Ras–Erk MAPK signaling controls many of the principal pathways involved in metazoan cell motility, drives metastasis of multiple cancer types and is targeted in chemotherapy. However, its putative roles in immune cell functions or in infections have remained elusive. Here, using primary dendritic cells (DCs) in an infection model with the protozoan Toxoplasma gondii, we show that two pathways activated by infection converge on Ras–Erk MAPK signaling to promote migration of parasitized DCs. We report that signaling through the receptor tyrosine kinase Met (also known as HGF receptor) contributes to T. gondii-induced DC hypermotility. Furthermore, voltage-gated Ca2+ channel (VGCC, subtype CaV1.3) signaling impacted the migratory activation of DCs via calmodulin–calmodulin kinase II. We show that convergent VGCC signaling and Met signaling activate the GTPase Ras to drive Erk1 and Erk2 (also known as MAPK3 and MAPK1, respectively) phosphorylation and hypermotility of T. gondii-infected DCs. The data provide a molecular basis for the hypermigratory mesenchymal-to-amoeboid transition (MAT) of parasitized DCs. This emerging concept suggests that parasitized DCs acquire metastasis-like migratory properties that promote infection-related dissemination.
The mitogen-activated protein kinase (MAPK) Ras–Erk (used here to refer to Erk1 and Erk2; also known as MAPK3 and MAPK1, respectively) signaling axis plays important roles in cell migration, cell proliferation and survival, and cancer metastasis (Eblen, 2018). Ras GTPases function as molecular switches by cycling between an inactive and active state. Their function in cell motility is linked to their localization to the plasma membrane, which is mediated by farnesylation (Cox et al., 1992). At the plasma membrane, Ras is locally activated by guanine-nucleotide-exchange factors (GEFs). In turn, Ras activates the Raf–Mek–Erk signaling cascade, which regulates cell migration (Chernyavsky et al., 2005; Shi et al., 2019). When Ras is in proximity to the cell membrane, its activation can be mediated by diverse mechanisms, including receptor tyrosine kinase (RTK) signaling, integrin activation and Ca2+ influx (Giehl et al., 2000; Rosen et al., 1994; Schlaepfer et al., 1994).
In cancer cells, the RTK Met (also termed c-Met, hepatocyte growth factor receptor or HGFR) promotes cell migration and metastasis upon binding its ligand, hepatocyte growth factor (Hgf,scatter factor or SF) (Webb et al., 1998). Activated Met recruits RasGEFs, which locally activate Ras, leading to Erk phosphorylation and cell motility (Hartmann et al., 1994; Wang et al., 2002). Additionally, voltage-gated Ca2+ channels (VGCCs) mediate Ca2+ influx, which activates Ca2+-sensing proteins locally at the plasma membrane (Dolmetsch et al., 2001; Rosen et al., 1994). In the context of cell migration, Ca2+ influx activates calmodulin (CaM) and downstream kinases, which impacts cytoskeleton organization and cell motility by regulating actin dynamics through the Ras–Erk pathway (Li et al., 2004; Lundberg et al., 1998). Despite the central roles played by MAPK signaling in immune responses (Stupack et al., 2000), the impact of Ras–Erk signaling on the diverse functions of immune cells has only started to be understood (Ebert et al., 2016; Scheele et al., 2007). Furthermore, in primary dendritic cells (DCs), Ras–Erk signaling remains largely unexplored (Riegel et al., 2019).
Owing to host–pathogen co-evolution by reciprocal selection, the study of host–pathogen interactions has emerged as a powerful approach to gain insight into basic cell biology. The protozoan Toxoplasma gondii is a model obligate intracellular pathogen due to its wide host range among warm-blooded vertebrates and its ability to actively invade nucleated cells (Sibley, 2004).
One-third of the global human population is estimated to be chronically infected with T. gondii (Pappas et al., 2009). Upon ingestion and after crossing the intestinal epithelium, T. gondii tachyzoites rapidly disseminate, ultimately establishing chronic infection in the brain (Montoya and Liesenfeld, 2004). Early on, tachyzoites encounter DCs, which play a determinant role in mounting a robust protective immune response (Liu et al., 2006). Paradoxically, T. gondii exploits the inherent migratory ability of DCs for dissemination via a ‘Trojan horse’ mechanism (Courret et al., 2006; Lambert et al., 2006, 2009; Sangare et al., 2019). Within minutes of active invasion by T. gondii, DCs in mice adopt a hypermigratory phenotype that mediates rapid systemic dissemination (Kanatani et al., 2017; Weidner and Barragan, 2014). We recently reported that T. gondii induces mesenchymal-to-amoeboid transition (MAT) in DCs (Ólafsson et al., 2018). Hypermigratory parasitized DCs are characterized by high-velocity amoeboid locomotion (termed hypermotility), cytoskeletal reorganization with loss of podosomes, decreased pericellular proteolysis and redistribution of integrins (Kanatani et al., 2015; Ólafsson et al., 2018; Weidner et al., 2013). Hypermigration is triggered by non-canonical GABAA-receptor-mediated (Fuks et al., 2012) activation of VGCCs, including subtype CaV1.3 (Kanatani et al., 2017), and Timp-1-mediated, Itgb1-dependent activation of focal adhesion kinase (Fak, also known as PTK2) (Ólafsson et al., 2019). However, the signaling elements downstream of VGCC and Itgb1–Fak signaling, and their respective contribution to the hypermigratory phenotype of DCs, remain unknown.
