Head-to-head comparisons of Toxoplasma gondii and its near relative Hammondia hammondi reveal dramatic differences in the host response and effectors with species-specific functions

Summary

Toxoplasma gondii and Hammondia hammondi are closely-related coccidian intracellular parasites that differ in their ability to cause disease in animal and (likely) humans. The role of the host response in these phenotypic differences is not known and to address this we performed a transcriptomic analysis of a monocyte cell line (THP-1) infected with these two parasite species. The pathways altered by infection were shared between species ~95% the time, but the magnitude of the host response to H. hammondi was significantly higher compared to T. gondii. Accompanying this divergent host response was an equally divergent impact on the cell cycle of the host cell. In contrast to T. gondii, H. hammondi infection induces cell cycle arrest via pathways linked to DNA-damage responses and cellular senescence and robust secretion of multiple chemokines that are known to be a part of the senescence associated secretory phenotype (SASP). Remarkably, prior T. gondii infection or treatment with T. gondii-conditioned media suppressed responses to H. hammondi infection, and promoted the replication of H. hammondi in recipient cells. Suppression of inflammatory responses to H. hammondi was found to be mediated by the T. gondii effector IST, and this finding was consistent with reduced functionality of the H. hammondi IST ortholog compared to its T. gondii counterpart. Taken together our data suggest that T. gondii manipulation of the host cell is capable of suppressing previously unknown stress and/or DNA-damage induced responses that occur during infection with H. hammondi, and that one important impact of this T. gondii mediated suppression is to promote parasite replication.

Keywords:

Cell cycle and cell division – Parasitic diseases – Toxoplasma gondii – DNA damage – Host cells – Sporozoites – Parasitic cell cycles – Parasite replication

Introduction

A successful infection in T. gondii results in rapid replication of infecting parasites as tachyzoites, dissemination to a variety of tissues, followed by the clearance of actively replicating stages and encystment of bradyzoite stages in muscle and neuronal tissues. Therefore the ability of T. gondii to suppress host responses are important to promote its rapid growth and dissemination during the acute phase such that a sufficient number of parasites can ultimately convert to the cyst stage and therefore be transmitted to the next host.

A critical feature of T. gondii infection is the ability to alter fundamental biological processes of the host cell. T. gondii causing changes in multiple pathways including those responsible for the regulation of metabolism [1] and innate immune signaling [2], and many of these are known to be important for parasite replication and dissemination during the acute phase. Many of the transcriptional changes that occur during T. gondii infection depend on the secretion of specific effectors directly into the host cell [1, 3, 4], and T. gondii is likely to harbor hundreds of such effectors [5, 6]. Rhoptry-derived proteins such as ROP16 regulate host transcription factors like STAT3 [7] and STAT6 [8], while dense granule proteins GRA15 [9], GRA16 [10] and GRA24 [11] modulate host NF-κB, p53 and p38 MAP kinase host pathways, respectively. GRA18 modulates the Wnt signaling pathway in mouse cells, owing to its ability to stabilize the transcription factor β-catenin [12], and GRA25 induces CCL2 production by human foreskin fibroblasts [13]. The parasite effector TgIST represses the interferon (IFN)-γ response by recruiting the Mi-2/NuRD repressor complex and subsequently blocking STAT1-related IFNγ-stimulated transcription [14, 15]. For the dense granule-derived effectors, many require the presence of a protein complex on the parasitophorous vacuole membrane (PVM) consisting of at least three proteins named MYR1, 2 and 3 [16, 17]. This complex has also been shown recently to be critical for the induction of the chemokine CCL22 in human placental cells [18]. Importantly T. gondii has no impact on CCL22 production by human foreskin fibroblasts (HFFs) [18], further demonstrating that responses to T. gondii infection can vary dramatically across cell types. Taken together it is clear that multiple means of modulating the host cell exist in T. gondii.

An unanswered question is how important these host manipulations are in determining the many unique life cycle characteristics of T. gondii compared to its near Apicomplexan relatives, including the lack of an obligately heteroxenous life cycle and the ability to infect an uncharacteristically large number of intermediate hosts. As a means to address this question we have turned to the nearest extant relative of T. gondii, Hammondia hammondi as a parasite lineage that is naturally deficient in the aforementioned life cycle features. H. hammondi was first discovered in the feces of a cat in Iowa, USA in 1975 [19], and while morphologically indistinguishable from T. gondii, H. hammondi has a comparatively restricted natural intermediate host range (being incapable of infecting birds) [20, 21]. Prior work shows that H. hammondi shares >95% of its ~8000 genes in near perfect synteny with T. gondii and expresses functional orthologs of key T. gondii effectors such as ROP18 and ROP5 [22, 23]. This is important from an epidemiological and evolutionary perspective, as H. hammondi is not known to cause clinical disease in any naturally infected intermediate (human and other animals) or definitive host [20]. In addition, in tissue culture H. hammondi replicates for a short period of time and then enters into a unique terminally differentiated bradyzoite state that is a) unable to be subcultured in vitro, b) incapable of infecting an intermediate mouse host and c) only capable of infecting a feline host [20, 23]. Paradoxically, despite this natural bradyzoite developmental program and expression of canonical bradyzoite genes during tachyzoite-like replication [23, 24], H. hammondi cannot be induced to form bradyzoites using stresses like high pH medium that robustly induce cyst formation in T. gondii [23]. While it is not yet known if H. hammondi is capable of infecting humans, and there are no tests as yet capable of distinguishing these closely-related species serologically [25], H. hammondi and T. gondii oocysts are known to co-circulate, suggesting that humans are likely to encounter this parasite [26].

Since multiple T. gondii effectors modulate host transcriptional regulation, and to date many of these have been found to be functionally conserved between T. gondii and H. hammondi [22, 24], we sought to test the hypothesis that differences in the response of the host cell to T. gondii and H. hammondi could be linked to phenotypic differences between these species, including replication rate, life cycle flexibility and pathogenesis. To do this we performed the first thorough analysis of the cell-autonomous host response to H. hammondi and compared it to T. gondii. We found that the majority of infection-induced changes in the host cell were conserved between these two species, but also that the magnitude of these changes in H. hammondi-infected cells was much larger (often by many-fold) than in T. gondii-infected cells. We confirmed that this effect is manifested at the protein level for a subset of chemokines, and that a similar effect could be observed in vivo during mouse infections. In addition to these qualitatively shared pathways, we also identified a small, but important, subset of host cell pathways that were altered during T. gondii infection but were either unaltered or even inverted in H. hammondi-infected cells. Some of these transcriptional responses indicated that H. hammondi-infected cells were in a distinct cell cycle state compared to T. gondii. Remarkably we also found that prior infection with T. gondii suppressed host responses to H. hammondi and that this suppressive effect was partially dependent on the T. gondii effector IST [14, 15] and that the molecular mechanism for this difference was specific polymorphisms in the TgIST and HhIST coding sequence. Finally we found that prior exposure of non-immune cells to media conditioned with T. gondii-infected THP-1 cells improved growth conditions for H. hammondi, linking observed differences in the host response to affecting parasite replication rate in neighboring cells. Taken together these data indicate a high level of conservation of host responses to T. gondii and H. hammondi, but extensive divergence in their abilities to target specific host cellular pathways that can be linked to polymorphisms in secreted effectors.

Results

H. hammondi sporozoites induce significantly more potent gene expression changes in THP-1 cells as compared to T. gondii

To compare the impact of T. gondii and H. hammondi infection on the transcriptional host response during infection, we performed RNA-sequencing (seq) on THP-1 cells (a monocyte cell line [27]) infected with representatives of three major lineages of T. gondii and two H. hammondi isolates. In all cases (unless specified), we used parasites that were freshly excysted from oocysts and then grown for 24 hours in HFFs prior to being quantified and used to infect THP-1 cells in suspension.

