A motogenic GABAergic system of mononuclear phagocytes facilitates dissemination of coccidian parasites.

Gamma-aminobutyric acid (GABA) serves diverse biological functions in prokaryotes and eukaryotes, including neurotransmission in vertebrates. Yet, the role of GABA in the immune system has remained elusive. Here, a comprehensive characterization of human and murine myeloid mononuclear phagocytes revealed the presence of a conserved and tightly regulated GABAergic machinery with expression of GABA metabolic enzymes and transporters, GABA-A receptors and regulators, and voltage-dependent calcium channels. Infection challenge with the common coccidian parasites Toxoplasma gondii and Neospora caninum activated GABAergic signaling in phagocytes. Using gene silencing and pharmacological modulators in vitro and in vivo in mice, we identify the functional determinants of GABAergic signaling in parasitized phagocytes and demonstrate a link to calcium responses and migratory activation. The findings reveal a regulatory role for a GABAergic signaling machinery in the host-pathogen interplay between phagocytes and invasive coccidian parasites. The co-option of GABA underlies colonization of the host by a Trojan horse mechanism.

Introduction

Gamma-aminobutyric acid (GABA), first identified as a plant and microbe metabolite, is a principal neurotransmitter in the central nervous system (CNS) of vertebrates (Roth et al., 2003). Moreover, recent findings implicate GABAergic signaling in the disease environment of cancer and other inflammatory conditions in humans (Neman et al., 2014; Bhat et al., 2010; Takehara et al., 2007; Li et al., 2017). Neurons and other GABAergic cells synthesize GABA via glutamate decarboxylases (GAD65/67) (Soghomonian and Martin, 1998). GABA is shuttled in and out of cells via GABA transporters (GATs) (Höglund et al., 2005) and acts via activation of GABA-A receptors (GABA-A Rs) (Olsen and Sieghart, 2008) and GABA-B Rs (Bettler et al., 2004). The GABA-A Rs are pentameric ionotropic chloride channels, normally comprised of three types of subunits: 2 αs, 2 βs and a third type of subunit. Nineteen different mammalian GABA-A R subunits (α1–6, β1–3, γ1–3, δ, ε, π, θ and ρ1–3) can combine to form numerous variants of functional heteromeric receptors in neuronal cells. The strength and polarity of GABA signaling is regulated by cation-chloride cotransporters (CCCs) (Kaila et al., 2014). GABA-A R activation by GABA can elicit opening of voltage-dependent calcium (Ca2+) channels (VDCCs) with subsequent Ca2+ influx into the neuronal cell (Bortone and Polleux, 2009).

Owing to host-pathogen coevolution with reciprocal selection, studies of host-pathogen interactions provide a powerful tool to gain insight into biological processes. The obligate intracellular protozoan Toxoplasma gondii actively invades nucleated cells (Sibley, 2004) and has a broad range of hosts among warm-blooded vertebrates. One third of the global human population is estimated to be chronically infected by T. gondii (Pappas et al., 2009) and severe manifestations may occur upon immunosuppression and during pregnancy (Montoya and Liesenfeld, 2004). Similarly, the related coccidian Neospora caninum represents a pathogen of major importance in veterinary medicine (Dubey et al., 2007). Upon ingestion and after crossing the intestinal epithelium, the tachyzoite stages of these coccidian parasites rapidly disseminate in their intermediate hosts, ultimately establishing latent infection in the CNS (Montoya and Liesenfeld, 2004; Dubey et al., 2007).

The mononuclear phagocyte system comprises dendritic cells (DCs), monocytes, macrophages and brain microglia, which mediate multiple immunological functions and are crucial to counteract microbial infection (Guilliams et al., 2014). Early on during infection, tissue-invasive coccidian tachyzoites encounter DCs and other phagocytes, which play a determinant role in mounting a robust host-protective immune response (Liu et al., 2006; Mashayekhi et al., 2011). Paradoxically, T. gondii and N. caninum exploit the inherent migratory ability of DCs and monocytes for dissemination via a Trojan horse mechanism (Sangaré et al., 2019; Lambert et al., 2006; Lambert et al., 2009; Courret et al., 2006; Collantes-Fernandez et al., 2012). Within minutes of active invasion by T. gondii, DCs adopt a hypermigratory phenotype that mediates rapid systemic dissemination in mice (Kanatani et al., 2017; Weidner and Barragan, 2014). GABAergic inhibition of DCs hampers dissemination of T. gondii (Kanatani et al., 2017; Fuks et al., 2012), however the precise mechanisms of action have remained uncharacterized.

Some components of GABAergic signaling have been detected in DCs, monocytes, macrophages, T cells and B cells (Bhat et al., 2010; Fuks et al., 2012; Alam et al., 2006; Wheeler et al., 2011; Bhandage et al., 2014). Yet, the precise functions of GABA in immune cells have remained elusive. Here, we have performed a systematic analysis of the GABAergic system of various types of human and murine DCs and monocytes and report a conserved GABAergic machinery implicated in migratory responses. Further, we show that coccidian parasites hijack GABAergic signaling in parasitized phagocytes to promote infection-related dissemination.

Results

Human and murine mononuclear phagocytes exhibit hypermotility and secrete GABA upon challenge with T. gondii and N. caninum

To address the impact of coccidian infection on mononuclear phagocyte motility, primary cells from human donors and mice were challenged with freshly egressed tachyzoites of T. gondii and N. caninum. Upon invasion by tachyzoites (Figure 1A), a rapid increase of migrated distances (Figure 1B) and elevated velocities (Figure 1C) were recorded in infected murine bone marrow-derived DCs (mBMDCs), compared with unchallenged cells. Similarly, human monocytes, monocyte-derived DCs (hMoDCs) and primary myeloid DCs (hMDCs) freshly isolated from blood of human donors consistently exhibited hypermotility upon challenge (Figure 1D,E,F, Figure 1—figure supplement 1A,B,C). Further, phagocytes expressed transcripts of enzymes for GABA synthesis and catabolism (Figure 1G), which were up- and downregulated, respectively, upon challenge with T. gondii (Figure 1H). Consistent with this, elevations of GABA concentrations in the supernatants were detected shortly after challenge and increased over time (Figure 1I). Further, challenge of mononuclear phagocytes with separate strains of the two coccidia resulted in elevated concentrations of GABA in cell supernatants (Figure 1J). Together, this indicated a putative link between GABA and migratory activation, motivating a further analysis of GABAergic signaling in mononuclear phagocytes.

Figure 1.

Migratory activation and GABAergic responses of mononuclear phagocytes challenged with T. gondii and N. caninum.

(A) Representative micrographs of mBMDCs infected by T. gondii tachyzoites (PRU-RFP, arrowheads) and by-stander non-infected BMDCs, stained for F-actin (Alexa Fluor 488 Phalloidin) and nuclei (DAPI). Scale bars, 20 μm, inset image 10 μm. (B) Representative motility plots of unchallenged mBMDCs (green) and T. gondii- or N. caninum-infected mBMDCs (red and orange, respectively). X- and y-axes show distances in μm. (C) Box-and-whisker dot plots show, for each condition in (B), median velocities (μm/min) of cells shown in motility plots (n = 3–6 independent experiments). (D) Motility plots of hMonocytes, hMoDCs and hMDCs, respectively, unchallenged (blue) or infected with T. gondii (PRU-RFP, red). (E) Quantifications of velocities (μm/min) in (D) are shown as box-and-whisker dot plots (n = 3–6 independent experiments). (F) Normalized mean velocities of unchallenged and T. gondii-infected hMDCs from human donors (n = 5 independent donors, 50–60 cells per donor). (G) Relative mRNA expression (2-ΔCt) of GABA synthesis enzymes (GAD65 and GAD67) and catabolic enzyme (GABA-T) in unchallenged mBMDCs, hMoDCs, hMDCs and hMonocytes, respectively, determined by real-time qPCR. (H) Corresponding heat maps for the indicated cell types show (%) transcriptional changes upon challenge with T. gondii relative to unchallenged cells at indicated time-points, as detailed in Materials and methods. (X) indicates no amplification (n = 3–6 independent experiments). (I) GABA (ng/ml) secreted in supernatants of hMoDCs challenged with T. gondii (ME49/PTG) at indicated time-points (n = 3 human donors). (J) GABA (ng/ml) secreted in supernatants of mBMDCs, hMonocytes and hMoDCs, respectively, challenged with different strains of T. gondii (PRU and RH) or N. caninum (Nc-1 and Nc-Liverpool) was quantified by ELISA (n = 6–8 independent experiments). Bar graphs show mean + SEM. Statistical significance was tested by Mann-Whitney test for (C, E, F), ordinary one-way ANOVA with Dunnett’s multiple comparison test for (J) and paired t-test for (I), *p<0.05, **p<0.01, ***p<0.001, ns p≥0.05.

(A) Representative bivariate dot plots show characterization of hMDC preparations using flow cytometry. Cells were checked for contaminating T cells (CD3), B cells (CD20) and monocytes (CD14) and confirmed to be HLA-DR+ and CD1c+. (B) Representative motility plots of unchallenged and T. gondii-infected hMDCs from five independent human donors (1-5). X- and y-axes indicate distances in μm. (C) Box-and-whisker dot plots show, for each indicated condition, velocities (μm/min) of unchallenged and T. gondii-infected hMDCs, as in (B). (D) Representative motility plots of unchallenged and T. gondii-infected hMDCs treated with picrotoxin and TPMPA (inhibitors of GABA-A Rs), or bumetanide (NKCC1 inhibitor), or benidipine and nifedipine (inhibitors of VDCCs). (E) Dot plots show, for each indicated condition, median velocities (μm/min) of unchallenged and T. gondii-infected hMDCs, as in (D). Each dot indicates median velocity of 50–60 cells tracked from one individual donor (n = 3). Statistical significance was assessed by Mann-Whitney test for (C) and by ordinary one-way ANOVA with Tukey’s multiple comparison test for (E), **p<0.01, ***p<0.001.

Figure 1—figure supplement 1.

hMDCs exhibit a hypermotile phenotype upon T. gondii challenge.

(A) Representative bivariate dot plots show characterization of hMDC preparations using flow cytometry. Cells were checked for contaminating T cells (CD3), B cells (CD20) and monocytes (CD14) and confirmed to be HLA-DR+ and CD1c+. (B) Representative motility plots of unchallenged and T. gondii-infected hMDCs from five independent human donors (1-5). X- and y-axes indicate distances in μm. (C) Box-and-whisker dot plots show, for each indicated condition, velocities (μm/min) of unchallenged and T. gondii-infected hMDCs, as in (B). (D) Representative motility plots of unchallenged and T. gondii-infected hMDCs treated with picrotoxin and TPMPA (inhibitors of GABA-A Rs), or bumetanide (NKCC1 inhibitor), or benidipine and nifedipine (inhibitors of VDCCs). (E) Dot plots show, for each indicated condition, median velocities (μm/min) of unchallenged and T. gondii-infected hMDCs, as in (D). Each dot indicates median velocity of 50–60 cells tracked from one individual donor (n = 3). Statistical significance was assessed by Mann-Whitney test for (C) and by ordinary one-way ANOVA with Tukey’s multiple comparison test for (E), **p<0.01, ***p<0.001.