Here, we use primary DCs and T. gondii to investigate alternative pathways for migratory activation of immune cells. We identify Ras-dependent sustained Erk phosphorylation downstream of VGCC–CaM–CaMkII signaling and Met signaling, with an impact on non-canonical migratory activation of T. gondii-infected DCs. The findings provide novel mechanistic insights into infection-related MAT of DCs.
Erk is rapidly phosphorylated in parasitized DCs, and gene silencing of Erk1 and Erk2 abolishes DC hypermotility
Because the MAPKs Erk1 and Erk2 (Erk) play a key role in cancer cell metastasis (Viala and Pouyssegur, 2004) and leukocyte motility (Stupack et al., 2000), we investigated Erk activity in primary murine DCs challenged with T. gondii tachyzoites. We found that infection affected Mapk3 (Erk1) and Mapk1 (Erk2) mRNA expression (Fig. S1A). However, the total Erk protein expression of DCs was non-significantly altered upon T. gondii challenge 6 h post-infection (h.p.i.) (Fig. S1B). In sharp contrast, elevated Erk phosphorylation was observed at this time point, which was abrogated upon treatment with the Mek inhibitor U0126 (Fig. 1A). Because DC hypermigration begins within minutes of T. gondii invasion (Weidner et al., 2013), we assessed the early kinetics of Erk phosphorylation upon T. gondii challenge. As we found that the ratio of phosphorylated Erk to total Erk was rapidly elevated in T. gondii-challenged DCs (Fig. 1B), we investigated its putative role in DC hypermotility. Importantly, inhibition of Erk phosphorylation through Mek antagonism significantly decreased the mean velocity of T. gondii-infected DCs, without significantly impacting the baseline motility of DCs (Fig. 1C; Fig. S1C). At the concentrations used, Mek antagonism did not significantly affect DC viability, infection frequency or tachyzoite replication (Fig. S1D–G).
To determine the respective roles of Erk1 and Erk2 in DC hypermotility, we transduced DCs with lentivirus encoding an eGFP reporter and shRNA targeting Erk1 (shErk1), Erk2 (shErk2) or non-expressed (shLuc) mRNAs. First, the knockdown efficiency of shErk1 and shErk2 was quantified. shErk1- and shErk2-transduced DCs exhibited significantly reduced levels of Erk1 and Erk2 mRNA (Fig. 1D) and protein (Fig. 1E), respectively, relative to mock- and shLuc-transduced DCs. Next, the motility of transduced parasitized DCs was analyzed (Fig. 1F,G). Importantly, gene silencing of either Erk1 or Erk2 significantly reduced hypermotility in T. gondii-infected DCs and had a non-significant effect on baseline motility (Fig. 1H). From these data, we conclude that challenge of DCs with T. gondii is accompanied by a rapid phosphorylation of Erk that underlies T. gondii-mediated migratory activation of DCs.
Erk phosphorylation via Ras downstream of VGCC–CaM–CaMkII-mediated Ca2+ signaling governs DC hypermotility
Based on findings in neuronal cells showing that Erk is activated downstream of VGCC signaling (Kotturi et al., 2003), and because VGCCs are implicated in T. gondii-mediated hypermotility (Kanatani et al., 2017), we hypothesized that VGCC–Ras–Raf–Mek signaling mediated Erk phosphorylation in T. gondii-infected DCs. To this end, we analyzed the phosphorylation of Erk in T. gondii-infected DCs upon treatment with the VGCC inhibitors nifedipine (targeting L-type VGCCs) and CPCPT (a selective inhibitor of VGCC subtype CaV1.3). Interestingly, a significant inhibition of Erk phosphorylation was observed upon VGCC antagonism with both nifedipine and CPCPT in T. gondii-infected DCs (Fig. 2A). VGCC inhibition had a non-significant impact on the baseline phosphorylation of Erk in unchallenged DCs (Fig. S2A). Because motility-related Ras signaling is strongly associated with localized farnesylated-Ras activity at the plasma membrane (Charest et al., 2010; Sasaki et al., 2004), we pharmacologically blocked plasma membrane Ras activity. For this, we used the Ras farnesylation inhibitor salirasib and found that it significantly inhibited Erk phosphorylation in T. gondii-infected DCs (Fig. 2B), with a non-significant effect on baseline Erk phosphorylation (Fig. S2B). At the concentrations used, non-significant effects were observed on DC viability, infection frequencies and tachyzoite replication (Fig. S2C–E). Conversely, VGCC agonism with Bay K8644 (Bay K), a structural analog of nifedipine with a positive ionotropic effect, led to elevated Erk phosphorylation, which was inhibited by salirasib treatment (Fig. 2C). Importantly, hypermotility was abrogated by Ras antagonism (Fig. 2D,E), in line with abrogation by VGCC inhibition, CaV1.3 gene silencing (Kanatani et al., 2017) and Mek inhibition (Fig. 1C; Fig. S1C), which together indicate the involvement of a VGCC–Ras–Mek signaling axis.