We first used principal component analysis (PCA) to look broadly at differences and similarities in the host transcriptome across strains and species [28]. According to this analysis the first PC (PC1, 68% variance) indicated a clear separation between parasite species, while the second (PC2, 20% variance) encapsulated differences between strains of each species and/or experiments that were performed together. These data indicate that THP-1 cells infected with all three T. gondii types (TgGT1, TgME49 and TgVEG) are more similar to each other than to THP-1 cells infected with either strain of H. hammondi (HhEth1 and HhAmer; Fig 1A). Infection status (Mock vs. Infected) also was distributed across PC1, in that mock-infected cells clustered together within PC1 regardless of strain or species (Fig 1A). Importantly the two H. hammondi strains were much further separated from all mock-infected samples along this PC, suggesting 1) a distinct response by the host cells to this species and/or 2) more potent induction of transcriptional changes by the parasite. PC2 showed a clear separation between TgGT1-infected cells and all other samples, including T. gondii and H. hammondi. While it was clear from PCA and other analyses (below) that TgGT1 infection induced extensive changes in transcript abundance compared to mock-infection (Fig 1A, 1C and 1H), the TgGT1-infected cells were clearly the most distinct T. gondii strain along PC2 due to strain differences and/or variation between experiments (TgME49 and TgVEG infections were performed together alone with HhEth1 and HhAmer respectively, while TgGT1 was assayed in a separate experiment. Hierarchical clustering of the distances between samples also further confirmed the differential expression profiles seen in THP-1 cells infected with H. hammondi as compared to T. gondii (Fig 1B).

We next compared host gene expression of parasite-infected cells to mock-infected cells for all types/strains using DESeq2 [28] with the thresholds of log₂ fold-change ≥ 1 or ≤ -1 with P_{adjusted (adj)} < 0.01 and identified host transcripts that were of higher or lower abundance after infection (S1 Table). A striking difference was observed in the host transcriptional response to H. hammondi compared to T. gondii, regardless of strain. Specifically, HhEth1 and HhAmer-infected THP-1 cells had significant changes in ~27% and ~32% of the 15452 queried transcripts, respectively (Fig 1I and S1 Table), while in T. gondii this ranged from ~4% for TgGT1 to ~7% for TgVEG (Fig 1H and S1 Table). MA-plots showing average expression vs. log fold-change of each gene further illustrate the stark contrast in the quantitative impact that infection by T. gondii and H. hammondi has on the host cell transcriptome (Fig 1C–1E vs. 1F and 1G). We also identified strain-specific gene expression profiles for T. gondii (as expected based on extensive comparative transcriptomics between members of the type I, II and III lineages) and to a lesser extent, H. hammondi (Fig 1H and 1I).

Hyperinduction of inflammatory cytokines by H. hammondi compared to T. gondii is recapitulated in vivo

While mice are critical intermediate hosts for both T. gondii and H. hammondi, little is known about differences in the host response to T. gondii and H. hammondi in vivo. Therefore we infected mice intraperitoneally with parasites of each species, collected peritoneal lavage supernatants and peritoneal cells, and quantified cytokine transcript and protein levels. Since H. hammondi replicates ~4X slower than T. gondii in vitro [23], we also monitored relative replication rates and parasite loads using species-specific RT-qPCR for the parasite GRA1 gene (as in [23]; S1 Fig). Overall we observed significantly higher levels of host cytokine transcripts and secreted protein in T. gondii- compared to H. hammondi-infected mice. However H. hammondi parasite burden was significantly lower than T. gondii in vivo (S1 Fig) indicating that H. hammondi replicates more slowly than T. gondii in vivo consistent with its slower in vitro growth phenotype [23]. To compensate for this growth difference we set up an independent experiment where we normalized cytokine transcript and protein level to GRA1 transcript level, a reasonable proxy for parasite burden [23]. We observed similar cytokine responses in this experiment as the first, and when we normalized cytokine transcript levels (specifically Cxcl10 and Ccl22) in peritoneal cells to GRA1 transcript level we found that per parasite H. hammondi infection resulted in significantly higher levels of transcript at 30 min, 20 and 48 h post-infection for Cxcl10 and 20 h post-infection for Ccl22 (Fig 2A and 2B). We also observed significantly higher levels of Cxcl10 and Ccl22 protein in peritoneal lavage fluid from H. hammondi-infected mice compared to T. gondii (Fig 2C and 2D). These data suggest that the more robust proinflammatory response to H. hammondi compared to T. gondii also occurs in vivo.

Pathways altered by T. gondii and H. hammondi are mostly shared, with a small number of critical differences

While it was clear that H. hammondi-infected cells displayed a more robust transcriptional change than T. gondii-infected cells (Fig 1), this could occur by differential induction of distinct sets of genes, and/or via differences in the overall magnitude of the induction of the same gene sets. To assess this we used Gene Set Enrichment Analysis (GSEA; [29]) on each transcriptional profile using the curated “Hallmark” gene sets database. Overall we identified 47 gene sets that were significantly enriched in cells infected with T. gondii and/or H. hammondi (compared to mock-infection; FDR-q value<0.05), and 41 of these were shared between species (Fig 3A and 3B, S1 Table). Interestingly, H. hammondi infections induced quantitatively higher enrichment of these shared transcriptional changes. For example, for the IFNγ response gene set H. hammondi Eth1 and Amer had normalized enrichment scores (NES) of 11.2 and 11.4 respectively while TgGT1, TgME49 and TgVEG-infected cells had and NES of 8.0, 8.6 and 10.1, respectively (Fig 3A, S1 Table). This is perhaps most simply illustrated by a clustered heatmap containing of a subset of the IFNγ response gene set, where infection by all strains and species results in increased transcript abundance but the magnitude of that induction is highly species-specific (Fig 3C, S2 Table and S2C Fig).

We used Ingenuity Pathway Core Analysis (IPA) to identify additional pathways and upstream regulators driving species-specific transcriptional changes. We identified 280 canonical pathways significantly enriched in T. gondii and/or H. hammondi infections (p <0.01, S3 Table) compared to mock-infected cells including dendritic cell maturation, IL-1 signaling, IL-6 signaling and interferon signaling (S2A and S2B Fig). Our IPA analysis reaffirm our GSEA results that while H. hammondi infection alters many of the same canonical pathways and gene sets in THP-1 cells as does infection with T. gondii, the magnitude of the effect is greater (e.g., dendritic cell maturation had z-scores of 6.38 and 6.20 for HhAmer and HhEth1 respectively; 5.57, 4.69 and 4.09 for TgGT1, TgME49 and TgVEG respectively; S2A Fig, S3 Table). This could be due to 1) T. gondii suppression of these host responses and/or 2) direct induction of a more dramatic host response during infection by H. hammondi. Since IFNγ-signaling is critical for controlling T. gondii infection [30–32] we examined all members of the interferon signaling pathway and identified a subset of them (BAK1, 1FITM1, 1FITM2, JAK1, JAK2 and PSMB8) that were of significantly higher abundance in H. hammondi-infected THP-1 cells but were comparatively unchanged during infection with T. gondii (S2B Fig). Clearly H. hammondi induces a more robust host response and/or fails to suppress the activation of this and other pathways compared to T. gondii.

