cells were removed by 3 washing steps with warm RPMI. 1 × 10⁵ in vitro generated Th1, Th2 or Th2/1 cells were added and co-cultured in 200 μl for 24 h. Lipopolysaccheride (LPS, L5529, Sigma) was added for co-stimulation at 0.5 μg/ml to some wells. Control cultures received 1 or 10 ng/ml recombinant murine IFN-γ. Nitrite concentrations in culture media were quantified by the Griess reaction.

Proximal small intestinal tissue samples were fixed in formalin and histochemically stained with hematoxylin and eosin (H&E) to assess histomorphology and with periodic acid Schiff (PAS) for goblet cell quantification.

Enteritis was scored as described earlier (Rausch et al., 2009). Images were acquired using the AxioImager Z1 microscope (Carl Zeiss MicroImaging, Inc., Jena, Germany) at 100× magnification. All evaluations were performed in a blinded manner.

All data were assessed for Gaussian distribution. Statistically significant differences between two groups were analyzed by using Student's t-test (parametric) or Mann-Whitney U-test (non-parametric) using GraphPad Prism 7 software (San Diego, CA). For multiple comparisons, the Kruskal-Wallis test with Dunn's correction was performed. P-values ≤ 0.05 were considered to indicate statistical significance.

The regression analyses were carried out using SAS 9.4. Quantitative variables were tested for normality (PROC UNIVARIATE) and when necessary transformed to logarithmic values. After graphic assessment of the assumption of linear relationships (PROC GPLOT) and assessment of correlations between variables (PROC CORR) associations between variables were investigated by multivariable linear models with the procedure PROC REG. Models with different independent factors were compared by using the adjusted R² and the global F-statistics. The variable “group” as well as the main variable of the respecting analyses were always kept in the model, but other factors were removed in a backward selection according to the p-value and the change of R². Model diagnostics included residues assessment (normality), test for multicollinearity and variance inflation. Observations with great influence (according to leverage hat-values or Cook's D) were excluded, if the model changed significantly after removing the observation. P-values ≤ 0.05 were considered to indicate statistical significance.

In order to assess the immune response to S. ratti, mice were infected with 200 or 2,000 infective stage 3 larvae and dissected at day 10 or 20 post infection (Figure 1A). Worm burdens were assessed at day 6 by quantification of Strongyloides DNA in faces. Expectedly and as shown before, mice infected with 2,000 larvae displayed significantly higher worm burdens than mice inoculated with 200 larvae (Figure 1B; Eschbach et al., 2010). Histological examination of proximal small intestinal tissue samples showed that the infection did not result in immunopathological changes by day 10, while some mice displayed mild signs of cellular infiltration at day 20 post infection (Figure 1C). Numbers of mucus-producing goblet cells were at best mildly and transiently, but not significantly increased by day 10 post infection (Figure 1D).

Expectedly, infected mice displayed significantly increased frequencies of CD4⁺ T cells expressing GATA-3 and IL-4 in spleen, gut-draining mesenteric lymph nodes (mLN) and small intestinal lamina propria (siLP) when compared to uninfected controls (Figures 2A–D). Th2 cells were also elevated in blood and lung tissue of infected mice (SI Figures 1A,B). T-bet⁺ Th1 cells tended to increase in the spleen of infected mice, reaching significance at day 20 post infection, while IFN-γ expression by CD4⁺ T cells from spleens was transiently elevated at day 10 (Figures 2E,F). Frequencies of T-bet expressing cells in mLN, siLP and other organs were similar in all groups (Figures 2G,H and SI Figures 1C,D) while IFN-γ responses in the small intestine were significantly reduced in infected mice (Figure 2H).

Collectively, S. ratti infection led to local and systemic Th2 responses confirming previous studies (Eschbach et al., 2010; Blankenhaus et al., 2011), while transiently elevated IFN-γ production and expansion of T-bet expressing Th1 cells was restricted to the spleen. Pathological changes and Th2-driven increases of goblet cells in the small intestine were only mild, pointing out the relatively asymptomatic nature of S. ratti infection in mice.