Next, because calmodulin (CaM) and calmodulin kinase II (CaMkII) act downstream of VGCC-mediated Ca2+ influx and impact Ras–Raf activity and Erk phosphorylation (Agell et al., 2002), we investigated the putative roles of CaM and CaMkII in T. gondii-mediated Erk phosphorylation and DC motility. Clearly, antagonism of CaM (using w7) or CaMkII (using sto-609) significantly reduced T. gondii-induced Erk phosphorylation (Fig. 2F). However, a significant reduction in Erk phosphorylation was also observed in unchallenged DCs upon CaM inhibition, but not CaMkII inhibition (Fig. S2B), indicating a role for CaM in baseline Erk phosphorylation. Non-significant effects were observed on DC viability, infection frequency and tachyzoite replication (Fig. S2C–E). Importantly, CaM and CaMkII antagonism inhibited hypermotility of Toxoplasma-infected DCs (Fig. 2G,H). In contrast, a non-significant effect on baseline motility was noted in unchallenged DCs upon CaM inhibition and CaMkII inhibition (Fig. 2G,H). Jointly, this indicates a higher relative dependency of hypermotility on CaM–CaMkII signaling compared with baseline motility, and is consistent with the higher relative dependency of hypermotility on Erk signaling compared with baseline motility (Fig. 1C,H).
Taken together, these data indicate that L-type VGCC-CaV1.3 signal transduction occurs via the CaM–CaMkII–Ras pathway in parasitized DCs, with subsequent phosphorylation of Erk and an impact on DC migration.
Hgf–Met signaling promotes DC motility via Ras activation and Erk phosphorylation
We recently reported the involvement of the non-receptor tyrosine kinase Fak in DC hypermotility (Ólafsson et al., 2019). However, receptor tyrosine kinases (RTKs) also play central roles in cell migration and metastasis (Lemmon and Schlessinger, 2010). Because aberrantly activated RTK Met mediates metastasis in a number of cancers (Benvenuti and Comoglio, 2007; Webb et al., 1998), we investigated the expression of Met and its ligand Hgf in DCs challenged with T. gondii. Kinetics analyses showed that DCs upregulated the transcription of Met and Hgf mRNAs shortly after T. gondii challenge (Fig. S3A,B), motivating further functional investigation.
First, we investigated the impact of Met activation with recombinant Hgf (rHgf) on Erk phosphorylation and DC motility. Indeed, treatment with rHgf induced Erk phosphorylation in unchallenged DCs (Fig. 3A). Further, rHgf treatment stimulated motility in both unchallenged and T. gondii-challenged DCs (Fig. 3B). Importantly, T. gondii-induced Erk phosphorylation was significantly inhibited upon treatment with the Met inhibitor su11274 (targeting submembrane Met phosphorylation) and was not rescued by rHgf treatment (Fig. 3C). Moreover, inhibition of Met phosphorylation inhibited infected DC hypermotility and non-significantly affected baseline motility (Fig. 3D,E; Fig. S3C), DC viability, infection frequency and tachyzoite replication (Fig. S3E–G). Unexpectedly, challenge with T. gondii non-significantly affected the amounts of secreted Hgf in supernatants, relative to unchallenged DCs (Fig. 3F). Because Ras acts downstream of Met in its activation of Erk (Wang et al., 2002), and Ras antagonism inhibited hypermotility (Fig. 2E) and Erk phosphorylation (Fig. 2B), we investigated the impact of salirasib treatment on rHgf-mediated Erk phosphorylation. Similar to its inhibitory effect on VGCC-mediated Erk phosphorylation, salirasib treatment significantly inhibited rHgf-mediated Erk phosphorylation (Fig. 3G). Further, rHgf-mediated DC motility, but not baseline motility, was abolished upon treatment with salirasib, and rHgf treatment did not rescue hypermotility in salirasib-treated infected DCs (Fig. 3H). This indicates that Hgf–Met signaling impacts hypermotility via Ras.