T. gondii and H. hammondi-infected cells have transcriptional profiles indicating distinct cell cycle states

Only two gene sets, Myc targets v1 and v2, were significantly enriched in all T. gondii-infected cells regardless of strain type and either negatively enriched or not significantly enriched H. hammondi-infected cells (Fig 3A and 3B; S3C Fig, bottom panels). Given the fact that T. gondii is known to induce c-Myc translocation into the nucleus and also induces S phase transition in human host cells [33, 34], we wanted to determine if the lack of induction in the Myc targets v1 and v2 gene sets in H. hammondi-infected cells reflected an inability of this species to manipulate cell cycle gene expression and ultimately progression through the cell cycle. We therefore looked specifically at transcript levels for a number of cell cycle and MYC target genes in T. gondii- and H. hammondi-infected cells and found multiple E2F-family related genes were of lower abundance in H. hammondi-infected cells compared to T. gondii-infected cells (e.g., E2F1, E2F2 and MCMs 4, 5, 6 and 7; Fig 4A and 4B and S2 Table). This observation is consistent with the lower absolute NES values (for negative enrichment) in H. hammondi infection compared to T. gondii strains for the G2M Checkpoint and E2F Targets gene sets (Fig 3A, S1 Table), and suggests that H. hammondi-infected cells may be in a different cell cycle state. Consistent with this idea, we found that that H. hammondi-infected cells had significantly higher transcript abundance for GADD45A, B and G (genes involved in DNA damage-induced growth arrest [35, 36]) and CDKN1A and CDKN1C compared to T. gondii-infected cells (Fig 4A), suggesting that H. hammondi-infected cells may have a phenotype consistent with DNA damage-induced cell cycle arrest. A role for activation of DNA-damage-mediated pathways is also supported by IPA, in which we identified 38 H. hammondi-specific pathways including DNA damage-induced 14-3-3σ signaling which contains multiple DNA damage-induced cell cycle checkpoint-related genes [37] (S2B Fig and S3 Table; highlighted in blue). Combined with the differential MYC targets gene sets that were suppressed during H. hammondi infection compared to T. gondii (Figs 3A and 4B), these data suggest that T. gondii and H. hammondi-infections may have a divergent impact on the cell cycle of the host cells that they infect. While T. gondii induces progression through S phase and into G₂/M, H. hammondi infection may lead to cell cycle arrest (in either G₁/S or G₂/M) via DNA damage-related stress responses.

Upstream regulator analysis reveals differential regulation of cell cycle-related transcription factors by H. hammondi infections

Using IPA upstream regulator analysis, we identified cascades of upstream transcription regulators that might be responsible for driving the observed transcriptional changes after infection with T. gondii and H. hammondi. We shortlisted upstream transcription regulators in response to H. hammondi or T. gondii infection (compared to mock-infected THP-1 cells) using |z scores| ≥ 2 and Fisher’s exact test, P <0.01. We identified FOXM1 as a significant T. gondii-specific upstream transcription regulator (average Z-score in T. gondii of 2.31 compared to -0.79 in H. hammondi), and CDKN2A as a H. hammondi-enriched upstream regulator (average Z-score in T. gondii of 1.96 compared to 5.08 in H. hammondi; Fig 4C). From the analysis we also identified type/strain-specific and common transcription regulators activated or inhibited in T. gondii and/or H. hammondi infections (Fig 4C, S4 Table). Common upstream transcription regulators included many well-known signaling mediators involved in T. gondii infection (e.g. multiple IRFs and STATs including IRF1, 3 and 5 and STAT1) as well as the NFκB p65-encoding gene RELA (S4 Table).

H. hammondi and T. gondii-infected THP-1 cells are in distinct phases of the cell cycle

Having shown that T. gondii and H. hammondi differentially alter cell-cycle related genes, we analyzed cell cycle progression of sporozoite-infected cells using flow cytometry. Consistent with the literature [33, 34], TgVEG-infected THP-1 cells showed a distinct cell cycle profile compared to uninfected cells, having a significantly higher proportion of cells in G₂/M (Fig 4D). In contrast, the cell cycle profile of HhAmer-infected cells was more similar to uninfected THP-1 cells, with a marked lack of cells in G₂/M and higher numbers of cells in G₀/G₁ phase (Fig 4D). Similar results were obtained when comparing THP-1 cells infected with either RH88 tachyzoites or HhAmer sporozoites (S3A Fig). We also repeated this experiment in triplicate in U2OS cells and found that cell cycle manipulation differences between T. gondii and H. hammondi were not cell-type specific. Specifically, in contrast to T. gondii-infected U2OS cells which exhibited a prominent G₂/M peak (S3B Fig) and a significantly higher proportion of cells in G2/M compared to uninfected cells (Fig 4E), H. hammondi-infected U2OS cells failed to show this increase in G2/M during infection (S3B Fig and Fig 4E).

One mechanism of cell cycle arrest is via DNA damage responses leading to increased CDKN1A transcription leading to increased p21 signaling in the nucleus. FOXM1 is a component of the FOXM1-MMB complex which is active during G₂/M and promotes transcription of E2F target genes, while CDKN2A (and CDKN1A as described above) are both involved in DNA-damage and stress-induced cell cycle arrest via CDK inhibition [35, 38]. Therefore we investigated the possibility that the p53-p21-DREAM pathway was differentially modulated by T. gondii and H. hammondi since the DREAM transcriptional repressor regulates a large set of cell cycle-associated genes [39]. We performed GSEA on the Fischer DREAM Targets gene set [39] and found that target genes for part of this pathway were suppressed in H. hammondi infected cells compared to T. gondii (Fig 4F; NES = -13.12, FDR-q = 0). Given the known impact of cell cycle arrest on T. gondii replication rate [33], and the fact that H. hammondi replicates slower than T. gondii in vitro [23] and in vivo (Fig 2), we postulated that p21-mediated cell cycle arrest might play a role in determining T. gondii and H. hammondi replication rate. To test this hypothesis we treated HFFs with 4 μM cisplatin (to induce p21 expression) or vehicle and quantified parasite growth. While cisplatin treated cells had significantly higher nuclear p21 staining compared to vehicle-treated cells (p = 0.01; Fig S4C and S4G), T. gondii infected and proliferated within cisplatin and vehicle-treated cells equally (P>0.05; Fig 4G), indicating that p21 upregulation alone was not sufficient to alter T. gondii replication. These data suggest that the differences in replication rate between T. gondii and H. hammondi in HFFs are unlikely due to differences in p21 level.

Of the chemokines that we quantified at the transcriptional level using RNAseq and the protein level in supernatants of infected THP-1 cells using Luminex Bead Arrays, CXCL10 stood out as having the most H. hammondi-specific expression phenotype and as shown in Fig 2 this response could be recapitulated in mice. Therefore we used this chemokine as a sentinel for H. hammondi-specific responses. To determine if the lack of CXCL10 induction in T. gondii VEG sporozoite-infected cells was strain- or life-stage-specific, we infected THP-1 cells with TgVEG sporozoites, RH88 tachyozites or with H. hammondi sporozoites. Consistent with TgVEG, TgRH88 also failed to induce CXCL10 secretion by THP-1 cells (Fig 5A), while H. hammondi infected cells produced ~150-fold more CXCL10 compared to cells infected with either T. gondii strain (Fig 5A). CXCL10 induction required live parasites, in that heat-killed H. hammondi failed to induce CXCL10 (Fig 5B). Since H. hammondi-infection induced genes related to DNA damage and cell cycle arrest, and since a subset of cell cycle arrest phenotypes (like quiescence regulated by the DREAM complex and cellular senescence) can be transmitted to bystander cells in a paracrine fashion, we used Transwell inserts to separate infected THP-1 cells from bystanders (Fig 5C) and quantified CXCL10, CCL22 and CDKN1A transcript in both classes of cells. We used CXCL10 and CDKN1A as sentinels for paracrine regulation of senescence and quiescence-associated phenotypes, and CCL22 as a chemokine induced by both parasite species but which is not typically associated with quiescence or senescence associated phenotypes. Since the SASP response can signal other cells in a paracrine manner, we used CXCL10 secretion as a sentinel chemokine for SASP. We found that bystander cells from H. hammondi-infection had significantly higher CXCL10 transcript as compared to those from mock infection (by ~10-fold; Fig 5D), and RT-qPCR for H. hammondi GRA1 transcript confirmed the absence of parasites in the bystander cells (S4 Fig). In contrast to CXCL10, we did not observe any increase in transcript abundance for CCL22 and CDKN1A in bystander cells (Fig 5D). CXCL10 was detected in supernatants in both conditions, since this chemokine could freely diffuse after its secretion. Taken together these data provide further support for the hypothesis that H. hammondi infection, and not T. gondii infection, leads to a host cell phenotype with hallmarks of quiescent and/or senescent cells capable of altering cell signaling pathways in neighboring cells.