A conserved repertoire of GABA-A R subunits in human and murine phagocytes

To address GABAergic signaling in phagocytes, human and murine cells were screened for expression of the 19 known GABA-A R subunits (Supplementary file 1), which in neuronal cells combine to form multiple pentameric receptor variants (Olsen and Sieghart, 2008). Interestingly, mBMDCs consistently expressed mRNAs for 10 out of the 19 GABA-A R subunits (α3, α4, α5, β2, β3, γ1, γ2, δ, ρ1, ρ2) (Figure 2A, Supplementary file 2) and expression of GABA-A R subunits was conserved in a murine DC line (JAWSII) (Figure 2—figure supplement 1A). Further, hMoDCs from healthy donors consistently expressed the α4, ρ1, ρ2 and ρ3 subunits, while expression of the α3, β1 and β3 subunits was only detected in some donors (Figure 2B, Supplementary file 2). Finally, freshly isolated hMDCs from human donors consistently expressed β2, θ, ρ1 and ρ2 subunits of GABA-A Rs (Supplementary file 2). Interestingly, challenge with T. gondii modulated the subunit mRNA expression in murine and human DCs (Figure 2C,D, Figure 2—figure supplement 1B). Of note, transcriptional expression of the α6, β2 and θ subunits, undetectable in unchallenged hMoDCs, was consistently observed in challenged hMoDCs (Figure 2B,D). Immunocytochemical analyses with available antibodies yielded signal consistent with expression of α3, α5, β3 and ρ1 subunits (Figure 2E). We conclude that murine and human phagocytes constitutively transcribed one or more α subunits, one or more β subunits and one or more additional subunits, including ρ GABA-A R subunits (Supplementary file 2). Additionally, the modulated subunit expression upon challenge with T. gondii motivated an assessment of GABA-A R function upon infection.

Figure 2.

Modulated expression of GABA-A R subunits in DCs upon T. gondii infection.

(A, B) Relative mRNA expression (Mean + SEM, 2-ΔCt) of GABA-A R subunits in unchallenged (A) mBMDCs (n = 7 independent experiments) and (B) hMoDCs (n = 3 independent experiments). (C, D) Heat maps show (%) transcriptional changes in the expression of GABA-A R subunits upon challenge of (C) mBMDCs and (D) hMoDCs with T. gondii (PRU-RFP) relative to unchallenged cells at indicated time-point (n = 3–4 independent experiments). (E) Representative micrographs of unchallenged mBMDCs stained with antibodies against GABA-A R α3, α5, β3 and ρ1 subunit (and Alexa Flour 488-conjugated secondary antibodies), respectively, and Alexa Fluor 647 Phalloidin (F-actin) and DAPI (nuclei). Scale bars, 10 μm, for β3 5 μm (n = 3 independent experiments).

(A) Relative mRNA expression (mean + SEM, 2-ΔCt) of GABA-A receptor subunits in unchallenged JAWSII, a mouse DC line, (n = 3 independent experiments). (B) Heat map shows (%) transcriptional changes in the expression of GABA-A receptor subunits upon challenge of JAWSII with T. gondii (PRU-RFP) relative to unchallenged cells at indicated time-point (n = 3 independent experiments).

Figure 2—figure supplement 1.

Expression of GABA-A receptor subunits in JAWSII is modulated upon T. gondii infection.

(A) Relative mRNA expression (mean + SEM, 2-ΔCt) of GABA-A receptor subunits in unchallenged JAWSII, a mouse DC line, (n = 3 independent experiments). (B) Heat map shows (%) transcriptional changes in the expression of GABA-A receptor subunits upon challenge of JAWSII with T. gondii (PRU-RFP) relative to unchallenged cells at indicated time-point (n = 3 independent experiments).

Selective pharmacological inhibition of GABA-A Rs abrogates T. gondii-/N. caninum-induced hypermotility in human and murine phagocytes

To functionally assess the putative implication of GABA-A Rs in parasite-induced migratory activation of DCs, motility assays were performed in the presence of general and subunit-selective GABA-A R antagonists and modulators. A broad range GABA-A R open-channel blocker (picrotoxin) and α, β and ρ subunit selective inhibitors (L655 708, SCS and TPMPA, respectively), efficiently inhibited T. gondii-/N. caninum-induced hypermotility in mBMDCs (Figure 3A,C, Figure 3—figure supplement 1A,B), hMoDCs (Figure 3B,D) and, freshly isolated hMDCs and monocytes (Figure 1—figure supplement 1D,E, Figure 3—figure supplement 1C), with a non-significant impact on baseline cell motility. Further, we took advantage of the finding that blockade of GABA transporters (SNAP) inhibited hypermotility in parasitized mBMDCs (Figure 3E,F) with reduced GABA concentrations in the supernatants (Figure 5F) to attempt reconstitution of hypermotility by allosteric modulators of GABA-A Rs. Importantly, hypermotility was rescued by an allosteric modulator of β2/β3-containing GABA-A Rs (etomidate), but not by an allosteric modulator of δ subunit-containing GABA-A Rs (allopregnanolone) (Figure 3E,F). Because mBMDCs transcriptionally express β2, β3 and δ subunits (Figure 2), the data jointly indicated an implication of β subunits in hypermotility and also differential effects or implication by specific subunits. Together, this indicated that GABA-A Rs are implicated in the motogenic activation of phagocytes upon coccidian challenge and, specifically raised the question of participation of α, β and ρ GABA-A Rs subunits in hypermotility of phagocytes.

Figure 3.

Impact of pharmacological modulation of GABA-A Rs on T. gondii-induced hypermotility of DCs.

(A, B) Representative motility plots of unchallenged mBMDCs and T. gondii (PRU-RFP)-infected (A) mBMDCs and (B) hMoDCs treated with GABA-A R inhibitor picrotoxin (open channel blocker) or subunit specific inhibitors L655,708 (α-specific), SCS (β-specific) and TPMPA (ρ-specific) at concentrations stated in Materials and methods. X- and y-axes indicate distances in μm. (C, D) Box-and-whisker dot plots show, for each condition, median velocities (μm/min) as indicated in (A) and (B) (n = 3–6 independent experiments). (E) Representative motility plots and (F) velocities of unchallenged mBMDCs and T. gondii-infected mBMDCs treated with SNAP (GAT inhibitor) in presence of etomidate and allopregnanolone (allosteric modulators of β and δ subunit-containing GABA-A Rs, respectively). (n = 3–4 independent experiments). Statistical significance was tested by ordinary one-way ANOVA with Dunnett’s multiple comparison test for (C, D, F), ***p<0.001, ns p≥0.05.

(A, B) Box-and-whisker dot plots show median velocities (μm/min) of mBMDCs challenged with N. caninum Nc-1 strain (A) and Nc-Liverpool strain (B) and treated with GABA-A R inhibitors, NKCC1 inhibitor or VDCC inhibitors as indicated, respectively (n = 3 independent experiments). (C) Box-and-whisker dot plots show median velocities (μm/min) of hMonocytes challenged with T. gondii (PRU) and treated with GABA-A R inhibitor, NKCC1 inhibitor or VDCC inhibitor, respectively (n = 3 independent experiments). Statistical significance was assessed by ANOVA with Dunnett’s multiple comparison test for (A, B, C), ***p<0.001.

Figure 3—figure supplement 1.

Effect of pharmacological inhibitors of GABA-A Rs, NKCC1 and VDCCs on hypermotility.

(A, B) Box-and-whisker dot plots show median velocities (μm/min) of mBMDCs challenged with N. caninum Nc-1 strain (A) and Nc-Liverpool strain (B) and treated with GABA-A R inhibitors, NKCC1 inhibitor or VDCC inhibitors as indicated, respectively (n = 3 independent experiments). (C) Box-and-whisker dot plots show median velocities (μm/min) of hMonocytes challenged with T. gondii (PRU) and treated with GABA-A R inhibitor, NKCC1 inhibitor or VDCC inhibitor, respectively (n = 3 independent experiments). Statistical significance was assessed by ANOVA with Dunnett’s multiple comparison test for (A, B, C), ***p<0.001.

Gene silencing of GABA-A R subunits inhibits DC hypermotility

To determine which GABA-A R subunits were implicated in hypermotility, we designed a gene silencing approach in primary DCs. First, based on subunit expression data (Figure 2) and on the impact of selective pharmacological inhibitors on migration (Figure 3), we designed probes (Supplementary file 3) to knock-down the mRNA expression of α3, β3 and ρ1 subunits in mBMDCs and the α4 and ρ2 subunits in hMoDCs, respectively. Second, the shRNA probes were validated in neuronal cell lines known to express GABA-A Rs (Figure 4—figure supplement 1A–E) and significantly silenced target transcription in mBMDCs (Figure 4A) and hMoDCs (Figure 4B). Finally, the impact of gene silencing on hypermotility was assessed. The hypermotility phenotypes remained unaffected in mock-transduced and control shRNA-treated conditions (Ólafsson et al., 2019; Figure 4C,D,E,F). In contrast, hypermotility was abolished in shβ3- and shρ1-treated mBMDCs (Figure 4C,D,G), demonstrating the implication of β3 and ρ1 subunits. In shα3-treated cells (with two separate constructs), hypermotility was significantly reduced, albeit not abolished, indicating a contribution by the α3 subunit. In hMoDCs, hypermotility was significantly reduced in shα4-treated cells, but not in shρ2-treated cells (Figure 4E,F,G), indicating primarily a dependence on the α4 subunit for hypermotility. Jointly with pharmacological inhibition, these data demonstrate a critical dependency of DC hypermotility on the β3 and ρ1 subunits for mBMDCs and on the α4 subunit for hMoDCs.

Figure 4—figure supplement 1.

ShRNA transduction efficiency.

(A–B) Representative micrographs show (A) mBMDCs and the murine neuronal cell line NE4C, and (B) hMoDCs and the human neuronal cell line SH-SY5Y treated with control shRNA (shLuc) or shRNA to indicated target gene, respectively. (C, D) Bar graphs show mean % (+ SEM) GFP+ cells, representative of the lentiviral transduction efficiency quantified in mBMDCs and NE4Cs (C) and hMoDCs and SH-SY5Y (D). (E) Bar graph shows mean (+ SEM) mRNA knock-down (%) assessed in unchallenged transduced NE4Cs. Statistical significance was assessed by Student’s t-test, *p<0.05.

Figure 4.

Targeted gene silencing of GABA-A R subunits impacts DC hypermotility.

(A, B) The mRNA expression in unchallenged (A) mBMDCs and (B) hMoDCs, treated with shRNA for control Luc and GABA-A R subunits, related to mock-treated cells (Mean + SEM, %). For shα3, two separate constructs were used (n = 3–5 independent experiments for mBMDCs and 6–7 for hMoDCs). (C, E) Representative motility plots of (C) mBMDCs and (E) hMoDCs, treated as in (A–B) and challenged with T. gondii (PRU-RFP). X- and y-axes indicate distances in μm. (D, F) Histograms of accumulated distances migrated (μm) by (D) mBMDCs and (F) hMoDCs as in (C and E), respectively. Dotted lines indicate median values (n = 3–4 independent experiments). (G) Box-and-whisker dot plots show, for each indicated condition, median velocities (μm/min) of unchallenged and T. gondii-infected mBMDCs and hMoDCs (n = 3–4 independent experiments). Statistical significance was tested by ordinary one-way ANOVA with Dunnett’s multiple comparison test for (A, B, G) and by Mann-Whitney test for (D, F), *p<0.05, **p<0.01, ***p<0.001, ns p≥0.05.