As we did not observe elevated Hgf secretion upon T. gondii challenge and because T. gondii activates integrin Itgb1–Fak signaling in DCs (Ólafsson et al., 2019), we hypothesized that, in addition to Hgf, Hgf-independent activation of Met might occur via Fak (Hui et al., 2009). Indeed, Erk phosphorylation was significantly reduced in T. gondii-infected DCs upon treatment with inhibitors targeting Fak and the associated kinase Pyk2 (Fig. 3I). The treatments had a non-significant effect on Erk phosphorylation in unchallenged DCs (Fig. S3D), DC viability, infection frequency and tachyzoite replication (Fig. S3E–G).
Taken together, the data show that Met–Ras signaling is activated in DCs upon T. gondii infection, with an impact on motility via phosphorylation of Erk. Further, our data suggest that, in parasitized DCs, Met is activated both by its ligand Hgf and via tyrosine kinase-mediated transactivation.
Gene silencing of Met inhibits hypermotility of T. gondii-infected DCs
To determine the role of Met in T. gondii-induced hypermigration, we applied a gene silencing approach in primary DCs. First, we analyzed Met protein expression in DCs by flow cytometry (Fig. S4A,B). Consistent with Met transcription data, signal corresponding to Met protein was significantly elevated in CD11c+ DCs infected by T. gondii (Fig. 4A). Of note, parasitized DCs exhibited significantly higher signal corresponding to plasma membrane-associated Met and total Met compared with the signal exhibited by non-infected bystander DCs (Fig. 4A). This indicated a selective upregulation of Met in parasite-invaded DCs. Next, we silenced Met mRNA expression with a lentivirus encoding an eGFP reporter and shRNA targeting Met mRNA (shMet) or non-target control mRNA (shLuc). We confirmed that DCs transduced with shMet exhibited significantly reduced Met mRNA (Fig. 4B, Fig. S4C) and Met total protein (Fig. 4C) relative to shLuc transduced DCs. We then subjected Met-silenced T. gondii-challenged DCs to motility analyses (Fig. S4D). Importantly, gene silencing of Met abolished hypermotility in T. gondii-infected DCs (Fig. 4D,E), consistent with the effect of pharmacological antagonism of Met (Fig. 3E). Taken together, these data show that Met signaling is activated in T. gondii-infected DCs with an impact on motility.
In this study we investigated the putative role of the MAPKs Erk1 and Erk2 (Erk) in T. gondii-induced hypermigration of primary DCs. We report that T. gondii-induced VGCC and Met signaling converge on the Ras–Erk signaling axis to mediate hypermotility of parasitized DCs.
Our studies establish that Erk signaling plays a central role in Toxoplasma-induced hypermigration of DCs. A rapid and sustained Erk phosphorylation was observed upon T. gondii challenge, leading to the activation of DC motility. Despite transcriptional changes following infection, total Erk protein amounts remained stable, likely due to the 50 h half-life time of Erk (Schwanhausser et al., 2011). Using shRNA-mediated gene silencing of Mapk3 and Mapk1 and pharmacological antagonism of Erk phosphorylation (by Mek inhibition), we demonstrate that phosphorylated Erk is necessary for DC hypermotility. Interestingly, knockdown of either Erk1 or Erk2 was sufficient for reducing hypermotility (although baseline motility was maintained), despite shared substrate redundancy between Erk1 and Erk2 (von Kriegsheim et al., 2009). This indicates a dependency on both Erk isoforms for hypermotility but not for baseline DC motility. Moreover, the observed compensatory elevation of Erk1 expression upon silencing of Erk2, and elevation of Erk2 upon silencing of Erk1, were not sufficient to maintain hypermotility. This rapid reciprocal compensatory expression between Erk1 and Erk2 advocates for a tight regulation of the Erk MAPK pathway in primary DCs. Furthermore, the crucial dependence of DC hypermotility on Erk signaling cannot be generalized to other MAPK pathways as, for example, infected DCs remain hypermotile upon inhibition of the MAPK p38 (ten Hoeve et al., 2019). Taken together, these observations prompted us to investigate signaling pathways upstream of Erk activation in parasitized DCs.
Our data show that VGCC–CaM–CaMkII–Ras signaling acts upstream of Erk to mediate migratory activation of DCs. Previously, we showed that the onset of the hypermigratory phenotype in DCs depends on live intracellular parasites and the discharge of T. gondii secretory organelles, independently of TLR–MyD88 signaling or chemotaxis (Fuks et al., 2012; Lambert et al., 2006; Ólafsson et al., 2018; Sangare et al., 2019; Weidner et al., 2013). However, TLR–MyD88 signaling and chemotactic stimuli can also activate Erk. Specifically, in T. gondii-infected macrophages both TLR–MyD88-dependent and a strong component of TLR–MyD88-independent Erk activation have been described (Kim et al., 2006). However, T. gondii-induced hypermigration is fully maintained in MyD88-deficient DCs (Lambert et al., 2006; Ólafsson et al., 2018). Similarly, hypermigration is independent of chemotactic stimuli (but can act synergistically with chemotaxis upon stimulation) (Fuks et al., 2012; Kanatani et al., 2015; Weidner et al., 2013). Thus, alternative signaling underlies the migratory activation of DCs by T. gondii and, consequently, the hypermotility-related activation of Erk observed here.