Consistent with this observation, the Fridman Senescence Up gene set [40] was significantly enriched in H. hammondi-infected cells with an NES of 3.4 (Fig 6A; FDR-q = 0), suggesting that H. hammondi-infected cells may have some similarities to senescent cells. This would explain the observed differences in the host cell cycle as well as the increased production of inflammatory cytokines via the Senescence-Associated Secretory Phenotype (SASP) [41]. This hypothesis was further supported by IPA analyses showing significant enrichment of genes in the DNA damage-14-3-3σ signaling pathway (S2A Fig), and the IPA analysis results (Fig 4) showing that CDKN2A is an important upstream regulator of pathways uniquely altered by H. hammondi infection and conversely that FOXM1 signaling targets are enriched in T. gondii-infected cells (Fig 4). One of many defining characteristics of senescent cells is the production of lysosomal senescence-associated β-galactosidase (gal) and β-gal enzyme activity can be detected both in cells and supernatants [42]. We evaluated this in parasite-infected THP-1 cells but found no significant increases in β-gal activity in THP-1 cells infected with either parasite species, even though β-gal activity did increase in Phleomycin-treated THP-1 cells (as a control for β-gal activity; S5 Fig). However, we did find that THP-1 cells infected with H. hammondi had significantly higher levels of IP10/CXCL10, RANTES/CCL5, and MIP-1a/CCL3 transcript compared to T. gondii (Fig 6B), and this increased transcript abundance was also reflected in significantly higher levels of these chemokines in supernatants from H. hammondi-infected cells (Fig 6C and S5 Table). These chemokines are also well-known indicators of the SASP [43].

T. gondii-mediated suppression of H. hammondi-induced CXCL10 is TgIST-dependent

Next, we speculated that T. gondii might be capable of suppressing the hyperinflammatory transcriptional and cytokine secretory response by H. hammondi. To test this hypothesis we exposed THP-1 cells to TgVEG sporozoites for 4 h prior to infection with H. hammondi (controls were exposed to mock conditions as above) and used ELISA to quantify transcriptional changes across conditions. We found that TgVEG infection significantly and dramatically suppressed H. hammondi-mediated CXCL10 secretion (Fig 7A). However, this suppression only occurred when THP-1 cells were first exposed to T. gondii, since THP-1 cells pre-infected with H. hammondi sporozoites followed by T. gondii infection produced similar levels of CXCL10 (Fig 7A). In similar experiments, we harvested RNA and subjected it to RNAseq analysis, and found that the suppressive phenotype of prior infection with T. gondii on H. hammondi alterations of the abundance of transcripts encoding senescence- and DNA damage-associated gene products (including members of the GADD45 family; S6 Fig). Interestingly, suppression of H. hammondi-induced CXCL10 production also occurred when THP-1 cells were first exposed to T. gondii/THP-1 conditioned media (harvested 4 h post-infection; Fig 7B), suggesting that T. gondii-mediated suppression is due, at least in part, to parasite- and/or host cell-derived soluble mediators released during infection. However, our data clearly showed that the suppressive effect was most potent when T. gondii parasites were used as opposed to T. gondii-conditioned media. Given the dramatic differences in replication rate between T. gondii and H. hammondi sporozoites [23], we tested whether T. gondii/THP-1 conditioned media could alter the division rate of H. hammondi. Pre-treatment of HFFs with T. gondii/THP-1 conditioned media resulted in a highly significant (P = 2x10^-16) increase in H. hammondi vacuole size (Fig 7C). This change was characterized by an increase in the number of 2- and 4-parasite vacuoles (from 38 to 50% and 14 to 28%, respectively; Fig 7C). In contrast, when we pre-treated HFFs with H. hammondi/THP-1 conditioned media, we observed only a small, but significant (P = 0.005; Fig 7D) decrease in vacuole size compared to mock-treated controls. Overall these data further show that T. gondii and H. hammondi infection induce distinct cellular responses with the capacity to have distinct effects on bystander host cells. Perhaps more importantly they suggest that manipulation of the host cell by T. gondii may play a role in determining replication rate itself and that this could be exploited to improve cultivation methods for H. hammondi.

Given the impact of prior T. gondii infection on CXCL10 secretion we hypothesized that the suppressing factor might be a T. gondii-secreted protein. Since CXCL10 secretion is induced by IFNγ treatment in a variety of cell types (CXCL10 is also known as interferon-γ-inducible protein 10 or “IP-10” [44]) and T. gondii IST is known to block IFNγ-induced responses during infection [14, 15], we performed multiple assays with T. gondii strains lacking TgIST. We found that compared to wild type T. gondii (RH strain), prior infection with T. gondii ΔIST parasites was significantly less effective at suppressing H. hammondi-mediated CXCL10 production (Fig 7E). In a separate experiment, we compared the ability of TgΔIST and the same strain complemented with a wild type copy of T. gondii IST (TgΔISTcmp) to suppress H. hammondi-induced CXCL10 secretion. Again, we found that THP-1 cells preinfected with TgΔIST parasites secreted significantly more CXCL10 after H. hammondi infection than those pre-infected with TgΔISTcmp (Fig 7F; P<0.0001). While there may be other effectors produced by T. gondii that are also capable of suppressing CXCL10 and other IFNγ-mediated responses, these data show that the IST effector plays a key role in this phenomenon.

H. hammondi fails to suppress IFNγ-induced IRF1 activation and has a significantly less functional IST gene compared to T. gondii

Given that T. gondii IST plays a role in suppressing H. hammondi-induced CXCL10 production, we hypothesized that this might be due, at least in part, to deficiencies in the function of H. hammondi IST. TgIST is required for T. gondii to suppress IFNγ-induced IRF1 nuclear translocation [14, 15], and so we first quantified IRF1 nuclear staining in IFNγ-treated HFFs infected with H. hammondi, T. gondii, or Neospora caninum. N. caninum lacks a clear ortholog of TgIST and has been shown previously to be incapable of suppressing IFNγ-induced signaling [15, 45]. As expected, pre-infection of HFFs with TgRH88 tachyzoites or TgVEG sporozoites significantly suppressed IRF1 activation in response to IFNγ treatment (P<0.05; Fig 8A and 8B), while prior infection with N. caninum tachyzoites did not (Fig 8A and 8B). When the same experiment was performed with H. hammondi sporozoites, prior infection failed to suppress IFNγ-induced IRF1 activation (Fig 8A and 8B), suggesting that sequence differences in the IST locus may determine species-specific differences in the host response. Comparison of the TgIST (TGME49_240060) and HhIST (HHA_240060) predicted protein sequences using BLASTP shows multiple gaps in the alignment and significant differences in the predicted size of the amino acid sequence (Fig 8C). While the size difference may be an artifact of sequence assembly breakdown in repetitive sequences found in the C-terminus of HhIST (there is a sequence assembly gap in the C-terminus of HHA_240060; Fig 8C; ToxoDB.org), the overall level of identity between TgIST and HhIST in the aligned regions is low (68% for both regions; Fig 8C) compared to the predicted proteome-wide average of ~89%. Moreover, H. hammondi IST lacked any significant predicted nuclear localization sequences, while TgIST harbored multiple predicted NLS’ (3 with a posterior probability >0.5 and one with a posterior probability >0.4; red and pink boxes, respectively; NLStradamus 4 state hidden markov model; [46]). Alignments of the first two predicted NLS sequences between TgIST and HhIST show extensive polymorphism, suggesting that one functional difference between TgIST and HhIST was a lack of NLS targeting sequences. To test this directly, we expressed Ty-tagged forms of TgIST, HhIST and HhIST with a C-terminal NLS in U2OS cells. As expected, based on Ty-tag staining TgIST trafficked primarily to the nucleus of transfected cells and effectively suppressed IFNγ-induced IRF1 activation (Fig 8D, top). In contrast, HhIST could be found in the nucleus and cytoplasm of the transfected cell (Fig 8D, bottom). When we quantified IRF1 staining in transfected cells and compared it to untransfected cells in the same field of view, we found that HhIST was less effective at blocking IFNγ-induced IRF1 compared to TgIST (P = 0.01; Fig 8E). Adding a C-terminal NLS to HhIST partially reversed this deficiency, as HhIST-NLS-expressing host cells had nuclear IRF1 staining intensity that was not significantly different (P = 0.9) from TgIST (Fig 8E). These data implicate sequence differences in HhIST in determining deficiencies in the inability of this parasite species to suppress host innate immune responses, and suggest that differences in nuclear localization of IST in the host cell may be at least partially responsible for species-specific differences in the ability to suppress IFNγ-induced signaling.