Asking whether infections with S. ratti led to the expansion of Th2/1 cells as previously shown for other helminth infections (Peine et al., 2013), we screened the organs of naïve and infected mice for CD4⁺ T cells simultaneously expressing GATA-3 and T-bet. We detected significantly increased frequencies of GATA-3⁺T-bet⁺ Th2/1 cells in all organs of infected mice (Figures 3A–D and SI Figures 2A,B). The proportions of T-bet co-expressing Th2/1 cells within the total GATA-3⁺CD4⁺ population were highest in spleen and siLP of S. ratti infected mice (Figure 3E). T-bet/IFN-γ co-expressing cells were detected in similar proportions within IL-4 and IL-13 producers from spleens of mice infected for 10 days (Figure 3F) and the frequencies of IFN-γ⁺ Th2/1 hybrids in Th2 cytokine producing populations were similar when comparing day 10 and 20 post infection (Figure 3G).

As shown previously (Peine et al., 2013), Th2/1 hybrid cells expressed lower levels of GATA-3 and IL-4 than conventional Th2 cells (Figure 3H) and IFN-γ production was significantly decreased compared to Th1 cells (Figure 3I). Notably, T-bet expression by Th2/1 cells was significantly elevated compared to infection-derived Th1 cells (Figure 3I), which might reflect the acute activation status of the Th2/1 cells.

Thereby, our previous demonstration of the induction of Th2/1 hybrid cells in nematode and Schistosome infections (Peine et al., 2013) can be generalized to mice infected with different doses of threadworms.

To evaluate whether CD4⁺ T cells with combined characteristics of Th2 and Th1 cells also occur in human helminth-infected patients we investigated blood samples of patients infected with S. stercoralis and endemic uninfected control donors. The demographic profile of the cohorts is given in Table 1. No differences in age range, gender and socio-economic status were observed between the two studied groups. All individuals were healthy and free of symptoms. Hematological features of both cohorts are depicted in Table 2. No differences concerning hemoglobin levels, red and white blood cell counts, hematocrit or platelet counts were observed between the groups. The frequencies of lymphocytes and eosinophils were significantly elevated in the infected cohort (SI Figures 3A,D), while neutrophil frequencies were significantly lower in S. stercoralis infected patients (SI Figure 3B). Blood monocyte and basophil frequencies were not different between the groups (SI Figures 3C,E).

To compare the immune status of helminth-infected individuals and uninfected control subjects, PBMC from S. stercoralis-infected donors and healthy endemic controls were stimulated with PMA/ionomycin and assessed for markers of Th1 and Th2 differentiation by flow cytometry. The S. stercoralis-infected group displayed significantly elevated frequencies of CD3⁺CD4⁺GATA-3⁺ Th2 cells compared to the uninfected control group (Figures 4A,B) and a higher ratio of Th2:Th1 cells (SI Figure 4). While frequencies of IL-4 and IL-5 expressing T cells were similar in both cohorts, the S. stercoralis-infected group displayed significantly increased levels of IL-13 expressing cells (Figure 4B). Linear regression analysis confirmed a significant effect of the group in terms of IL-13 levels (p = 0.0407) and showed that increased GATA-3 expression was associated with increased IL-13 expression (p = 0.0279) (SI Figure 5A). Furthermore, levels of IL-4, IL-13 and IFN-γ significantly increased with age (p = 0.0030, p = 0.0062, and p = 0.0407, respectively, data not shown).

No differences were observed for Th1 cells expressing T-bet and IFN-γ between the cohorts (Figure 4C), but increased T-bet expression levels were associated with higher IFN-γ expression (p < 0.0001) (SI Figure 5B).

Hence, S. stercoralis infection in humans is marked by Th2 differentiation, while frequencies of Th1 cells are indistinguishable from healthy controls.

We next asked whether hybrid Th2/1 cells sharing characteristics of Th2 and Th1 cells were detectable in blood samples of the study cohorts. We detected CD3⁺CD4⁺ T cells distinctly co-expressing IL-4 and IFN-γ in the infected as well as the healthy control group (Figures 4A,D). Furthermore, the S. stercoralis-infected cohort displayed significantly increased frequencies of CD3⁺CD4⁺ T cells co-expressing GATA-3 and T-bet compared to healthy controls (Figure 4D). Expectedly, IL-4 single producing cells clustered in the GATA-3⁺ population, while some, but not all IFN-γ⁺ cells expressed elevated levels of T-bet (Figure 4E). IL-4⁺IFN-γ⁺ Th2/1 hybrid cells, however, featured a GATA-3^lowT-bet^high phenotype and clustered with the fraction of IFN-γ⁺ cells expressing the highest levels of T-bet (Figure 4E). Investigating the proportions of IFN-γ⁺ cells within the total IL-4⁺ population confirmed that the proportions of Th2/1 cells were similar in uninfected controls and S. stercoralis infected patients, hence the ratios of Th2:Th2/1 cells were similar in both cohorts (Figure 4F). However, higher values of conventional Th2 cells (IL-4+, IL-13+ or IL-5+) correlated with increased Th2/1 hybrid cell frequencies co-expressing IFN-γ and the respective Th2 cytokines (p < 0.0001) irrespective of the Strongyloides infection status (SI Figure 6).