T. gondii promotes DC hypermigration via GABAergic signaling (Fuks et al., 2012), which triggers the activation of the VGCC CaV1.3 (Kanatani et al., 2017). VGCC-mediated Ca2+ influx impacts Ca2+-sensing proteins locally at the cell membrane (Rosen et al., 1994; Zhou et al., 2015). Here, by pharmacological agonism and antagonism of the VGCC signaling axis, we extend these findings by linking Erk phosphorylation to VGCC activation and CaM–CaMkII–Ras signaling. We show that inhibition of L-type VGCCs and CaV1.3 with nifedipine and CPCPT, respectively, effectively blocked Erk phosphorylation in T. gondii-infected DCs. We previously showed that VGCC agonism enhances migration of unchallenged and T. gondii-challenged DCs (Kanatani et al., 2017). Importantly, reinforcing the link between VGCCs, and specifically CaV1.3, and Erk, we show that L-type VGCC agonism (Bay K) enhanced Erk phosphorylation via CaM–CaMkII–Ras signaling whereas the opposite effect was observed upon antagonism (nifedipine, CPCPT). These data are in line with paradigms in neuronal cells, where VGCC-dependent CaM activity regulates cytoskeleton organization and cell migration via Ras–Erk signaling (Rosen et al., 1994). Further, in lymphocytes, treatment with nifedipine and Bay K blocks and stimulates Erk phosphorylation, respectively (Kotturi et al., 2003). Thus, our data describe, for the first time, VGCC (CaV1.3)-mediated Erk activation in leukocytes and provide evidence for its dependence on CaM–CaMkII–Ras signaling.
Cancer cells seldom rely on one single pathway to orchestrate metastasis. Instead, metastasis generally relies on the activation of redundant and complementary pathways (Tracey et al., 2013). Consequently, we reasoned that diverse signaling cascades might underlie hypermotility in parasitized DCs.
Here, we identify a role for the receptor tyrosine kinase Met in the migratory activation of primary DCs. Upon T. gondii challenge, transcriptional upregulation and increased Met protein expression was indicative of activation. Importantly, antagonism of Met and gene silencing of Met inhibited DC hypermotility. Functionally reinforcing the observation that Met is upregulated on the plasma membrane of parasitized DCs, rHgf treatment and infection had a synergistic effect on migratory activation. Furthermore, treatment with rHgf induced Erk phosphorylation and motility in unchallenged DCs. Conversely, Met antagonism inhibited Erk phosphorylation in unchallenged rHgf-treated DCs and T. gondii-infected DCs. Finally, Ras inhibition blocked rHgf–Met-induced Erk phosphorylation. Consistent with this idea, aberrant Met signaling mediates invasive cell growth and metastasis (Benvenuti and Comoglio, 2007) and afferent lymphatic migration of epidermal DCs upon skin inflammation (Baek et al., 2012). Altogether, our data demonstrate that Met mediates migratory activation of DCs through Ras–Erk signaling.
Because Hgf is the only known ligand for Met, we investigated the secretion of Hgf by DCs upon T. gondii challenge. Surprisingly, the concentration of Hgf in supernatant from DCs was non-significantly affected by T. gondii challenge, suggesting that alternative mechanisms activate Met signaling in T. gondii-infected DCs. Ligand-independent activation of Met, or transactivation, is regulated by adapter proteins, integrins and tyrosine kinase crosstalk (Fischer et al., 2004; Hui et al., 2009; Mitra et al., 2011). Pharmacological antagonism of Fak and Pyk2 blocked Erk phosphorylation to a similar extent to Met antagonism in T. gondii-infected DCs, providing evidence for tyrosine kinase-mediated transactivation of Met. In breast cancer cells, the integrin Itgb1 transactivates Met (Barrow-McGee et al., 2016). We recently described a Timp-1–CD63–Itgb1–Fak signaling axis in T. gondii-infected DCs, and found gene silencing and antagonism of Itgb1 and Fak abolished hypermotility (Ólafsson et al., 2019). Furthermore, Timp-1 was recently shown to promote activation of the RTK c-Kit in DLD-1 colorectal cancer cells (Nordgaard et al., 2019). Taken together, the data advocate for a potentiation of the motogenic action of Met through the Timp-1–CD63–Itgb1–Fak axis. However, additional possibilities that are not mutually exclusive with this idea exist. The secreted T. gondii proteins MIC1, MIC3 and MIC6 can directly phosphorylate the RTK Egfr (Muniz-Feliciano et al., 2013), which could hypothetically contribute to Met transactivation. Collectively, the data demonstrate that T. gondii activates the Met signaling pathway in infected DCs. Consistent with results from our previous studies, a milieu of RTK crosstalk with Erk phosphorylation in T. gondii-infected immune cells emerges, reminiscent of that described in metastatic cancer cells (Kaufmann et al., 2009; Volinsky and Kholodenko, 2013).