Discussion

It is well known that T. gondii tachyzoites activate a potent host immune response. Despite that, once inside a host cell, T. gondii is able to modulate, survive and even evade immune responses, all of which lead to replication and ultimate dissemination to distant host tissues. This has largely been attributed to the powerful strategy that T. gondii employs to co-opt host gene expression [10, 47] and to protect itself from host proteins designed to destroy the parasite-containing vacuole [48, 49]. In contrast little is known about the impact of H. hammondi on the host cell, and whether its comparatively “avirulent” lifestyle can be attributed at all to differences in the host response. Many of the unique aspects of H. hammondi biology, including its comparatively slower replication rate, spontaneous (and terminal) cyst formation and inability to be lethal even in IFNγ knockout mice [20], suggest that it is engaged in an inflexible developmental program once sporozoites are released from the oocyst. In the extreme, then, one could hypothesize that the nature and magnitude of the host response may be completely irrelevant to the outcome of infection with H. hammondi. However our data provide strong evidence to the contrary.

Our study is the first to compare host responses to sporozoites of T. gondii and H. hammondi. This particular life stage can be a challenging experimental subject, and batch-to-batch variability in excystation rates and the viability of the obtained sporozoites are typical and have been reported previously [23, 50]. This can likely explain observed differences in the magnitude of host responses to infection as well as differences the T. gondii-mediated suppressive phenotype (e.g., Fig 7E versus 7F). However, care was taken to use oocysts within 6 months of purification from cat feces and to compare sporozoites to sporozoites whenever possible (e.g., for all experiments involving comparisons between T. gondii VEG and H. hammondi).

At first glance, our comparisons of the transcriptomes T. gondii and H. hammondi-infected host cells indicate that these parasites modulate the host cell using qualitatively similar strategies. Over 95% of the underlying altered pathways were shared between T. gondii and H. hammondi, and fell into categories like inflammation, apoptosis, cell growth, and metabolism that have long been known to be targeted by T. gondii [1, 47] (Fig 3). However when the magnitude of the response was taken into account (whether measured by normalized enrichment scores from GSEA or cytokine secretion), it is clear that H. hammondi-infected cells are altered more robustly compared to those infected with T. gondii. This effect, which is also recapitulated in an animal model of infection when differences in parasite burden is taken into account (Fig 2), suggests either that a) H. hammondi has fewer countermeasures to counteract the host response than T. gondii or b) H. hammondi has a factor or factors that cause hyperactivation of the immune response (via secreted effectors, and/or production of pathogen- or damage-associated molecular patterns). Regardless of the root causes (which are likely to be complex), the rapid proinflammatory response to H. hammondi that is not induced and/or suppressed during T. gondii infection is likely to contribute to dissemination differences between these organisms (e.g., the lack of significant brain tropism in H. hammondi; [20]).

Intriguingly our results show that the most obvious differential host response in THP-1 cells infected with T. gondii and H. hammondi is the regulation of cell cycle (Figs 3, 4 and S4), a phenomenon long-described in T. gondii but whose role in determining infection outcome is unknown. Unlike T. gondii infection that actively induces host c-Myc [16], the MYC targets v1 and v2 gene sets were conversely suppressed in THP-1 cells infected with H. hammondi (Figs 3 and 4). T. gondii infection leads to an accumulation of cells in the G₂/M phase [33, 34] and in our data this was accompanied by increased transcript abundance of FOXM1-targeted genes (Fig 4; [38]). In contrast, H. hammondi leads to increased transcription of multiple GADD45 genes which block cell cycle progression as a part of the DNA damage response [35, 38]. DNA damage can induce p53 activation and increase transcription of CDKN1A [51], and we observed this outcome only in H. hammondi-infected cells (Fig 4A). In addition, our data showed H. hammondi-mediated suppression of the p53-DREAM pathway target gene set which could lead to host cell G1/S cell cycle arrest (Fig 5A; [52]). The precise impact of the robust activation of the P53 signaling pathway on H. hammondi replication in vitro is unknown, since experimental manipulation of p21 levels did not alter T. gondii replication.

Multiple T. gondii effectors have been implicated in modulating the host cell cycle and levels of host cell cycle regulating effectors like P53 and p21. These include GRA16 [10], which alters P53 levels and parasite virulence, and HCE1/TEEGR [53, 54] which interacts directly with host cyclins and modulates the host cell cycle. The fact that multiple T. gondii effectors target these overlapping host cell pathways highlights the importance of the replication status of the host cell during T. gondii infection. The fact that H. hammondi fails to modulate the host cell cycle in the same fashion as T. gondii suggests that this may be a derived trait. It is likely that there are functional differences between T. gondii and H. hammondi paralogs of GRA16 and HCE1/TEEGR, and now that forward genetics approaches are viable in H. hammondi [23] this can be tested directly through gene knockouts and/or cross-species complementation experiments.

DNA damage responses can have dramatic effects on cell biology that are independent of cell cycle arrest, and one of these is the Senescence Associated Secretory Phenotype (SASP; [41–43]). We observed increased production of multiple chemokines typically associated with the SASP, including CXCL10 which not only was uniquely present in H. hammondi-infected supernatants but was also activated in bystander cells via paracrine signaling (Fig 5). We did not detect elevated β-gal activity in host cells infected with H. hammondi, cell types and different cellular conditions could also affect the reliability of any types of senescence markers [41–43]. It is intriguing to postulate that the manipulation of the host cell cycle by T. gondii, and specifically its ability to induce S-phase transition and ultimately G₂/M arrest, is a means to prevent the host cell from becoming senescent and activating the SASP in both the infected cell and ultimately in neighboring bystander cells. Disrupting the ability of the host cell to secrete cytokines like CXCL10 would certainly be advantageous given the importance of this chemokine to maintain T-cell populations capable of controlling T. gondii proliferation [30, 55].