Assessing the expression levels of GATA-3, T-bet, IL-4 and IFN-γ by mean fluorescence intensities (MFI) confirmed that human IL-4⁺IFN-γ⁺ Th2/1 hybrid cells expressed significantly lower levels of GATA-3 and IL-4 than IL-4 (or IL-13, not shown) single producers (Figure 4G). The mean expression levels of T-bet and IFN-γ, however, were similar for IFN-γ single and IL-4⁺IFN-γ⁺ double producing cells (Figure 4G).

Taken together, CD4⁺ T cells with a Th2/1 hybrid phenotype marked by the co-expression of IL-4 and IFN-γ were detectable in Strongyloides-infected patients and healthy controls. The frequencies of Th2/1 cells were positively influenced by the magnitude of the overall Th2 response. In contrast to murine Th2/1 hybrids expressing intermediate levels of GATA-3, IL-4 and IFN-γ (Figure 3), human Th2/1 hybrid cells were marked by high IFN-γ production and low GATA-3 expression.

Next, we assessed if S. stercoralis infection was marked by changes in the serum antibody profile. The infected cohort displayed significantly increased total levels of the Th2-associated antibody classes IgE and IgG4 compared to the uninfected control group (Figures 5A,B). IgE levels were positively correlated with IL-13 levels (p = 0.0011, data not shown). Interestingly, also the level of IgG3 was significantly increased in the S. stercoralis-infected group, while the levels of IgG1, a subclass associated with Th1 responses in humans, were similar in both cohorts (Figures 5C,D).

T helper cells not only produce effector cytokines in response to T cell receptor (TCR) triggering by their cognate antigen, but may also react to combinations of IL-1 cytokine family members and STAT activators (Guo et al., 2012). Th2 cells have been shown to produce IL-5 and IL-13 in response to the combined stimulation by IL-33 and IL-7 (Guo et al., 2015), while stimulation by IL-18 and IL-12 results in IFN-γ production by Th1 cells (Robinson et al., 1997; Yoshimoto et al., 1998). We thus asked if Th2/1 hybrid cells induced in helminth infections share innate effector functions with Th2 and Th1 cells. To this end, we isolated CD4⁺ T cells from mice acutely infected with S. ratti to compare their TCR-independent and TCR-induced effector responses. In response to IL-33 and IL-7, both Th2 cells and Th2/1 cells failed to produce significant amounts of IL-13 (Figures 6A,B) despite the expression of the IL-33 receptor ST2 (Figure 6C). After stimulation with IL-18 and IL-12, we detected similar IFN-γ responses of by Th1 and Th2/1 hybrid cells (Figures 6A,D).

We next compared in vitro generated murine Th1 and Th2/1 cells for their efficiency in macrophage activation. As shown previously (Peine et al., 2013), naïve cells stimulated in presence of IL-4, IFN-γ and IL-12 differentiated into a homogenous population of Th2/1 hybrids co-expressing GATA-3 and T-bet, many of which co-produced IL-4 and IFN-γ in response to re-stimulation (Figures 7A,B). Similar to Th2/1 cells induced in vivo (Figure 3), in vitro-generated Th2/1 hybrid cells produced significantly lower levels of IFN-γ than Th1 cells (Figure 7C), despite similar expression levels of T-bet (Figure 7A). In co-cultures with peritoneal macrophages, spontaneous IFN-γ secretion by Th1 cells triggered similar levels of nitric oxide (NO) production by macrophages as induced by IFN-γ added exogenously (Figure 7D). Th2/1 hybrid cells induced approximately 10-fold lower NO-responses (Figure 7D), consistent with their lower IFN-γ production upon stimulation (Figure 7C), while classic Th2 cells failed to trigger an NO response (Figure 7D). However, when macrophages where additionally stimulated with lipopolysaccharide (LPS), the limited amount of IFN-γ produced by Th2/1 cells was sufficient to significantly enhance NO-production over the levels produced in response to the LPS-trigger alone (Figure 7E).

In conclusion, Th2/1 hybrid cells induced in helminth infection are reactive to cytokine triggers leading to TCR-independent IFN-γ production. IFN-γ produced by Th2/1 hybrid cells is sufficient for the classical activation of macrophages under appropriate conditions.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.