The data raise questions on how the regulation of hypermigration is fine tuned. Amoeboid hypermigration of parasitized DCs represents a dramatic activation of DC locomotion, which is sustained over time and integrates multiple cell signaling pathways (Fuks et al., 2012; Kanatani et al., 2015; Ólafsson et al., 2018; Weidner et al., 2013). Notably, blockade of VGCC–Ras signaling or Met–Ras signaling independently leads to inhibition of hypermotility, but not baseline motility of DCs. We speculate that the onset and maintenance of hypermigration requires a threshold signaling activity that is achieved by co-engaging tightly regulated VGCC and Met RTK signaling. Supporting this concept, in neuronal models, VGCC and NGF RTK signaling synergistically converge on Ras for neuronal survival (Vaillant et al., 1999). Additionally, cross-regulation between VGCC and RTK signaling may take place (Cullen and Lockyer, 2002; Vela et al., 2007).
For the experimental conditions used, the pharmacological inhibitors had a non-significant effect on DC viability and parasite replication. However, an impact on the secretion of putative parasite-derived effectors by pharmacological inhibitors, or subtle effects on parasite metabolism without an impact on viability, cannot be ruled out. However, the inhibitor data is consistent with gene silencing data targeting effectors of the host cell. The data are also in line with previous work targeting upstream components of the two converging signaling pathways by gene silencing and with pharmacological modulators (Fuks et al., 2012; Kanatani et al., 2017; Ólafsson et al., 2019).
Based on our data, we propose a model for the migratory activation of DCs upon challenge with T. gondii (Fig. 5). T. gondii infection activates Ras signaling through Met and GABAAR–VGCC–CaM–CaMkII signaling. Subsequently, the Ras–Raf–Mek cascade phosphorylates Erk, which drives migratory activation in DCs. The data show that T. gondii orchestrates the migratory activation of parasitized DCs through the Ras–Erk signaling node, which in turn is activated via tyrosine kinase signaling, integrins and GABA/Ca2+ ion channel activation (Kanatani et al., 2017; Ólafsson et al., 2019). Our data also provide a molecular basis for the MAT and hypermigration of parasitized DCs. Interestingly, amoeboid motility is a feature of metastasizing cells, and the signaling in parasitized DCs is consistent with emerging paradigms of signaling in metastatic cells (Lambert et al., 2017). Future studies will unravel the downstream effectors of Erk1 and Erk2 in DC hypermotility. In this context, putative parasite-derived effectors have recently been identified, which may interact with host cell MAPK signaling (Weidner et al., 2016), WAVE complex and actin dynamics (Sangare et al., 2019), and Rho GTPases (Drewry et al., 2019) in infected leukocytes. Thus, our data provide a signaling framework for assessing parasite-derived effectors and non-canonical migratory activation of leukocytes. Future investigations need to address how putative effectors emanating from rhoptries and dense-granule organelles impact the onset and regulation of the hypermigratory phenotype of parasitized phagocytes. Finally, this work highlights conserved signaling mechanisms between neurons, metastasizing cells and immune cells in the context of infection, which should be taken into consideration when designing therapies that target Met, VGCCs or Ras–Erk signaling.
MATERIALS AND METHODS
The Regional Animal Research Ethical Board, Stockholm, Sweden, approved experimental procedures and protocols involving extraction of cells from mice, following proceedings described in EU legislation (Council Directive 2010/63/EU).
Cells and parasites
Murine bone marrow-derived DCs (DCs) were generated as previously described (Lambert et al., 2006). Briefly, cells from bone marrow of 6–10-week-old male or female C57BL/6 mice (Charles River) were cultivated in RPMI 1640 with 10% fetal bovine serum (FBS), gentamicin (20 μg/ml), glutamine (2 mM) and HEPES (0.01 M) – referred to as complete medium (CM; all reagents from Life Technologies) – and supplemented with recombinant mouse GM-CSF (10 ng/ml; Peprotech). Loosely adherent cells were harvested after 6–10 days of maturation. T. gondii tachyzoites of the RFP-expressing Prugniaud strain (Pru and Pru-RFP, type II) (Lambert et al., 2009) or GFP-expressing Ptg strain (type II) (Weidner et al., 2013) lines were maintained by serial 2-day passages in human foreskin fibroblast (HFF-1 SCRC-1041, American Type Culture Collection) monolayers. Cell and parasite cultures were periodically tested for mycoplasma (Biontex). The hypermigratory phenotype induced by type II strains in DCs has been previously characterized (Ólafsson et al., 2018).