Similar to paracrine effects of the H. hammondi-induced SASP on bystander cells, T. gondii induces host- and infection-altering factors to be produced by host cells as well. H. hammondi has a markedly slow replication rate compared to T. gondii (shown in multiple studies including [23, 56]), but this slow rate of division appears to be flexible depending upon the host cell environment. The ability to alter the replication of an intracellular parasite like H. hammondi using T. gondii-conditioned medium (Fig 7) leads to a number of interesting biological questions about the nature and identity of the factors capable of controlling parasite replication. Supernatants from T. gondii-infected cells have been found to have a variety of effects on recipient cells that vary depending on the donor and recipient cell types (e.g., [57, 58]), although to date the factors themselves have not been identified. Our data provide further compelling evidence for their importance in determining the outcome of T. gondii infections. In addition to this interesting biology there are additional practical implications of this work with respect to the H. hammondi/T. gondii comparative system. The use of T. gondii-conditioned media may be a first step in further improving culture conditions for H. hammondi, with the ultimate goal of promoting long term in vitro culture which is not yet possible [23, 56].

Finally our data point to a key role for the effector TgIST in suppressing H. hammondi-induced host responses. While the relevance to infections in vivo is currently unknown, it suggests that there is some level of overlap between the pathways that are robustly induced by H. hammondi and those that are suppressed by T. gondii IST. Based on our data comparing the ability of T. gondii and H. hammondi IST to suppress IFNγ-induced IRF1 activation, these data also suggest that T. gondii IST has traits that evolved uniquely in this lineage, since they are not shared with H. hammondi IST and since N. caninum does not appear to express a functional form of this gene [14, 15]. The lack of predicted nuclear localization signals in H. hammondi IST as well as its increased functionality in ectopic expression assays suggest that one innovation in TgIST may have been the ability to be robustly trafficked to the host cell nucleus. This may have permitted further optimization of the TgIST gene product to robustly suppress IFNγ induced responses through its interaction with STAT1 targeted genes. This difference in functionality at the TgIST locus may explain some of the well-documented differences in virulence between T. gondii and H. hammondi. Regardless, it is possible that the lack of a fully functional HhIST gene in H. hammondi fits with its developmental program. Unlike T. gondii, H. hammondi fails to alter its developmental trajectory in the host based on its immune status, and therefore it is possible that it survives the robust immune response that it induces using a pre-programmed period of replication and development. This likely results in lower cyst burdens since it may only replicate and disseminate for 5–7 days, but H. hammondi cysts are only infectious to cats where even a single cyst can lead to the production of hundreds of millions of oocysts.

Summary

Comparisons between T. gondii and its near relatives can shed important new light on the evolutionary origins of pathogenesis in a parasite that has achieved a near global presence in humans and other animals. Here we show that in addition to having dramatic differences in replication rate and development, T. gondii and H. hammondi also differ in their abilities to manipulate the host cell. This was somewhat surprising given our prior work showing that pathogenic determinants like ROP18 and ROP5 were functionally conserved in H. hammondi [22, 24], but we have now identified at least one dense granule protein (IST) that is a molecular determinant of these differences in host cell modulation. Moreover we show for the first time a potential link between species-specific differences in host cell modulation in the intrinsic growth rate of these parasites, and have discovered that T. gondii may be inducing host cells to produce soluble factors that increase permissiveness to parasite replication in neighboring cells. These discoveries illuminate previously unknown mechanisms of host cell manipulation by T. gondii that would not have been possible to identify without the use of H. hammondi as a comparative model. Given that H. hammondi is a highly successful parasite of animals in its own right [26], the hypothesis emerging from this and our prior work is that T. gondii and H. hammondi share nearly all of their genes, but sequence-level divergence at key loci have resulted in the species-specific traits. In the case of T. gondii these traits led to the emergence of T. gondii as a highly virulent parasite of animals and humans with a global distribution and the unique ability to dynamically regulate its developmental state based on environmental cues [23].

Materials and methods

Cells

Human foreskin fibroblasts (HFFs) and human osteosarcoma bone cells (U2OS) were maintained in cDMEM (100 U/ml penicillin/streptomycin, 100 μg/ml streptomycin, 2 mM L-glutamine, 10% FBS, 3.7 g NaH₂CO₃/L, pH7.2; ThermoFisher Scientific). Human monocytes (THP-1) were maintained in cRPMI-1640 (100 U/ml penicillin/streptomycin and 10% FBS; ThermoFisher Scientific) respectively. All cells were grown at 37°C in 5% CO₂. One day prior to parasite infection, THP-1 cells growth media were replenished with 20% (v/v) of new media (cRPMI). All THP-1 cell infections were performed in cDMEM.

Mice

Balb/c mice were purchase from Jackson Laboratory and were female aged 6–8 weeks. All animal experiments were approved by the local IACUC at the University of Pittsburgh (Protocol #18032113), with euthanasia conducted according to AVMA guidelines.

Parasites

Oocysts of Toxoplasma gondii (Tg) genotype I (GT1), II (ME49) and III (VEG) and Hammondia hammondi (Hh) American (Amer) and Ethiopian-1 (Eth1) isolates were harvested from cat feces 7–11 days after feeding mouse tissues (brain for T. gondii, leg muscle for H. hammondi) infected with parasites to 10–20 week old cat free of pathogens [20, 50]. Unsporulated oocysts were isolated via sucrose floatation and placed at 4°C in 2% H₂SO₄ to encourage sporulation and for long-term storage. Rh88 tachyzoites were propagated in HFF cells. To obtain pure tachyzoites, HFF cells were washed with PBS, scraped, lysed with 25 gauge needle and filtered with 5 μM syringe-driven filter.

The following knock-out parasite strains were used in this study: RHΔIST [15], RHΔGRA16 [10] and RHΔHCE1 [54]. RHΔHCE1 and RHΔGRA16 are gifts from John Boothroyd. T. gondii RH88 and RH-YFP tachyzoites and the knock-out parasites were maintained in HFFs cultured in cDMEM. Prior to infection, monolayer with parasites was scraped and syringe lysed with a 25 gauge and a 27 gauge needle, counted with a hemocytometer and added to HFFS, THP-1 or U2OS cells at the desired multiplicity of infection (MOI). Mock-infection was performed by filtering the parasite extracts with a 0.2 μM syringe-driven filter and added the same volume of parasite-free extract to the host cells.

T. gondii and H. hammondi oocyst excystation

We performed oocyst production and excystation as described in detail in [50]. Briefly, sporulated oocysts were washed 3 X in Hank’s balanced salt solution (HBSS; Life Technologies) and treated with 10% bleach (in PBS) for 30 min with shaking at room temperature. Next, bleach was washed off using HBSS and parasites were added to 4 g sterile glass beads (180 μM; Sigma-Aldrich). Oocysts were vortexed on high speed for 15 s on/15 s off for a total duration of 2 min to disrupt the oocyst wall mechanically. Sporocysts were pelleted by centrifugation at 1,000 x g for 10 min. Pellet was resuspended in 5 ml of pre-warmed and freshly made, 0.22 μM filtered excystation media (0.1 g porcine trypsin (Sigma-Aldrich), 2 g Taurocholic Acid (Sigma-Aldrich) in 40 ml PBS, pH 7.5). Sporocysts were incubated in 37°C water bath with 5% CO₂ for 45 min and syringe-lysed using a 25 and a 22 gauge needle. H. hammondi sporocysts were syringe-lysed with a 25 gauge needle. To quench the excystation media, 7 ml of cDMEM was added. For some experiments, excysted parasites (sporozoites) were pelleted and resuspended in cDMEM and used to infect monolayers of HFFs overnight at 37°C in 5% CO₂. HFFs were then scraped, syringe-lysed and filtered with 5 μM syringe-driven filter (Millipore) to obtain pure parasites. In other experiments, freshly excysted sporozoites were used. These included all in vivo experiments, experiments analyzing induction of CXCL10 in Transwells, heat-killed parasite assays and parasite replication assays. Within an experiment, all T. gondii VEG and H. hammondi sporozoites were treated the same, and in all cases parasites were used within 24 hr of excystation.