(A–B) Representative micrographs show (A) mBMDCs and the murine neuronal cell line NE4C, and (B) hMoDCs and the human neuronal cell line SH-SY5Y treated with control shRNA (shLuc) or shRNA to indicated target gene, respectively. (C, D) Bar graphs show mean % (+ SEM) GFP+ cells, representative of the lentiviral transduction efficiency quantified in mBMDCs and NE4Cs (C) and hMoDCs and SH-SY5Y (D). (E) Bar graph shows mean (+ SEM) mRNA knock-down (%) assessed in unchallenged transduced NE4Cs. Statistical significance was assessed by Student’s t-test, *p<0.05.

Abolished DC hypermotility by inhibition of GABA synthesis (GAD67) and secretion is rescued by GABA-A R agonism

The constitutive elements of a GABA synthesis and transport in human DCs have remained uncharacterized (Fuks et al., 2012). In addition to GABA synthesis enzymes (Figure 1G), hMoDCs and mBMDCs expressed GABA transporters (Figure 5A, C), whose expression was modulated by challenge with T. gondii (Figure 5B, D). Pharmacological inhibition of GABA synthesis (SC) (Figure 5E, H) or transportation (SNAP) (Figure 5E, G) inhibited hypermotility and reduced GABA concentrations (Figure 5F) in the supernatants of hMoDCs, as previously shown in mBMDCs (Fuks et al., 2012). Further, exogenous GABA and GABA-A R agonism (muscimol) fully reconstituted hypermotility in hMoDCs in presence of GABA synthesis inhibitor (Figure 5E, H). This, together with the identification of GAD67 as putative principal GABA synthesis enzyme in hMoDCs (Figure 1G) motivated gene silencing of GAD67 (Figure 5I, Figure 4—figure supplement 1B,D, Supplementary file 3). Importantly, the amounts of secreted GABA in the supernatant were strongly reduced in GAD67-silenced infected hMoDCs, approached amounts secreted by unchallenged cells and contrasted with maintained GABA secretion by mock- or shLuc-transduced infected cells (Figure 5J). Finally, GAD67-silenced infected hMoDCs exhibited abrogated hypermotility (Figure 5K). Jointly, these data demonstrate a critical dependence of the hypermigratory phenotype on GABA synthesized by GAD67 in hMoDCs.

Figure 5.

Impact of GABA synthesis enzymes and GABA transporters on hypermotility.

(A, C) Relative mRNA expression (2-ΔCt) of GABA transporters in unchallenged (A) mBMDCs and (C) hMoDCs (n = 3 independent experiments). (B, D) Heat maps show (%) transcriptional changes in the expression of GABA transporters upon challenge of (B) mBMDCs and (D) hMoDCs with T. gondii (PRU-RFP) relative to unchallenged cells at indicated time-points. (X) indicates no amplification (n = 3–4 independent experiments). (E) Representative motility plots of unchallenged and T. gondii-infected hMoDCs treated with GABA transporter inhibitor (SNAP) and GABA synthesis inhibitor (SC) in absence and presence of GABA or GABA analog (muscimol). X- and y-axes indicate distances in μm. (F) GABA (ng/ml) secreted in supernatants of hMoDCs challenged with T. gondii (ME49/PTG) in presence of SNAP was quantified by ELISA (n = 3 independent experiments). (G, H) Box-and-whisker dot plots show, for each indicated condition, velocities (μm/min) of unchallenged and T. gondii-infected hMoDCs, as in (E) (n = 3 independent experiments). (I) The mRNA expression (2-ΔCt) in control shLuc-treated and shGAD67-treated unchallenged hMoDCs related (%) to mock-treated cells (n = 5 independent experiments). (J) GABA secreted (ng/ml) in supernatants of mock-, shLuc- and shGAD67-treated hMoDCs challenged with T. gondii (PRU-RFP) (n = 5 independent experiments). (K) Histograms show accumulated distances migrated (μm) by control shLuc-treated and shGAD67-treated hMoDCs, respectively, challenged with T. gondii (PRU-RFP). Dotted lines indicate median values (n = 3–4 independent experiments). Bar graphs show mean + SEM. Statistical significance was tested by paired t-test for (F), ordinary one-way ANOVA with Dunnett’s multiple comparison test for (G, H, I, J) and Mann-Whitney test for (K), *p<0.05, **p<0.01, ***p<0.001, ns p>0.05.

The GABA signaling regulator NKCC1 impacts hypermotility of phagocytes

GABA-A R function in the CNS is regulated by CCCs, which are subdivided in Na-K-Cl cotransporters (NKCCs) and K-Cl cotransporters (KCCs) (Kaila et al., 2014). However, CCCs have remained uncharacterized in phagocytes. A transcriptional expression screen (Supplementary file 1) detected mRNAs of NKCCs and KCCs in mBMDCs (Figure 6A) and in hMoDCs (Figure 6B). Notably, upon T. gondii challenge, a strong upregulation of NKCC1 was observed in mBMDCs, which was less accentuated in hMoDCs (Figure 6C,D). Immunolabeling and western blotting indicated presence of NKCC1/2 proteins in mBMDCs (Figure 6E,F). Interestingly, antagonism with bumetanide, at concentrations known to inhibit NKCC but not KCC activity in neurons (Orlov et al., 2015), resulted in impaired T. gondii-/N. caninum-induced hypermotility in mBMDCs (Figure 6G,I, Figure 3—figure supplement 1A,B), hMoDCs (Figure 6H,J), hMDCs and monocytes (Figure 1—figure supplement 1D,E, Figure 3—figure supplement 1C). To test this further, we applied gene silencing on NKCC1 (Figure 6K,L, Supplementary file 3) after validation in cell lines (Figure 4—figure supplement 1A–E). Importantly, hypermotility was abolished in NKCC1-silenced mBMDCs and hMoDCs (Figure 6M,N). Next, we addressed if NKCC1 and GABA-A R functions were interconnected. First, we observed that stimulation with GABA or with the GABA-A R agonist muscimol failed to reconstitute hypermotility of mBMDCs in the presence of the NKCC1 antagonist bumetanide (Figure 6O), contrasting with the reconstitution of hypermotility by GABA and muscimol in cells with abrogated GABA synthesis (Figure 5G). Second, shNKCC1- or shGAD67-transduced hMoDCs were stimulated with GABA. Upon silencing of GABA synthesis (shGAD67), addition of exogenous GABA reconstituted hypermotility. In contrast, in NKCC1-silenced cells, GABA failed to reconstitute hypermotility (Figure 6P,Q, Figure 6—figure supplement 1). These data indicate a link between NKCC1 and GABA/GABA-A R function in parasitized phagocytes. Altogether, the data demonstrate a functional implication of the Na-K-Cl cotransporter NKCC1 in T. gondii-/N. caninum-induced hypermotility of phagocytes.

Figure 6.

NKCC1 is a determinant of GABAergic hypermotility.

(A, B) Relative mRNA expression (2-ΔCt) of cation chloride transporters (CCCs) in unchallenged (A) mBMDCs and (B) hMoDCs (n = 3 independent experiments). (C, D) Heat map depicts (%) transcriptional expression changes of CCCs in (C) mBMDCs and (D) hMoDCs challenged with T. gondii (PRU-RFP) relative to unchallenged cells at indicated time points. (X) indicates no amplification (n = 3 independent experiments). (E) Immunostaining of mBMDCs challenged with T. gondii (ME49/PTG-GFP) stained with NKCC1/2 monoclonal antibody (Alexa Flour 594-conjugated anti-mouse secondary antibody) and DAPI (nuclei). Scale bars: 10 μm. (F) Representative Western blot of lysates from T. gondii-challenged mBMDCs for indicated time, immunoblotted with phospho-NKCC1 and total NKCC1/2 antibodies. GAPDH was used as loading reference. (n = 4 independent experiments). (G, H) Representative motility plots of unchallenged and T. gondii-infected (G) mBMDCs and (H) hMoDCs treated with NKCC1 inhibitor (bumetanide). X- and y-axes indicate distances in μm. (I, J) Box-and-whisker dot plots show, for each indicated condition, median velocities (μm/min) of unchallenged and T. gondii-infected (I) mBMDCs and (J) hMoDCs as in (G, H) (n = 3 independent experiments). (K, L) The mRNA expression (2-ΔCt) of control shLuc- and shNKCC1-treated unchallenged (K) mBMDCs and (L) hMoDCs related (%) to mock-treated cells (n = 7 independent experiments for mBMDCs and n = 4 for hMoDCs). Bar graphs show mean + SEM. (M, N) Histograms show accumulated distances migrated (μm) by control shLuc-treated and shNKCC1-treated (M) mBMDCs and (N) hMoDCs, respectively, challenged with T. gondii (PRU-RFP). Dotted lines indicate median values (n = 3 independent experiments). (O) Velocities of unchallenged and T. gondii-infected mBMDCs treated with GABA or muscimol in presence of bumetanide. (P) Representative motility plots of T. gondii-infected shGAD67- and shNKCC1-treated hMoDCs in presence of GABA. (Q) Velocities of unchallenged and T. gondii-infected shLuc-, shGAD67- and shNKCC1-treated hMoDCs with or without GABA. Statistical significance was tested by ordinary one-way ANOVA with Dunnett’s multiple comparison test for (I, J, K, L, O, Q) and by Mann-Whitney test for (M, N), *p<0.05, **p<0.01, ***p<0.001, ns p≥0.05.

Dotted lines indicate median values (n = 3 independent experiments). Statistical significance was tested by Mann-Whitney test, *p<0.05, ns p≥0.05.

Figure 6—figure supplement 1.

Histograms of accumulated distances migrated (μm) by mock- or shLuc-treated unchallenged and T. gondii-infected hMoDCs (upper panel), T. gondii-infected shGAD67- or shNKCC1-treated hMoDCs with or without GABA (lower panel).

Dotted lines indicate median values (n = 3 independent experiments). Statistical significance was tested by Mann-Whitney test, *p<0.05, ns p≥0.05.

The VDCC CaV1.3 mediates hypermotility and transient extracellular Ca2+ influx into DCs in response to GABA

We recently reported VDCC expression by murine DCs (Kanatani et al., 2017). To address the roles of VDCCs in human DCs, we challenged hMoDCs and hMDCs with T. gondii. First, we detected mRNA expression of 9 out of 10 known VDCC subtypes in hMoDCs, with highest relative expression of CaV1.3, 1.4 and 3.1 (Figure 7A, Supplementary file 1, 2). Moreover, challenge with T. gondii led to a prominent upregulation of CaV1.3, 2.2 and 2.3 mRNAs (Figure 7B). This result motivated pharmacological inhibition, including broad VDCC inhibition (benidipine), inhibition of CaV1 subtypes/L-type VDCCs (nifedipine) and targeted inhibition of CaV1.3 (CPCPT) (Yao et al., 2006; Kang et al., 2012). Importantly, VDCC inhibition abolished T. gondii-/N. caninum-induced hypermotility in hMoDCs (Figure 7C,D), hMDCs, monocytes and mBMDCs (Figure 1—figure supplement 1D,E, Figure 3—figure supplement 1A,B,C). Moreover, a structural analogue of nifedipine with positive ionotropic effect (Bay K8644) reconstituted hypermotility of infected hMoDCs in the presence of GABA synthesis inhibitor (SC) (Figure 7E) and also significantly enhanced motility in unchallenged hMoDCs (Figure 7D). The prominent effect of CaV1.3 inhibition prompted us to silence CaV1.3 expression (Figure 7F, Figure 4—figure supplement 1B,D, Supplementary file 3), which yielded abolished hypermotility (Figure 7G,H).