The following soluble reagents were used to treat DCs in motility assays and for samples analyzed by western blotting: w-7 hydrochloride (w7, 25 μM, Tocris), sto-609 acetate (sto., 25–50 μM, Tocris), rHgf (40 ng/ml, Sigma-Aldrich), su11274 (su., 10 μM, Tocris), U0126 (10 μM, Tocris), salirasib (sal., 25–100 μM, Tocris), nifedipine (nif., 50 μM, Sigma-Aldrich), 1-(3-Chlorophenethyl)- 3-cyclopentylpyrimidine-2,4,6-(1H,3H,5H)-trione (cpcpt, 10 μM, Sigma-Aldrich), bay K8644 (Bay K, 10 μM, Sigma-Aldrich), pf573228 (pf228, 10μM, Tocris), pf562271 (pf5, 10 μM, Selleckchem).
Lentiviral vector production and in vitro transduction
Self-complementary hairpin DNA oligonucleotides targeting mouse Met (shMet), Erk1 (shErk1) Erk2 (shErk2) mRNA, and a non-related sequence shRNA control (shLuc) were chemically synthesized (DNA Technology, Denmark), aligned and ligated in a self-inactivating lentiviral vector (pLL3.7) containing a CMV-driven eGFP reporter and a U6 promoter upstream of cloning restriction sites (HpaI and XhoI) (Table S1). Lentivirus production was performed using Lipofectamine transfection. Briefly, shLuc, shMet, shErk1 or shErk2 vectors were co-transfected with the packaging vector psPAX2 (12260, Addgene) and envelope vector pCMV-VSVg (8454, Addgene) into HEK293FT or HEK293T Lenti-X cells (Takara Bio) and the resulting supernatant was collected after 48–72 h. The supernatant was centrifuged at 3000 g for 15 min to eliminate cell debris, filtered through 0.45 mm cellulose acetate filters, and virus particles were concentrated by ultracentrifugation at 50,000 g for 1 h, then resuspended in RPMI. Titers were determined by infecting HEK293FT cells with serial dilutions of concentrated lentivirus and determination of percentage of infected cells by flow cytometry (FACS Fortessa, BD Biosciences). Bone marrow-derived DCs were transduced twice at a multiplicity of infection (MOI) of 1 on day 3 and 4, or once at MOI 2 on day 3 by spinoculation at 1200 g for 2 h. Three days post-transduction eGFP-expression was verified by epifluorescence microscopy before the cells were used in experiments.
Motility assays were performed as previously described (Ólafsson et al., 2019). Briefly, DCs were cultured in 96-well plates with CM±freshly egressed T. gondii tachyzoites (Pru-RFP for shRNA experiments and Ptg-GFP for all other experiments, MOI 3, 4 h) and soluble reagents as indicated. Bovine collagen I (1 mg/ml, Life Technologies) was then added and live cell imaging was performed for 1 h, 1 frame/min, using a 10× objective (Z1 Observer with Zen 2 Blue v. 4.0.3, Zeiss). Time-lapse images were consolidated into stacks and motility data was obtained from >30 cells/condition (Manual Tracking, ImageJ) yielding mean velocities (Chemotaxis and migration tool, v. 2.0, Ibidi). Infected DCs were defined by localization of tachyzoite-derived RFP or GFP fluorescence within the cell boundary. Transduced cells were defined by the expression of eGFP.
DCs were cultured in CM with or without freshly egressed T. gondii tachyzoites (Ptg-GFP, MOI 3). Total RNA was extracted using TRIzol reagent (Invitrogen). Quantitative PCR was performed using SYBR Green PCR master mix (Kapa Biosystems), forward and reverse primers (200 nM) and cDNA (100 ng) and run on the Rotor Gene 6000 system (Corbett) for 45 cycles. Amplicons were validated with melting curves and agarose gel (1%) analysis. Data was analyzed with RG-6000 application software (v1.7, Corbett). Gapdh and Actb were used as housekeeping genes to generate ΔCt values. 2(−ΔCt) values were generated to show relative knockdown efficiency and 2(−ΔΔCt) values were used to show expression fold-change upon infection. Fold changes in expression were calculated using the comparative DCT method against the non-infected DC control. Primers were designed using Getprime (David et al., 2017) (Table S2) and purchased from Invitrogen.