Human monocyte cell line infection

Prior to infection, THP-1 cells were seeded at 1 x 10⁵ cells/well in 24-well plates in cDMEM. To prepare parasites for the infections, HFFs monolayers containing parasite sporozoites were scraped, syringe-lysed, and pelleted (in some cases, freshly excysted oocysts were used). After resuspending the pellet in cDMEM, the parasite mixture was filtered through a 5 μM syringe-driven filter (Millipore). THP-1 cells were infected with either T. gondii or H. hammondi at multiplicity of infection (MOI) of 4, 2 or 1.6. Three biological replicates were made for each parasite infection. THP-1 cells were also mock-infected with parasite filtered through a 0.22 μM syringe-driven filter (Millipore). These protocols are described in detail in [50].

RNA isolation

RNA was collected at 24 h post-infection or 20 h post-secondary infection (for coinfection experiments) from parasite- and mock-infected cells using the RNeasy Kit according to the manufacturer instructions (Qiagen). QIAShredder spin columns (Qiagen) were used to homogenize the samples prior to RNA extraction and contaminating DNA was degraded using RNase-free DNase (Qiagen). RNA was eluted in 50 μl of RNase-free water and gel electrophoresis and Nanadrop RNA quantification were performed to ensure RNA integrity. One mock and three biological infection replicates were done for all experiments. Total RNA samples were kept at -80°C.

mRNA-sequencing and data processing

mRNA-sequencing libraries and Illumina next generation sequencing were performed at the Core Facility at the University of Pittsburgh. Integrity of the RNA was analyzed with Agilent 2100 Bioananalyzer and all purified RNA samples had RIN scores >9. RNA was sequenced using NextSeq 550 (Illumina) and were pooled and sequenced over four lanes. Strand-specific, 150 bp, single-end RNA-sequencing was performed. Read libraries were mapped to the human genome (Homo sapiens ensemble v81; hg38) and transcriptome (Homo sapiens ensemble v81; hg38) with default options on CLC Genomics Workbench v11.0. Fastq files have been deposited in the NCBI short read archive (Accession number: SRX3734421-8).

Differential expression analysis using DESeq2 and Pre-ranked Gene Set Enrichment Analysis

DESeq2 [28] was used to perform differential expression analysis of genes in THP-1 cells infected with T. gondii and H. hammondi. Raw reads (total gene reads; exported from CLC Genomics Workbench) with at least 1 read count in all samples were analyzed. Prior to analyzing differential gene expression, integrity of the data was examined using principal component analysis (PCA) and distances of all samples were calculated (embedded in the DESeq2 package). Genes were considered to be differentially expressed in THP-1 cells if the log₂ fold-change was ≥ 1 or ≤ -1 and with a P_adj value (alpha) <0.01.

Pre-ranked Gene Set Enrichment Analysis (GSEA; [29]) was performed to compare gene sets that were enriched in THP-1 cells in relation to parasite infections. Ranked list calculating the fold-change difference (subtracting normalized log₂ fold-change of infected host cells from mock infected host cells) was used for the analysis.

Ingenuity Core Pathway Analysis

Core analysis from Ingenuity Pathway Analysis (IPA; Qiagen) was used to examine biological relevance of the RNA-seq data. Canonical pathways and upstream regulators that are over-represented in T. gondii and H. hammondi infection were analyzed using log₂ fold-change and p_adj value obtained in DESeq2 [28]. Only genes that were deemed significantly expressed were used in the analysis (log₂ fold-change ≥ 1 or ≤ -1 and p_adj value < 0.01 for Infected vs. Mock). IPA default settings were used for the analysis. For both canonical pathway and upstream regulator analysis, pathways and genes with activation z scores ≥ 2 were deemed to be activated while activation z scores ≤ -2 were deemed to be inhibited for different analysis. Threshold for p value significant used in the analysis was <0.05 or–log(p-value) >1.3.

Infected mouse peritoneal cell collection

Mouse infections were performed in two independent experiments. In the first, 40,000 sporozoites (TgVEG or HhAmer) were grown overnight in HFFs prior to being harvested by needle passage and filtering, and resuspended in PBS. Parasites were intraperitoneally injected into 7–9 week old mice. Mice were euthanized with CO2 according to IUCAC protocols and dissected. Peritoneal cells were collected in 3 ml PBS as described previously [23]. The second experiment was identical except that the dose was 20,000 per mouse, and instead of culturing parasites overnight after excystation they were used immediately.

Multiplex cytokine analysis

THP-1 cells were infected with MOI 4 of T. gondii or H. hammondi (same infection set up as the RNA-seq experiments). THP-1 cells infected or mock-infected with parasites were pelleted at 1,000 x g for 10 min. Supernatant were collected and stored at -80°C. Luminex was performed at the Luminex Core Facility University of Pittsburgh Cancer Institute. Each sample was measured (fluorescence intensity) in duplicate.

Cytokine analysis using ELISA

Concentrations of the human pro-inflammatory cytokine CXCL10 and CCL22 and mouse Ifnγ, Il12p40, Cxcl10 and Ccl22 in mouse peritoneal cell supernatants were analyzed by ELISA according to the manufacturers instructions (Human and Mouse DuoSet ELISA respectively, R&D Systems). For relative chemokine concentration in comparison to parasite load, log₂ values of the absolute concentration of chemokines was calculated and relative transcript abundance in relation to parasite burden was calculated using the 2^-ΔΔCT method (ΔΔC_T = log₂ chemokine concentration–ΔC_T GRA1).

Transwell and heat-killed parasite infections

Freshly excysted TgVEG or HhAmer was added to THP-1 cells seeded in 24-well plates (control) or added onto Transwell inserts (0.4 μM polycarbonate membrane, Corning). For heat-killed parasite infection, freshly excysted sporozoites were heat-killed at 95°C for 5 min and cooled to room temperature before infecting THP-1 cells. Multiplicity of infection of 2 was used in these experiments. Supernatants were collected either by pelleting the cells or collected from the bottom chamber of the Transwell setup.

Reverse transcriptase-quantitative PCR (RT-qPCR)

RNA were extracted using RNeasy RNA extraction kit as above and according to manufacturer’s instructions (Qiagen). cDNA was reverse transcribed from 1 μg of RNA using SuperScript IV First-Strand synthesis system (ThermoFisher Scientific). RT-qPCR was performed using QuantStudio with SYBR Green (Applied Biosystems). The PCR mixture contained 1 X SYBR Green buffer (BioRad), 0.25 μl of forward and reverse primers (S6 Table) and cDNA. Genes were amplified using the standard protocol (95°C for 10 min and 40 cycles of 95°C for 15 sec and 60°C for 1 min). Data were acquired and analyzed using QuantStudio^TM Design & Analysis Software (ThermoFisher Scientific) and data exported to Microsoft Excel for threshold values (C_T) and 2^-ΔΔCT for fold change analysis. GAPDH and Gapdh was used as the human and mouse reference gene respectively (control) for the 2^-ΔΔCT analysis. For in vivo 2^-ΔΔCT analysis, relative expression of target genes were normalized against the ΔC_T of mock-infected mice. For relative expression normalized by comparison with parasite load, we used GRA1 as a reference gene for parasite burden. To do this, the ΔC_T of target genes and GRA1 were first obtained (C_T target gene or GRA1 –C_T Gapdh). Then, ΔΔC_T target gene was calculated (ΔC_T target genes–ΔC_T GRA1). Relative expression was finally calculated using the 2^-ΔΔCT methods for respective target genes. Primers validation and melt curve analysis were performed to ensure integrity of the RT-qPCR. No RT and water controls were also included in every RT-qPCR.