Figure 7.

CaV1.3 is the mediator for downstream effects of activated GABA signaling.

(A) Relative mRNA expression (2-ΔCt) of voltage-dependent calcium channels (VDCCs) in unchallenged hMoDCs (n = 3 independent experiments). (B) Heat map shows (%) transcriptional changes in the expression of VDCCs subunits upon challenge of hMoDCs with T. gondii (PRU-RFP) relative to unchallenged cells at indicated time-point. (X) indicates no amplification (n = 3 independent experiments). (C) Representative motility plots of unchallenged and T. gondii-infected hMoDCs treated with benidipine (VDCC broad inhibitor), nifedipine (L-type inhibitor), CPCPT (CaV1.3 specific inhibitor) and bay K8644 (VDCC activator). X- and y-axis are distances in μm. (D) Box-and-whisker dot plots show, for each indicated condition, velocities (μm/min) of unchallenged and T. gondii-infected hMoDCs, as in (C), and (E) treated with SC (GABA synthesis inhibitor) in presence of bay K8644 (n = 3 independent experiments). (F) The mRNA expression in control shLuc- and shCaV1.3-treated unchallenged hMoDCs related (%) to mock-treated cells (n = 10 independent experiments). (G) Representative motility plots of hMoDCs, treated as in (F) and challenged with T. gondii (PRU-RFP). (H) Histograms of accumulated distances migrated (μm) by hMoDCs as in (G). Dotted lines indicate median values (n = 3 independent experiments). Bar graphs show mean + SEM. Statistical significance was tested by ordinary one-way ANOVA with Dunnett’s multiple comparison test for (D, E, F) and by Mann-Whitney test for (H), *p<0.05, ***p<0.001, ns p≥0.05.

To test the implications of GABAergic signaling on Ca2+ responses, Ca2+ indicator dye-loaded mBMDCs were challenged with agonistic and antagonistic stimuli and Ca2+ responses were measured (Figure 8A, Video 1). Importantly, perfusion of exogenous GABA consistently generated transient cytosolic Ca2+ elevations, which were antagonized by the GABA-A R blocker picrotoxin (broad inhibitor) and with maintained responsiveness by purinergic Ca2+ channels to ATP (Figure 8B,C,D; Video 1). Further, transients generated by L-type VDCC agonism (Bay K) were effectively antagonized by CaV1.3 inhibition (CPCPT), indicative of a prominent role for CaV1.3 in the measured Ca2+ responses (Figure 8E,F,G, Figure 8—figure supplement 1). Finally, T. gondii-infected cells responded to GABA with transient Ca2+ influx. In individual cell recordings, Ca2+ influx was significantly reduced or abolished by application of inhibitors of β-subunit containing GABA-A Rs (SCS) and by picrotoxin, with maintained responses to ATP (Figure 8H,I,J). Jointly, these data identify a determinant role for GABA-A Rs in the GABA-induced Ca2+ influx via CaV1.3 in DC hypermotility.

Figure 8.

Ca2+ responses in unchallenged and T. gondii-infected DCs upon agonism and antagonism of GABA-A Rs and VDCCs.

(A) Representative time-lapse micrographs show live cell Ca2+ imaging of one single mBMDC preloaded with Ca2+ indicator dye (Fluo-8H AM) and sequentially perfused with GABA (1 mM), GABA (1 mM) + picrotoxin (100 μM) and ATP (150 μM). Colored lines indicate perfusion times (min:s) of treatments. Color scale depicts relative fluorescence intensity. (B) Representative live cell Ca2+ recording traces (relative fluorescence intensity, F/Fmedian) from four cells plotted against time (min) as in (A). Lines indicate respective perfusions times. (C) Mean Ca2+ response intensity (black dots) and SEM (gray whiskers) from one recording with 105 cells, plotted against time as in (B), and (D) maximum Ca2+ response intensity of 294 cells from three independent experiments. (E) Representative live cell Ca2+ recording traces from four cells sequentially perfused with bay K 8644 (40μM), bay K 8644 (40μM) + CPCPT (100 μM), 2nd application of bay K 8644 (40μM) and ATP (150 μM) at times indicated by lines. (F) Mean Ca2+ response intensity (black dots) and SEM (gray whiskers), from 72 cells plotted against time as in (E), and (G) maximum Ca2+ response intensity of 233 cells from three independent experiments. (H) Representative live cell Ca2+ recording traces from T. gondii (ME49-RFP)-infected cells sequentially perfused with GABA (1 mM), GABA (1 mM) + SCS (100 μM), GABA (1 mM), GABA (1 mM) + picrotoxin (100 μM) and ATP (150 μM) at times indicated by lines. (I) Mean Ca2+ response intensity (black dots) and SEM (gray whiskers) from 44 cells, plotted against time as in (H), and (J) maximum Ca2+ response intensity of 115 cells from three independent experiments. Bar graphs show mean + SEM. Statistical significance was assessed by ordinary one-way ANOVA with Dunnett’s multiple comparison test for (D, G, J), **p<0.01.

Representative time-lapse micrographs show live cell Ca2+ imaging of unchallenged single mBMDCs preloaded with Ca2+ indicator (Fluo-8H AM) and sequentially perfused with bay K 8644 (40μM), bay K 8644 (40μM) + CPCPT (100 μM), 2nd application of bay K 8644 (40μM) and ATP (150 μM). Colored lines indicate perfusion times (min:s) of treatments. Color scale depicts relative fluorescence intensity.

Video 1.

Representative time-lapsed micrographs from a live cell.

Ca2+ imaging recording of mBMDC preloaded with Ca2+ indicator dye (Fluo-8H AM) and sequentially perfused with GABA (1 mM), GABA (1 mM) + picrotoxin (100 μM) and ATP (150 μM) at 3.42–6.46, 10.35–13.32 and 17.08–19.52 min, respectively. The Fluo-8H AM fluorescence signal (green) is converted to rainbow color scale depicting relative fluorescence intensity ranging from blue (lowest) to red (highest). Scale bar 20 μm.

Figure 8—figure supplement 1.

Single-cell micrographs of Ca2+ influx show VDCC activation.

Representative time-lapse micrographs show live cell Ca2+ imaging of unchallenged single mBMDCs preloaded with Ca2+ indicator (Fluo-8H AM) and sequentially perfused with bay K 8644 (40μM), bay K 8644 (40μM) + CPCPT (100 μM), 2nd application of bay K 8644 (40μM) and ATP (150 μM). Colored lines indicate perfusion times (min:s) of treatments. Color scale depicts relative fluorescence intensity.

Gene silencing and pharmacological antagonism of GABAergic signaling slow DC migration and reduce parasite loads in mice

To address the impact of GABAergic signaling on DC-mediated parasite dissemination, we designed separate approaches that targeted GABA-A R function or the function of the GABA signaling regulator NKCC1 in mice. First, we controlled that inhibitors had non-significant effects on parasite invasion and replication (Figure 9—figure supplement 1A) and a persistent inhibitory effect on hypermotility (Figure 9—figure supplement 1B). Second, inhibitor-pretreated parasitized mBMDCs (CMTMR-labeled) and non-treated parasitized mBMDCs (CMF2HC-labeled) were simultaneously adoptively transferred to mice in a competition assay (Figure 9A). Fourteen to 18 h post-inoculation, organs were harvested and cells were characterized by flow cytometry (Figure 9B,C, Figure 9—figure supplement 2A, B,C). Importantly, the ratio of pretreated vs non-treated parasitized mBMDCs was significantly lower in spleen for both GABA-A R inhibitor and NKCC1 inhibitor treatments compared with that for non-treated condition (Figure 9D). In contrast, non-significant differences were observed in peritoneum (Figure 9D). This indicated selective reduced migration of mBMDCs pretreated with GABA-A R inhibitors or NKCC1 inhibitor compared with non-treated mBMDCs in individual mice. Third, to asses if the reduced migration of parasitized mBMDCs impacted the infection, parasite loads were assessed by plaquing assays and by qPCR. Importantly, compared with the non-treated condition, treatments significantly decreased the parasite loads in spleen and liver (Figure 9E, Figure 9—figure supplement 2D, E). Finally, we silenced the GABA-A R subunit ρ1 or NKCC1, two GABA signaling targets with a significant impact on mBMDC migration in vitro (Figure 4, Figure 6). When gene-silenced mBMDCs challenged with T. gondii were adoptively transferred into mice, significantly reduced parasite loads were quantified in peripheral organs (Figure 9F), corroborating results of pharmacological treatments (Figure 9E). Moreover, pharmacological inhibition of GABA-A Rs or NKCC1 yielded a significant reduction of parasites loads in the brain by day seven post-inoculation (Figure 9G). The data demonstrate a role for the GABA-A R subunit ρ1 and the GABA signaling regulator NKCC1 in mBMDC-mediated dissemination of T. gondii. Altogether, the findings support the notion that GABAergic signaling promotes the dissemination of T. gondii via parasitized DCs.

Figure 9—figure supplement 1.

Effect of GABAergic inhibition on parasite replication in DCs.

(A) Bar graphs show parasite replication assessed in T. gondii-infected mBMDCs in presence of picrotoxin (50 μM), bumetanide (40 μM) and geneticin (10 μM), a blocker of polypeptide synthesis in eukaryotic cells, as a positive control. Number of parasites per vacuole is plotted against % of total vacuoles counted at indicated time points, as described in Materials and methods, (n = 3 independent experiments). (B) Box-and-whisker dot plots show, for each indicated condition, velocities (μm/min) of unchallenged and T. gondii-infected mBMDCs treated with picrotoxin for 4 h and 8 h or treated for 1 h, washed and recorded 3 h and 7 h post washing (n = 3 independent experiments). Bar graphs show mean + SEM. Statistical significance was assessed by ordinary one-way ANOVA with Tukey’s multiple comparison test, **p<0.01, ***p<0.001, ns p≥0.05.

Figure 9.

In vivo impact of GABAergic inhibition on DC migration and parasite loads.