Flow cytometry and viability assays
DCs were cultured in CM with or without freshly egressed T. gondii tachyzoites (Ptg-GFP, MOI 3, 4 h). Following Fc blockade (clone 2.4G2, Cat. #553141, BD Pharmingen), cells were stained with anti-CD11c-PE-Cy7 (clone N418, Cat. #25-0114-82, eBiosciences). After fixation (PFA 4%), cells were either directly stained (to detect plasma membrane Met) or permeabilized (IntraPrep Permeabilization kit, Cat. #IM2389, Beckman Coulter) before staining (to detect total Met). For staining we used anti-Met-PE (clone eBioclone 7, Cat. #12-8854-82, eBiosciences) or isotype control (clone eBRG1, Cat. #14-4301-82, eBiosciences). All antibodies were used at a dilution of 1:100. Samples were run on a BD LSRFortessa Cell Analyzer (BD Biosciences). Data was analyzed in FlowJo (Tree Star Inc, OR).
For infection frequencies, DCs were cultured in CM±reagents (as indicated) and freshly egressed T. gondii tachyzoites (Ptg-GFP, MOI 3) for 2 h. After fixation (PFA 4%), samples were run on an easyCyteTM 8HT (Millipore), in accordance with the manufacturer's guidelines. Samples were gated on FSC, SSC and GFP. A minimum of 30,000 cells were analyzed per condition. Data were analyzed in FlowJo (Tree Star Inc, OR). Fluorescence interference of su11274 precluded determination of infection frequency with this reagent. DC viability was assessed by flow cytometry (FACS Fortessa, BD) using LIVE/DEAD Fixable Violet staining (Cat. #L34963, Thermo Fisher Scientific) in the BV-421 channel. Sto-609 exhibited fluorescence in the BV-421 channel and was therefore assessed with adapted gating. Parasite replication was assessed on HFF monolayers after 24 h by counting 100 vacuoles/condition using epifluorescence microscopy (Z1 Observer with Zen 2 Blue v. 4.0.3, Zeiss) as detailed in Ólafsson et al. (2019).
Supernatants from DCs cultured in CM with or without freshly egressed T. gondii tachyzoites (Ptg-GFP, MOI 3) were collected at 2, 6, 12 or 24 h post-infection. Samples were analyzed by ELISA (MHG00, Thermo Fisher Scientific) according to the manufacturer's instructions. Samples were normalized to a CM control.
Cell lysates were collected and sonicated in RIPA buffer (150 mM NaCl, 5 mM EDTA, pH 8.0, 50 mM Tris-HCl, pH 8.0, 0.1% Triton-X100, 0.5% sodium deoxycholate and 0.1% SDS) with protease and phosphatase inhibitors (Thermo Fisher Scientific). Proteins were separated on 10% SDS-PAGE gels followed by transfer to PVDF Immobilon membranes (Millipore). Phospho-Erk was detected using an anti-phospho-Erk antibody (1:1000, clone D13.14.4E, Cat. #4370, Cell Signaling Technology), total Erk was detected using an anti-Erk antibody (1:2000, clone 137F5, Cat. #4695, Cell Signaling Technology) and Gapdh was detected with an anti-Gapdh antibody (1:3000, Cat. #ABS16, Millipore). HRP-conjugated anti-rabbit-IgG (1:3000, Cat. #7074S, Cell Signaling Technology) was used as secondary antibody. Proteins were visualized using ECL reagents (GE healthcare) in a Biorad ChemiDoc XRS+. Densitometry analysis was performed using ImageJ (NIH, MD).
Multiple comparisons of normally distributed data were carried out with one-way ANOVA followed by Tukey's HSD, Holm–Sidak's or Dunnett's multiple comparisons test. Two-way ANOVA followed by Holm–Sidak's multiple comparisons test was used to evaluate the interaction between time and infection. Two sample comparisons were carried out with paired or unpaired t-tests. Normality was tested by the Shapiro–Wilks test. In all statistical tests P-values<0.05 were defined as significant. All statistics were performed with Prism (v. 8, Graphpad).
We thank all members of the Barragan laboratory for critical input.
Conceptualization: E.B.Ó., A.B.; Methodology: E.B.Ó., A.L.t.H., X.L.-W., L.W., M.V.-G.; Validation: M.V.-G., A.B.; Formal analysis: E.B.Ó., A.L.t.H.; Investigation: E.B.Ó., A.L.t.H., X.L.-W., L.W.; Data curation: E.B.Ó.; Writing - original draft: E.B.Ó., A.B.; Writing - review & editing: E.B.Ó., A.B.; Visualization: E.B.Ó.; Supervision: A.B.; Project administration: A.B.; Funding acquisition: A.B.
This work was funded by the Swedish Research Council (Vetenskapsrådet, 2018-02411 to A.B.) and the Olle Engkvist Foundation (Stiftelsen Olle Engkvist Byggmästare, 193-609 to A.B.).
Peer review history
The peer review history is available online at https://jcs.biologists.org/lookup/doi/10.1242/jcs.241752.reviewer-comments.pdf
The authors declare no competing or financial interests.