Immunofluorescence

All IFAs were performed in HFFs seeded on glass coverslips. Infected cover slips were fixed with 4% paraformaldehyde in PBS for 20 min. Cover slips were rinsed once with 1X PBS and permeabilized with IFA buffer (PBS and 0.2% Triton-X). Cover slips were then blocked with the blocking buffer (PBS, 0.2% Triton-X and 5% BSA) for 30 min. All these steps were performed at room temperature (RT). CDKN1A/p21 was detected using the mouse monoclonal antibody p21 (Santa Cruz Biotechnology) either overnight at 4°C or for 1 h at RT. Secondary anti-mouse antibody conjugated to Alexa Fluor 488, or 647. Cover slips were mounted with DAPI (4’,6-diamidino-2-phenylindole) stain (Vector Laboratories). Fluorescence was detected using fluorescence microscope (Olympus) and images were analyzed using ImageJ or CellSense. The same exposure time was used for any given comparison data set. Growth of RH-YFP was determined by analyzing area of parasite growth from at least five field of views (FOV) from a single coverslip. Results were obtained from three separate coverslips.

Cell cycle analysis

Cell cycle profiles of THP-1 cells were analyzed with propidium iodide staining according to manufacturers instructions (ThermoFisher Scientific). THP-1 cells were infected with T. gondii or H. hammondi at an MOI of 2. For uninfected cells, THP-1 cells were seeded at the same time as the other cells in different treatment groups and an equal volume of media was added to the uninfected cells in place of parasite infection. At 20 h post-infection, cells were pelleted and washed 1X with chilled PBS and fixed in chilled 80% ethanol at -20°C for overnight. Cells were then washed with chilled PBS and rehydrated for 15 min with chilled PBS. Cells were stained with 2 μg/mL propidium iodide/RNaseA (ThermoFisher Scientific) solution for 15 min at room temperature with gentle agitation. Stained cells were analyzed immediately by flow cytometry using LSR II (BD Biosciences) at the United Flow Core at University of Pittsburgh or kept at -20°C in the dark for no longer than 24 hr until flow cytometry analysis. Cell cycle profiles were analyzed with ModFit LT 5.0 (VSH).

β-galactosidase activity assay

β-galactosidase activity was analyzed in THP-1 cells and secreted proteins using the β-galactosidase assay kit (ThermoFisher) according to manufacturers instructions. Briefly, THP-1 cells were inoculated with T. gondii or H. hammondi, treated with 1 μg/mL of phleomycin in cDMEM. At 72 h post-infection, cells were pelleted at 300 x g for 10 min and supernatant was collected for β-galactosidase activity assay. The cell pellets were washed 2 times with PBS and lyzed with IP lysis buffer (ThermoFisher) with Halt Protease Inhibitor Cocktail (ThermoFisher), before analyzing for β-galactosidase activity. β-galactosidase activity was determined after 1 h of incubation at 37°C and absorbance was read at 420nm.

IFNγ-induced IRF1 expression derived from parasite infection

HFFs grown on coverslips were infected with TgVEG and H. hammondi sporozoites (MOI 0.5) for 72 hours and with RH88 and N. caninum (NC-1) (MOI 0.5) for 3 hours prior to treatment with IFNγ. Infected cells were treated with recombinant human IFNγ protein (1 ng per mL of cDMEM; Gibco PHC4031) for 15 hours. Infected host cells were fixed with 4% paraformaldehyde. Coverslips were stored in blocking buffer (5% BSA, 0.1% Triton X-100) at 4°C until staining was performed. Coverslips were stained with goat Toxoplasma gondii Polyclonal Antibody (1:500) (Invitrogen PA1-7256) and rabbit IRF1 (1:200) (Cell Signaling Technology, 8478S) primary antibodies for 1 hour at room temperature. Coverslips were washed 3X with PBS and stained with Goat-anti Rabbit IgG Alexa Fluor 488 (1:1000) (Invitrogen A-11008) and Donkey anti-Goat IgG Alexa Fluor 594 (1:1000) (Invitrogen A-11058) secondary antibodies for 1 hour at room temperature. Coverslips were washed 3X with PBS and mounted with Prolong Diamond antifade mountant with DAPI (Invitrogen P36962).

IFNγ-induced IRF1 expression derived from mammalian expression

U2OS cells were grown on coverslips in cDMEM supplemented with 10% FBS and 2mM L-glutamine. For transfections, cells were maintained in cDMEM supplemented with only 2mM L-glutamine. Cells were transfected with 500ng of plasmid DNA using the Lipofectamine 3000 Transfection Reagent (Invitrogen L3000001) according to the manufacturer’s instructions. Cells were incubated at 37°C, 5% CO₂ for 17 hours. Cells were treated with IFNγ recombinant human protein (1 ng per mL of cDMEM; Gibco PHC4031) for 6 hours. Coverslips were fixed with 4% paraformaldehyde and stored in PBS at 4°C until staining was performed. Coverslips were incubated with blocking buffer (5% BSA, 0.15% Triton X-100) for 1 hour and stained with mouse Ty1 Tag monoclonal Ab (BB2) (1:100) (Invitrogen MA5-23513) and rabbit IRF1 (1:200) (Cell Signaling Technology, 8478S) primary antibodies for 1 hour at room temperature. Coverslips were washed 3X with PBS and stained with Goat-anti Rabbit IgG Alexa Fluor 647 (1:1000) (Invitrogen A-21244) and Goat anti-Mouse IGG Alexa Fluor 594 (1:1000) (Invitrogen A-11032) secondary antibodies for 1 hour at room temperature. Coverslips were washed 3X with PBS and mounted with Prolong Diamond antifade mountant with DAPI (Invitrogen P36962).

Ethics statement

Work described herein involves animal research, all of which was approved by the local IACUC committee at both the USDA and University of Pittsburgh under protocol numbers 15–017 (USDA) and 18032113 (U. Pittsburgh). All animals were euthanized according to AVMA guidelines. For felines, this involved the use of a sedative followed by lethal barbiturate injection, and for mice this involved exposure to CO₂ in a closed chamber at a flow rate between 30%-70% of the chamber volume according to AVMA guidelines.

Supporting information

S1 Fig [a]
VEG (TgVEG) and Amer (HhAmer) sporozoite infections in vivo.

S2 Fig [gsea]
Top canonical pathways enriched ( scores) in or -infected THP-1 cells compared to mock-infected cells.

S3 Fig [a]

S4 Fig [pdf]
Bystander cells lack any detectable expression of parasite transcript.

S5 Fig [tgveg]
β-galactosidase (gal) assay to detect activity of β-gal in THP-1 cells (Pellet) and supernatant.

S6 Fig [pdf]
Heatmap of log2(FPKM) RNAseq data from THP-1 cells infected with , strain VEG, or pre-infection with one species followed by infection with the other (HV→H. hammondi and then T. gondii; VH→T. gondii VEG followed by H. hammondi).

S1 Table [xlsx]
DESeq2 output for comparisons between THP-1 cells infected with different parasite strains and species and mock-treated THP-cells.

S2 Table [xlsx]
GSEA datasets found to be statistically significant for each strain and/or species comparison between infected and mock-treated THP-1 cells.

S3 Table [xlsx]
Log2 (FPKM) transcript count values across all THP-1 samples infected with different strains and species (or mock-infected) for genes belonging to either the Response or gene sets from the Hallmarks GSEA database.

S4 Table [xlsx]
Ingenuity Pathway Analysis output for THP-1 cells infected with or or subjected to mock treatment.

S5 Table [xlsx]
Signficant “upstream regulators” of downstream targets modulated by infection with T. gondii or H. hammondi compared to mock-treated THP-1 cells.

S6 Table [xlsx]
Backround-subtracted fluorescence intensity values of 30 target analytes found in supernatants from infected or mock-treated THP-1 cells using Luminex bead arrays.

S7 Table [docx]
Sequences of primers used in this study.