(A) Schematic illustration of simultaneous adoptive transfers of T. gondii (ME49/PTG-GFP)-challenged mBMDCs prelabeled with CMF2HC or CMTMR dye and pretreated as described in Materials and methods, respectively. (B, C) Representative bivariate plots show CD11c+ cells from (B) peritoneal cavity and (C) spleen of C57BL/6 mice inoculated with T. gondii-challenged mBMDCs prelabeled with CMF2HC or CMTMR dye. Cells were analyzed by flow cytometry at 14–18 h post-inoculation (gating strategy in Figure 9—figure supplement 2A, B,C). Plots in upper, center and lower rows show, respectively, non-treated cells, cells pretreated with GABA-A R inhibitors (GABA-A R i) and NKCC1 inhibitor (NKCC1 i). (D) Bar graph shows the ratio of treated cells (CD11c+GFP+CMTMR+) to non-treated cells (CD11c+GFP+CMF2HC+) in peritoneal lavage and spleen after adoptive transfer of T. gondii-challenged mBMDCs (n = 7–10 mice per group). (E) Parasite loads in spleen and liver of C57BL/6 mice at day 4 post-inoculation of pharmacologically treated cells (GABA-A R i, NKCC1 i) related to non-treated cells and measured by plaquing assays (n = 5–6 mice per group). (F) Parasite loads in spleen and liver of C57BL/6 mice at day 5 post-inoculation of gene-silenced cells (shρ1, shNKCC1) related to mock- and control shLuc-transduced cells and measured by plaquing assays (n = 6 mice per group). (G) Parasite loads in spleen and brain of CD1 mice at day 7 post-inoculation of pharmacologically treated cells (GABA-A R i, NKCC1 i) related to non-treated cells and measured by plaquing assays (n = 9–10 mice per group). Bar graphs show mean + SEM. Statistical significance was assessed by ordinary one-way ANOVA with Tukey’s multiple comparison test, *p<0.05, **p<0.01, ***p<0.001, ns p≥0.05.

(A) Bar graphs show parasite replication assessed in T. gondii-infected mBMDCs in presence of picrotoxin (50 μM), bumetanide (40 μM) and geneticin (10 μM), a blocker of polypeptide synthesis in eukaryotic cells, as a positive control. Number of parasites per vacuole is plotted against % of total vacuoles counted at indicated time points, as described in Materials and methods, (n = 3 independent experiments). (B) Box-and-whisker dot plots show, for each indicated condition, velocities (μm/min) of unchallenged and T. gondii-infected mBMDCs treated with picrotoxin for 4 h and 8 h or treated for 1 h, washed and recorded 3 h and 7 h post washing (n = 3 independent experiments). Bar graphs show mean + SEM. Statistical significance was assessed by ordinary one-way ANOVA with Tukey’s multiple comparison test, **p<0.01, ***p<0.001, ns p≥0.05.

(A, B) Cells collected at 14–18 h from (A) peritoneal cavity and (B) spleen were first gated for leukocyte populations (SSC-A vs. FSC-A), singlets (FSC-H vs. FSC-A) and further gated for expression of PECy7-CD11c. Further, this CD11c+ population was gated to identify adoptively transferred T. gondii-challenged mBMDCs (GFP+) pre-labeled with CMF2HC (non-treated cells) and CMTMR (treated with inhibitors of GABA-A Rs or NKCC1). (C) Representative bivariate plots show CD11c+ cells from spleen of non-infetced C57BL/6 mice gated to identify CMF2HC- or CMTMR-labeled cells. (D, E) Standard curves for B1 and AMA-1 genes of T. gondii, used to extrapolate parasite numbers in tissues by qPCR. The number of parasites per gram tissue, as in Figure 9D, was quantified by qPCR expression of B1 and AMA-1 (n = 5–6 mice per group). Bar graphs show mean + SEM. Statistical significance was assessed by ordinary one-way ANOVA with Tukey’s multiple comparison test, ***p<0.001, ns p≥0.05.

Figure 9—figure supplement 2.

Gating strategy for flow cytometry analysis.

(A, B) Cells collected at 14–18 h from (A) peritoneal cavity and (B) spleen were first gated for leukocyte populations (SSC-A vs. FSC-A), singlets (FSC-H vs. FSC-A) and further gated for expression of PECy7-CD11c. Further, this CD11c+ population was gated to identify adoptively transferred T. gondii-challenged mBMDCs (GFP+) pre-labeled with CMF2HC (non-treated cells) and CMTMR (treated with inhibitors of GABA-A Rs or NKCC1). (C) Representative bivariate plots show CD11c+ cells from spleen of non-infetced C57BL/6 mice gated to identify CMF2HC- or CMTMR-labeled cells. (D, E) Standard curves for B1 and AMA-1 genes of T. gondii, used to extrapolate parasite numbers in tissues by qPCR. The number of parasites per gram tissue, as in Figure 9D, was quantified by qPCR expression of B1 and AMA-1 (n = 5–6 mice per group). Bar graphs show mean + SEM. Statistical significance was assessed by ordinary one-way ANOVA with Tukey’s multiple comparison test, ***p<0.001, ns p≥0.05.

Discussion

Signaling pathways that can drive migration of immune cells, and are alternative to canonical chemokine-mediated migration, have remained poorly understood. Here, we establish that human and murine mononuclear phagocytes possess a conserved GABAergic system that, upon activation, promotes migration in vitro and in vivo. We identified and performed functional tests on the five principal components of GABAergic signaling, namely (i) GABA metabolism, (ii) GABA transportation and secretion, (iii) GABA-A R activation, (iv) GABA signaling regulators CCCs and (v) effector Ca2+ channel signaling by VDCCs (Figure 10). The data provide a molecular and cellular framework for assessing the role of the GABAergic system in immune cells.

Figure 10.

GABAergic signaling in mononuclear phagocytes with an impact on cell migration and coccidian parasite dissemination.

Schematic representation illustrates the molecular GABAergic signaling components (1-5) identified in human and murine mononuclear phagocytes. Their functions and experimental targeting approaches are detailed in the tabular representation, respectively. (6) Ca2+ influx sets the cell in a hypermotile state by activation of MAP kinases and cytoskeletal rearrangements (Ólafsson et al., 2020). In the tabular representation, ()Red colored text indicates molecular components commonly expressed by mouse and human phagocytes. Blue and black color indicates components only detected in mouse and human cells, respectively. (Indicates genes targeted by shRNA with an impact on cell migration in vitro (*), as described under Materials and methods. (*) Red asterisks indicate conditions additionally tested in vivo in mice. ()Indicates pharmacological agonists and antagonists with an impact on phagocyte motility in vitro tested on both mouse and human phagocytes. (*) Blue asterisks indicate conditions additionally tested in vivo in mice. n.d.: not determined.

We demonstrate that a conserved expression of GABAergic molecular components is functionally linked to motility and migratory activation of mononuclear phagocytes upon infection challenge. First, our studies identify GAD67 as the principal GABA synthesizing enzyme in phagocytes. Gene silencing and pharmacological inhibition of GAD67 abrogated secretion of GABA and migratory activation of phagocytes, which was reconstituted by GABA-A R agonism. This supports the notion that GABA is synthesized cytosolically and secreted in vesicle-independent fashion, likely by transport through GATs, for tonic modulations of GABA-A Rs in immune cells, similar to neurons (Kaufman et al., 1991; Feldblum et al., 1993). Second, our data establish that expression of specific GABA-A R subunits determine the motogenic function of GABA-A Rs in phagocytes. The expression of GABA-A R subunit types was diverse, in line with the expression diversity in neurons (Davis et al., 2000; Goetz et al., 2007). Yet, the different phagocyte types consistently expressed a repertoire of GABA-A R subunits sufficient to constitute functional channels, that is: at least one α, one β and one third type of subunit, or homopentamer-forming ρ subunits. The inhibitory effects by selective pharmacological antagonism on phagocyte hypermotility indicated implication of α, β and ρ subunits and was confirmed by gene silencing. Importantly, silencing of the α4 subunit (but not ρ2) or β3 and ρ1 subunits (but not α3) abolished hypermigration of human and murine DCs, respectively. Jointly, this narrows putative receptor pentamers acting in phagocytes but also highlights a hierarchy among GABA-A R subunits mediating migratory activation or function redundancy among the different subunits. Third, our data identify a determinant role for the CCC NKCC1 in the migratory activation of phagocytes. By pharmacological inhibition and gene silencing in vitro and in vivo, we show that NKCC1 plays a crucial role in the regulation of GABA signaling in phagocytes. Of note, NKCC1 was linked to GABA-A R function and its regulative characteristics together with limited variability -compared with the high diversity in expression of GABA-A R subunits- made NKCC1 a prime target for in vivo experimentation. Finally, we demonstrate that stimulation with GABA elicits Ca2+ influx transients in the DC cytosol. In line with this, human and murine phagocytes expressed a highly conserved repertoire of VDCC subtypes. Moreover, silencing of the VDCC subtype CaV1.3 in human DCs abrogated T. gondii-induced hypermotility, in line with our observations in murine DCs (Kanatani et al., 2017). Jointly, this demonstrates that CaV1.3 is determinant to the motogenic action of GABA-A R activation.

Our data establish that T. gondii and N. caninum, two coccidian parasites with a broad range of vertebrate hosts, induce GABAergic signaling in parasitized phagocytes. From a perspective of intracellular parasitism, hijacking a conserved GABAergic system offers the advantage of inducing migratory activation of shuttle phagocytes within minutes after active invasion (Weidner et al., 2013) to favor systemic dissemination in vertebrate hosts (Lambert et al., 2006; Courret et al., 2006). Activation of GABAergic signaling is relatively fast as GABA binding opens the GABA-A Rs in milliseconds (Farrant and Nusser, 2005) and the elements that synthesize and transport GABA, GADs and GATs, respectively, are constitutionally expressed and upregulated upon infection. Consequently, DCs initiated GABA secretion shortly after T. gondii invasion. Thus, the rapid onset and tight regulation of GABAergic signaling by-passes the need for, presumably slower, transcriptional regulation. Additionally, the GABA signaling regulator NKCC1 plays an important role in the hypermigration of parasitized phagocytes, presumably by impacting chloride concentration and thus, GABA-A R function. Consequently, inhibition of GABA-A Rs or NKCC1 in adoptively transferred pharmacologically pretreated or gene silenced DCs, significantly reduced the migration of parasitized DCs and parasite loads in mice. Importantly, hampered dissemination to peripheral vital organs was evident early during infection and, later, resulted in reduced parasite loads in the CNS. Jointly with present data, different approaches show that targeting (i) GABA synthesis/transportation (Fuks et al., 2012), (ii) GABA-A R signaling, (iii) GABA-A R regulation/NKCC1 hampers systemic dissemination and parasite loads in the brain. However, inhibition of VDCC signaling inhibited systemic dissemination but non-significantly impacted parasite loads in the brain (Kanatani et al., 2017), indicating that specific GABAergic signaling components contribute differently or indicating redundancy in VDCC signaling. Because invasive coccidian parasites need to reconcile their obligate intracellular existence with the need for dissemination, the hijacking of migratory leukocytes represents a secluded replication niche that facilitates dissemination. The identified GABAergic determinants provide a molecular framework for assessing if other protozoa, intracellular bacteria (Kim et al., 2018) or viruses (Zhu et al., 2017) utilize the GABAergic signaling of phagocytes for dissemination. Because hypermigratory parasitized DCs exhibit chemotaxis (Weidner and Barragan, 2014) and GABA impacts the secretion of pro-inflammatory cytokines (Bhandage et al., 2018), the impact of GABAergic signaling in the inflammatory microenvironment of infection needs to be further investigated, also in the setting of acute and chronic infection and neuroinflammatory responses in the CNS (Bhandage et al., 2019). Thus, hypermigration and chemotaxis are not antithetical and may, in fact, cooperatively potentiate the migratory potential of parasitized phagocytes and therefore also the dissemination of coccidia (Fuks et al., 2012; Weidner et al., 2013; García-Sánchez et al., 2019).

Our study provides the first exhaustive characterization of a GABAergic machinery in myeloid mononuclear phagocytes. Recent findings also indicate GABAergic responses by macrophages (Januzi et al., 2018), microglia (Bhandage et al., 2019), lymphocytes (Bhandage et al., 2018) and bovine immune cells (García-Sánchez et al., 2019). Altogether, this highlights that GABAergic signaling by immune cells may be more the rule than the exception. Along these lines, GABA has a motogenic role in embryonic interneuron migration in the developing fetus (Bortone and Polleux, 2009). Furthermore, GABAergic signaling has newly been linked to the metastasis of multiple cancer types (Sizemore et al., 2014; Wu et al., 2014), including gliomas, pancreatic cancer and breast cancer (Neman et al., 2014; Takehara et al., 2007; Smits et al., 2012). The peripheral GABAergic system also appears implicated in various autoimmune diseases, such as multiple sclerosis (Bhat et al., 2010), type I diabetes (Li et al., 2017; Bhandage et al., 2018) and rheumatoid arthritis (Tian et al., 2011), where GABAergic inhibition dampens the inflammatory response. It will be important to assess if the motogenic molecular components identified here are also implicated in the inflammatory responses and in cancer cell metastasis. Moreover, the patho-physiological cellular microenvironments result in expression of specific subtypes of GABA receptors (Bhandage et al., 2014; Smits et al., 2012) and thus, selective compounds have been identified in neuropsychiatric drug design (Korpi and Sinkkonen, 2006; Krall et al., 2015). Additionally, anesthetics targeting GABA-A Rs have been implicated in the impairment of immune cell function during human surgery (Wheeler et al., 2011). Thus, receptor subtypes or other GABAergic components may be targeted to modulate migration of GABAergic cells (Miao et al., 2010) and our data provide a molecular framework for therapeutic targets of clinical relevance.

Finally, the non-protein amino acid GABA has developed into an essential neurotransmitter of the evolved vertebrate CNS. However, GABA precedes the development of the CNS as a metabolism- and stress-related signaling molecule in prokaryotes, invertebrates and plants (Hudec et al., 2015; MacRae et al., 2012; Pinan-Lucarré et al., 2014). Here, we add that mononuclear phagocytes express a conserved GABAergic system linked to their migratory functions, which coccidian parasites can hijack for dissemination. Thus, GABA also acts as an interspecies signaling molecule in host-microbe interactions.

Materials and methods

Experimental animals

C57BL/6NCrl mice (6–10 weeks old) and female Crl:CD1(ICR) mice (6–8 weeks old) were purchased from Charles River (Sulzfeld, Germany) and maintained and/or bred under pathogen-free conditions at Experimental Core Facility (ECF), Stockholm University, Sweden.

Parasites and cell lines

T. gondii lines used include GFP-expressing RH (type I), ME49/PTG (type II) and RFP-expressing PRU or ME49 (type II) (Kim et al., 2001; Hitziger et al., 2005). N. caninum lines used include NC-1 and NC-Liverpool (ATCC 50977 and ATCC 50845, American Type Culture Collection, Manassas, Virginia, US). Tachyzoites were maintained by serial 2-day passaging in human foreskin fibroblast (HFF-1, ATCC SCRC-1041) monolayers cultured in DMEM (Thermofisher scientific, Stockholm, Sweden) with 10% fetal bovine serum (FBS; Sigma-Aldrich, Darmstadt, Germany), gentamicin (20 µg/ml; Thermofisher), glutamine (2 mM; Thermofisher), and HEPES (0.01 M; Thermofisher), defined as complete medium (CM). The murine DC cell line JAWSII (ATCC CRL-11904) and neuroectodermal cell lines NE4Cs (ATCC CRL-2925) were cultured in DMEM supplemented with 10% FBS, gentamicin, glutamine, HEPES and human neuronal cell line SH-SY5Y (ATCC CRL-2266) was cultured in Opti-MEM supplemented with 10% FBS and gentamicin. All cultures were regularly tested for mycoplasma.

Primary cells

Mouse bone marrow-derived DCs (mBMDCs) were generated as previously described (Fuks et al., 2012). Briefly, bone marrow cells extracted from legs of 6–10 week-old C57BL/6 mice were cultivated in RPMI 1640 (Thermofisher) with 10% FBS, gentamicin, glutamine and HEPES, additionally supplemented with recombinant mouse GM-CSF (10 ng/ml; Peprotech, Stockholm, Sweden) at 37°C. Medium was replenished on days 2 and 4. Loosely adherent cells harvested on days 6–8 were used for experiments. Human monocytes were isolated from buffy coats, obtained from Karolinska University Hospital, by negative selection using RosetteSep Human Monocyte Enrichment Cocktail (Stemcell Technologies, Cambridge, UK). The population obtained exhibited CD14+ (DakoCytomation, Glostrup, Denmark) and <1% CD3+/19+ (BD Biosciences, Stockholm, Sweden), as evaluated by flow cytometry. For differentiation into hMoDCs, cells were cultivated in DMEM with 10% FBS, gentamicin, glutamine and HEPES, additionally supplemented with recombinant human GM-CSF (75 ng/ml; Peprotech) and IL-4 (30 ng/ml; Peprotech) at 37°C. Medium was replenished on day 3 and day 6. Loosely adherent cells harvested on day 6–9 were used for experiments. DCs were typified by expression of CD1a, CD11b, CD14 (DakoCytomation), CD80, CD83, CD86, HLA-DR, CD11a, CD18, CD54 (BD Biosciences). CD1c+ hMDCs isolated by human Myeloid Dendritic Cell Isolation Kit (Miltenyi Biotec, Bergisch Gladbach, Germany) were 70–97% pure, as characterized by flow cytometry.

Reagents

Picrotoxin (50 μM), L-655 708 (11,12,13,13a-Tetrahydro-7-methoxy-9-oxo-9H-imidazo[1,5-a]pyrrolo[2,1 c][1,4]benzodiazepine-1-carboxylic acid, ethyl ester, 10 μM), SCS (Salicylidene salicylhydrazide, 1 μM), TPMPA ((1,2,5,6-Tetrahydropyridin-4-yl) methylphosphinic acid, 50 μM), etomidate (10 μM), allopregnanolone (100 nM), muscimol (5 μM), bumetanide (10 μM), nifedipine (10 μM), benidipine (10 μM), bay K 8644 (10 μM), bicuculline (50 μM), ATP (150 μM; all from Tocris Bioscience, Bristol, UK), SNAP ((S)-Nitroso-N-acetylpenicillamine, 50 μM), SC (semicarbazide 50 μM), GABA (5 μM), geneticin (10 μM; all from Sigma-Aldrich) and CPCPT (1-(3-Chlorophenethyl)−3-cyclopentylpyrimidine-2,4,6-(1H,3H,5H)-trione, 1 μM, Merck Millipore, Darmstadt, Germany) were used at the indicated concentrations, if not differently stated. All pharmacological treatments and live cell stainings were performed in CM at 37°C and 5% CO2.

Motility assays

Cell motility analyses was performed as previously described (Fuks et al., 2012; Weidner et al., 2013). Briefly, cells were challenged with freshly egressed tachyzoites and subjected to pharmacological treatments for 4–6 h. Cells were seeded in 96-well plates and imaged every min for 60 min (Zeiss Observer Z.1). Motility tracks for 50–60 cells per treatment were analyzed using ImageJ software for each experiment.

Immunocytochemistry

mBMDCs were seeded on gelatin-coated glass coverslips for 0.5–1 h, fixed with 4% PFA in PBS for 15–20 min at RT and permeabilized using 0.1% Triton X-100 in PBS. To visualize host cell F-actin, cells were stained with Alexa Fluor 488- or 594-conjugated phalloidin (Invitrogen). To probe GABA-A R subunits, cells were incubated with rabbit anti-GABA-A R α3 polyclonal antibody, rabbit anti-GABA-A R α5 polyclonal antibody, rabbit anti-GABA-A R ρ1 polyclonal antibody (all from Alomone labs, Jerusalem, Israel), mouse anti-GABA-A R β3 monoclonal antibody (NeuroMab, UC Davis, CA, US) and for NKCC, with mouse anti-NKCC1/2 monoclonal antibody (clone T4, Developmental Studies Hybridoma Bank, DSHB, Iowa, USA) ON at 4°C. Following staining with respective Alexa Fluor 488- or 594-conjugated secondary antibodies (Thermofisher) and DAPI, coverslips were mounted and imaged by confocal microscopy (LSM 780 and LSM 800, Zeiss).

Real-time quantitative PCR

Total RNAs were extracted using Direct-zol miniprep RNA kits (Zymo Research, Irvine, CA, USA) with TRI reagent (Sigma-Aldrich) and first-strand cDNA was synthesized using Superscript IV or Maxima H Minus Reverse Transcriptase (Thermofisher) using a standard protocol. Real-time quantitative PCR (qPCR) was performed in QuantStudio 5 384 Optical well plate system (Applied Biosystem, Stockholm, Sweden) and/or LightCycler 480 (Roche, Basel, Switzerland) in a standard 10 μl with the 2X SYBR FAST qPCR Master Mix (Sigma-Aldrich) with gene-specific primers (Supplementary file 1) as described (Bhandage et al., 2019). Relative expression (2-ΔCt) were determined for each target in reference to a normalization factor, geometric mean of reference genes, either importin 8 (IPO8) and TATA-binding protein (TBP) or Actin-β (ACTB) and glyceraldehyde 3-phosphate dehydrogenase (GAPDH).

GABA enzyme-linked immunosorbent assays

ELISA (Labor Diagnostica Nord, Nordhorn, Germany) was performed as previously described (Fuks et al., 2012). Briefly, cells were plated at a density of 1 × 106 cells per ml and incubated for 24 h, unless differently stated, in presence of T. gondii and N. caninum tachyzoites. GABA concentrations in supernatants quantified at a wavelength of 450 nm (VMax Kinetic ELISA Microplate Reader, Molecular Devices, Vantaa, Finland).

Live cell time-lapse calcium imaging

Cells were loaded with, a calcium indicator, 2 μM Fluo-8H AM (AAT Bioquest, Sunnyvale, CA, USA) for 30 min at 37°C and seeded on 5% 3-aminopropyltriethoxysilane (TESPA, Sigma-Aldrich)-coated coverslip for 2–5 min at 37°C. Time lapse images were acquired by confocal microscopy (LSM 800, Zeiss) with 20X or 63X objective at an interval of 1 s per image. Drugs were diluted in RPMI 1640 (without phenol red) and perfused at indicated time and concentration by a peristaltic pump (1 ml/min). Absolute fluorescence intensity values (F) were extracted using ZENBlue software (Zeiss) and relative intensity (F/Fmedian) at a given time were analyzed for individual cells. Further, mean intensity of all cells was calculated.

Western blot

mBMDCs and mouse brain hippocampal tissue were lysed in RIPA buffer with protease and phosphatase inhibitor cocktail (Thermofisher), sonicated, diluted with 4 x laemmli buffer and boiled. Protein were subjected to SDS-PAGE on 8% polyacrylamide gels, transfered onto PVDF membrane (Merck Millipore), blocked in 2.5% BSA and incubated ON with rabbit anti-phospho-NKCC1 polyclonal antibody (Thr212/Thr217, Merck Millipore), mouse anti-NKCC1/2 monoclonal antibody (clone T4, DSHB) and rabbit anti-GAPDH antibody (Merck Millipore) followed by incubation with respective HRP-conjugated secondary antibodies (Cell signaling, Leiden, Netherlands). Protein bands were revealed by enhanced chemiluminescence reagents (Thermofisher) in a ChemiDoc system (BioRad, Stockholm, Sweden).

Lentiviral vector production and transduction

Self-inactivating Lentiviral particles were produced from Lenti-X 293 T cells (Takara Bio, Gothenburg, Sweden) by co-transfecting (i) a lentiviral vector, pLL3.7 or pLKO.1, containing self-complementary hairpin DNA oligos targeting specific mRNA (Supplementary file 3), (ii) psPAX2 packaging vector and (iii) pCMV-VSVg envelope vector, as described previously (Kanatani et al., 2017; Ólafsson et al., 2019). The lentiviral supernatants were used for transduction. The lentivirus transduction efficiency was examined in murine NE4Cs and human SH-Sy5y cell lines and further, knock-down efficiency was determined in NE4Cs (Figure 4—figure supplement 1). Murine BMDCs were transduced on day 3 of culturing, whereas human MoDCs were transduced day 3 and day 5. Transduction efficiency was examined for eGFP expression by epifluorescence microscopy, followed by expression analysis by qPCR for knock-down of targeted mRNA. All the target conditions were compared to mock-condition and a positive control, non-related sequence (luciferase, Luc). Transduced cells were further used for experiments.

Adoptive transfers of T. gondii-infected DCs

Adoptive transfers were performed as previously described (Kanatani et al., 2017). Briefly, (i) mBMDCs were challenged with freshly egressed tachyzoites (ME49/PTG-GFP, 6 h, MOI 1.5) and 1 h post-challenge, treated with picrotoxin (50 μM) and bicuculline (50 μM), for inhibition of GABA-A R or with bumetanide (40 μM), for inhibition of NKCC1 or (ii) Mock-, shLuc, shρ1 and shNKCC1 transduced mBMDCs were challenged with freshly egressed PRU-RFP tachyzoites (6 h, MOI 1.5). Extracellular parasites were removed by centrifugation. Following resuspension, T. gondii-infected mBMDCs (equivalent to 50,000 colony-forming units of T. gondii) were adoptively transferred into recipient C57BL/6 or Crl:CD1(ICR) mice by an intraperitoneal (i.p.) injection. C57BL/6 mice injected with pharmacologically treated cells or with transduced cells were sacrificed at days 4 and 5 post-infection, respectively. Further, Crl:CD1(ICR) mice injected with pharmacologically treated cells were sacrificed at day 7 post-infection.

For competition assays, mBMDCs were challenged with freshly egressed tachyzoites (ME49/PTG-GFP, MOI 1.5). One h post-challenge, cells were pharmacologically treated with picrotoxin (50 μM) and bicuculline (50 μM) or with bumetanide (40 μM) for 5 h. Further, treated cells were stained with CMTMR (1 μM) and non-treated cells were stained with CMF2HC (2 μM) for 30 min. Cells were washed and adoptively transferred into C57BL/6 mice. Each mouse received two consecutive i.p. injections, first with treated CMTMR-prelabeled cells (5 × 106) and second with non-treated CMF2HC-prelabeled cells (5 × 106). Mice were sacrificed 14–18 h post-infection to collect organs and cells from peritoneal lavages. The organs were triturated, filtered through 40-μm cell strainer and subjected to further analysis.

Flow cytometry

Cells collected from peritoneum and spleen, after RBC lysis and Fc receptor blockade with CD16/32 antibody (clone 2.4G2; eBioscience, San Diego, CA, USA) for 15–20 min, were stained with CD11c-PE-cyanine-7 antibody (clone N418; eBioscience) for 30–40 min as per providers recommendation. Following extensive washing, cell samples were run on a LSRFortessa flow cytometer (Beckman Coulter, Pasadena, CA) and data were analyzed using FlowJo software (FlowJo LLC).

Plaquing assays

Plaquing assays were performed as described (Fuks et al., 2012). Briefly, organs were extracted and homogenized on 70 μm cell strainers. The numbers of viable parasites per g of tissue were determined by plaque formation on HFF-1 monolayers.

Replication assay

mBMDCs were challenged with freshly egressed tachyzoites (ME49/PTG-GFP, 1 h, MOI 2). Cells were washed by centrifugation to remove extracellular parasites and seeded in 96-well plates. Cells were imaged by epifluorescence microscopy to detect number of parasites per vacuoles at 6 h and 24 h post-infection in presence of pharmacological treatments.

Data mining and statistical analyses

Motility plots were compiled using ImageJ with manual tracking and chemotaxis plugin. X- and y-axes in the plots show distances in μm. For motility assays, box-whisker and scattered dot plots represents median velocities (μm/min) with boxes marking 25th to 75th percentile and whiskers marking 10th and 90th percentiles of the datasets. Gray circles represent velocities from individual cells. Accumulated distances travelled by the cells (% cells tracked are binned at a range of 2 μm distance) are represented as histograms. Bar graphs show mean + SEM. Heat maps represent transcriptional changes in mRNA expression (2-ΔCt) upon challenge with T. gondii. Red and blue color scales indicate percentage increase and decrease in expression, respectively, normalized to expression in unchallenged cells at the same time point, respectively. Data mining and statistical analyses were performed using GraphPad Prism 7.0 (La Jolla, CA, USA). The statistical significance is represented as p<0.05 (*), p<0.01 (**), p<0.001 (***) or non-significant p≥0.05 (ns).

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This study demonstrated that GABA regulates the cellular motility of human and murine mononuclear phagocytes. The GABAergic components were present in the phagocytes, such as the metabolic enzymes and transporters, GABA-A receptors and regulators, and voltage-dependent calcium channels. Infection of phagocytes by coccidian parasites activated GABAergic signaling and increased cell motility. Gene silencing and pharmacological modulators of the functional components affected the GABAergic signaling and cell migration. The finding reveals a layer of regulatory role of neurotransmitter machinery in the host-pathogen interplay.

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Summary:

Herein authors demonstrate that GABA regulates the cellular motility of human and murine mononuclear phagocytes. The authors showed that the GABAergic components were present in the phagocytes, such as the metabolic enzymes and transporters, GABA-A receptors and regulators, and voltage-dependent calcium channels. Infection of phagocytes by coccidian parasites activated GABAergic signaling and increased cell motility. Gene silencing and pharmacological modulators of the functional components affected the GABAergic signaling and cell migration. The finding reveals a layer of regulatory role of neurotransmitter machinery in the host-pathogen interplay.

Essential revisions:

1) The study covered the axis from GABA synthesis to signaling, which was comprehensive but also inevitably affected the strength of the evidence on some of the steps. For example, additional and more direct evidence would help consolidate the involvement of the synthesis/production of GADA on the initial and late stages post infection. While the perturbation studies were mainly done with pharmacological modulators or RNAi knockdown, genetic ablation such as using the CRISPR approach in the cell line would strengthen the finding and the conclusion.

2) The authors claim in the Abstract that "This study reveals a hitherto unappreciated role for GABAergic signaling in the host-pathogen interplay between phagocytes and invasive coccidian parasites.". The same group has published two previous papers in PLoS Pathogens, both using murine and human phagocytes, studying this signaling pathway, with sometimes the same assays being performed (measurement of GABA after T. gondii infection, pharmacological inhibition of SNAP to block motility etc). The current study completes the picture of the various components of the GABA signaling pathway for parasite-induced motility, but does not discover a novel unappreciated role. The authors have to clearly state and relate all work performed here that is the same or similar to their previous publications. For example, how is the measurement of GABA different from their 2012 publication? Same goes for all inhibitors used in the study that are sometimes the same as in their 2017 paper. What is new here? Is it the details of the subunits found that are novel? Is it using human versus mouse phagocytes for the inhibitor studies? Please clarify in the Results section and in the Discussion.

For the in vivo results (impact of dissemination into brain and organs (spleen etc) but not peritoneum – why is this different to their 2017 publication, where no effect of dissemination into the brain was found (another part of the GABA pathway was inhibited then). This needs to be clarified and discussed.

3) It seems this study is the first time GABA signaling has an effect on T. gondii dissemination into the brain. Why was this only analyzed in the acute phase and not the chronic phase?

4) Figure 4, verification of protein levels using western blot or immunostaining and direct determination of GABA downstream signaling such as calcium would help clarify the functional defects of the GABA-A R upon knockdown. Figure 6K and L, western blot to verify the knockdown efficiency is desirable.

5) Figure 1G, a complete characterization of the transcript dynamics from 0-24h during infection would be informative. It seems that the cells of human origin changed more drastically post-infection. Since the cellular motility increased within minutes post-infection, a more detailed detection of GABA in Figure 1H at different time points could also clarify the effect on GABA production post-infection. Also, the regulation on GABA releasing or transportation at the initial phase might be more relevant given the rapid response of cell motility to infection. The expression levels of those components might affect the interpretation of the contribution of GABA on cellular motility at different phases of post-infection.

6) In the proposed model (Figure 10), NKCC1 mediated influx of chloride, which became the substrate of GABA-A R, and regulated GABA signaling. By pharmacological inhibition and gene silencing of NKCC1, the results in Figure 6 suggested that NKCC1 played a role in regulating hypermotility of parasitized phagocytes. However, there were no experiments indicating that NKCC1 functioned through regulating activities of GABA-A R. The authors should establish the connections between NKCC1 and GABA-A R in parasitized phagocytes.

7) To demonstrate that calcium channels were the downstream mediator of GABA signaling, the author showed that perfusion of GABA generated transient cytosolic Ca2+ elevations (Figure 8A). The calcium responses in phagocytes infected by coccidian parasites should be measured to support that calcium influx did happen in parasitized phagocytes. In addition, the calcium influx of parasitized phagocytes treated with the GABA-A R blocker should also be shown.

8) SNAP was used as blockade of GABA transporters (Figure 3E). The concentration of GABA in the supernatant should be measured to support that SNAP did block the transport of GABA. The motility and velocities of unchallenged mBMDCs treated with SNAP and other modulators should be measured as negative controls (Figure 3E).

Essential revisions:

1) The study covered the axis from GABA synthesis to signaling, which was comprehensive but also inevitably affected the strength of the evidence on some of the steps. For example, additional and more direct evidence would help consolidate the involvement of the synthesis/production of GADA on the initial and late stages post infection. While the perturbation studies were mainly done with pharmacological modulators or RNAi knockdown, genetic ablation such as using the CRISPR approach in the cell line would strengthen the finding and the conclusion.

An important point related to the secretion of GABA at early and late time-points of infection is brought up. To address this, we have performed kinetic analyses of secreted GABA during 1-24 h. The data show an elevation of GABA in the supernatant shortly after infection and an increase over time, indicating that GABA production starts shortly after parasite invasion and accumulates during infection. The data reinforce the conclusions and is consistent with the observed maintained hypermotility of infected phagocytes over time (Results, subsection “Human and murine mononuclear phagocytes exhibit hypermotility and secrete GABA upon challenge with T. gondii and N. caninum”, new Figure 1I).

We have further strengthened the evidence for other steps of the GABAergic signaling cascade with new data, reinforcing the conclusions. These are specifically discussed under points 6, 7 and 8 below.

While genetic ablation, for example by CRISPR, in a cell line could strengthen the findings related to specific GABAergic determinants, the main goal of this paper was to describe the general role of GABAergic signaling in primary phagocytes. There is very little or no data on the expression and conservation of the GABAergic system in phagocytes. To this end, it was crucial to use primary cells from various sources and cells from relevant hosts for coccidian infections. Also, the hypermigratory phenotype is weaker or even absent in cell lines which would have complicated the phenotypic analyses and comparisons with primary phagocytes. This likely relates to the finding that, while primary human and murine phagocytes consistently express GABAergic components, we have observed that cell lines of various sources may have altered expression or undetectable expression of specific GABAergic components (our unpublished observations and see also Figure 2—figure supplement 1). On the technical note, we have performed extensive controls in cell lines and primary cells (see also Kanatani et al., 2017). In this study, we target GABAergic signaling in human and mouse phagocytes at 5 different levels in the GABAergic signaling cascade. We use pharmacological inhibitors as guidance for more specific targeting of genes by shRNA. The number of identified targets yielding an abolished migratory phenotype together with reconstitution experiments undoubtedly show that GABAergic signaling drives the hypermigratory phenotype.

2) The authors claim in the Abstract that "This study reveals a hitherto unappreciated role for GABAergic signaling in the host-pathogen interplay between phagocytes and invasive coccidian parasites.". The same group has published two previous papers in PLoS Pathogens, both using murine and human phagocytes, studying this signaling pathway, with sometimes the same assays being performed (measurement of GABA after T. gondii infection, pharmacological inhibition of SNAP to block motility etc). The current study completes the picture of the various components of the GABA signaling pathway for parasite-induced motility, but does not discover a novel unappreciated role. The authors have to clearly state and relate all work performed here that is the same or similar to their previous publications. For example, how is the measurement of GABA different from their 2012 publication? Same goes for all inhibitors used in the study that are sometimes the same as in their 2017 paper. What is new here? Is it the details of the subunits found that are novel? Is it using human versus mouse phagocytes for the inhibitor studies? Please clarify in the Results section and in the Discussion.

For the in vivo results (impact of dissemination into brain and organs (spleen etc) but not peritoneum – why is this different to their 2017 publication, where no effect of dissemination into the brain was found (another part of the GABA pathway was inhibited then). This needs to be clarified and discussed.

We agree that this needed additional clarification. GABAergic signaling encompasses a complex signaling cascade with multiple components. Our previous work has described for T. gondii infection that (1) mBMDCs and MoDCs secrete GABA, (2) that mBMDCs express some GABAergic components, (3) that pharmacological treatments targeting GABA inhibit hypermotility in mBMDCs and (4) gene silencing of CaV1.3 in BMDCs abolishes hypermotility.

In the current manuscript, we (1) identify and functionally assess the GABA receptor subunits (by selective pharmacological targeting and shRNA) that likely constitute the functional GABA-A receptors, (2) we identify GAD67 the principal GABA synthesis enzyme in phagocytes, (3) we identify NKCC1 as a main regulator of GABAergic signaling in phagocytes. Further, we extend previous findings (which were mostly on murine DCs) to (4) elutriated human myeloid DCs, and monocytes for the first time and (5) extend the concept of GABAergic activation beyond T. gondii to N. caninum, using 5 separate strains of the two coccidia. We have clarified and highlighted these aspects in the revised discussion.

No single experiment overlaps with our previous publications but sometimes the same pharmacological inhibitors are used together with additional inhibitors and agonists. We have previously identified CaV1.3 as the mediator of calcium fluxes in murine BMDCs (Kanatani et al., 2017). This could have been left out but we reasoned that VDCCs are an important constitutive component of the GABAergic system and therefore, this had to be addressed in a comprehensive approach. However, in the current paper hMoDCs are utilized when studying CaV1.3, which significantly reinforce and extend our previous conclusions in murine cells to human primary cells. There is no experimental overlap.

Our previous work is indicated in the Introduction with references, we have further clarified its reference in the Results section, and bring it up in the Discussion with a summary of the findings in Figure 10. We think this should make it clearer to the reader how the novel findings relate to the two previous papers. We have also provided precision to the above indicated sentence in the Abstract: “The findings reveal a regulatory role for a GABAergic signaling machinery in the host-pathogen interplay between phagocytes and invasive coccidian parasites”.

For the in vivo results (Figure 9), this paper primarily addresses the systemic dissemination of T. gondii rather than the passage to the brain. By a novel combination of approaches (competition assay, see our publ. Lambert et al., 2009, and adoptive transfer of shRNA/pharmacologically treated cells), we show that both targeting -here identified- GABA-A R subunits and -here identified- NKCC1 hampers parasite dissemination. The data is in line with our previous data of pharmacological inhibition of GABA synthesis and transportation (Fuks, et al., 2012, not addressed here in vivo). In Kanatani, et al., 2017, pharmacological inhibition of voltage-gated calcium channels was applied (not addressed here in vivo), with an impact on systemic dissemination (blood, MLN, spleen) but non-significant effects on parasite loads in the brain. Thus, different targets and in vivo methodologies are used in each paper. However, the approaches jointly indicate that targeting (1) GABA synthesis/transportation, (2) GABA-A R signaling, (3) GABA-A receptor regulation / NKCC1 or (4) VDCC signaling hamper T. gondii systemic dissemination. Approaches (1-3) show an impact on parasite loads in the brain while (4) non-significantly impacts parasite loads in the brain. More important, in the current manuscript, we simultaneously adoptively transferred treated and untreated cells and therefore the untreated cells serve as an internal control for each mouse. These aspects have now been further clarified in the revised Discussion.

3) It seems this study is the first time GABA signaling has an effect on T. gondii dissemination into the brain. Why was this only analyzed in the acute phase and not the chronic phase?

The review raises an interesting question. The review is correct that in a previous paper (targeting VDCC signaling) non-significant differences were observed on parasite loads in the brain (Kanatani et al., 2017). In contrast, an effect on parasite loads in the brain was observed in Fuks et al., 2012. In that paper, we targeted GABA synthesis and transport using pharmacological inhibitors. In the current paper, we target 2 novel functions: GABA-A receptor function and the GABA regulator NKCC1 with an impact on parasite loads in the brain. This raises an important question: Do all GABAergic signaling components impact equally on parasite dissemination? While we believe that different components impact differently or to different extent, we have not tested this extensively. We have clarified this aspect in the revised Discussion, providing references and avoiding excessive speculation.

Regarding testing the role of GABAergic signaling in the model of chronic phase of infection, we have not tested this (yet) for various reasons. The total parasite load in the chronic phase is likely determined by several host and parasite factors, of which the initial invasion is one. The focus here is on the role of GABAergic signaling on the initial systemic dissemination of acute infection. Given the rapid lytic cycle of T. gondii, it is unlikely that the T. gondii parasites in the CNS during chronic infection relate directly to the initial phagocytes that mediate the initial systemic dissemination and blood transportation of the parasite. However, we think this important question should be addressed in the future, also in light of our recent findings that brain-resident phagocytes (microglia) respond to T. gondii infection with hypermigration via GABAergic signaling (Bhandage et al., 2019). This has to be indeed addressed in a different experimental setup which involves different parasite stages (bradyzoites) and different host cell models. Thus, we agree that assessing brain cysts in the chronic phase of infection could add one additional characterization but it would not change the overall conclusions of the paper. Because an effect was observed in the acute phase, we opted not to pursue putative effects in the chronic phase, in line with approved animal ethics protocols. We have highlighted these aspects in the revised Discussion.

4) Figure 4, verification of protein levels using western blot or immunostaining and direct determination of GABA downstream signaling such as calcium would help clarify the functional defects of the GABA-A R upon knockdown. Figure 6K and L, western blot to verify the knockdown efficiency is desirable.

Whenever possible, a verification has been performed in the manuscript, for example by quantification of GABA upon silencing of the synthesis enzyme GAD67 (Figure 5I) or transporter inhibition by SNAP (Figure 5F). However, for other targets, specifically GABA-R subunits, quantifications of knockdown at the protein level have proven to be challenging. We have extensively attempted to quantify expression and knockdown by western blotting and immunofluorescence for GABA-A R subunits and NKCC1. While detection of subunits is feasible in highly-expressing neuronal tissue, weaker bands or no polypeptide bands are observed using purified primary phagocytes. Because expression in phagocytes is likely generally lower compared with neurons, reaching sufficient high numbers of transduced primary phagocytes (which would in theory allow quantitative analyses) is difficult. We have searched the literature for reference, but found that even in neuronal models targeting NKCC1 by shRNA, quantifications are not provided at the protein level (Mejia-Gervacio et al., Neural development, 2011), presumably due to the difficulty to quantify NKCC1 by Western blotting (MW 160-200 kDa, with several splice variants). Moreover, the currently available set of commercial antibodies for GABA-A R subunits yielded inconsistent detection and thus reliable quantifications were not possible. Examples of blots are provided in Author response image 1. The added data (new Figure 8H-I-J) also show the effects of pharmacological stimulation and inhibition of GABA-A receptors at the single cell level and that calcium influx is activated and inhibited, respectively, via GABA-A receptors.

Author response image 1.

Representative Western blots show detection of weak or undetectable polypeptide bands for GABA-A R subunits (alpha3, alpha5, rho1) in mBMDCs with commercially available antibodies.

Cell sorting can be used to enrich for transduced populations but was not an option in our case. Here, we were ultimately interested in the functional phenotypic analysis (motility) of transduced cells that had been invaded by Toxoplasma. First, we opted to use cell sorting as a way of enriching for transduced primary cells but our experience is that the harsh flow conditions of cell sorting had a dual effect: (1) Increased cell death/lysis with decreased parasite viability and (2) Stressing the cells with an impact on their migratory behavior making it difficult to control for effects by Toxoplasma infection and treatments. We therefore realized that avoiding this stress moment, while making quantifications of knock-down slightly less precise (likely underestimating knockdown), would be a better reflection of functional gene silencing levels in cells that were phenotypically characterized in motility assays. This did not affect functional motility counts as only transduced cells (positive for the reporter GFP+, Figure 4—figure supplement 1A-D) were assessed in the motility assays.

We validated the shRNA constructs and approach in cells known to express GABAergic signaling components (neuronal murine NE4C and human SH-SY5Y) shown in Figure 4—figure supplement 1A-E. We have clarified this in the manuscript (Results).

5) Figure 1G, a complete characterization of the transcript dynamics from 0-24h during infection would be informative. It seems that the cells of human origin changed more drastically post-infection. Since the cellular motility increased within minutes post-infection, a more detailed detection of GABA in Figure 1H at different time points could also clarify the effect on GABA production post-infection. Also, the regulation on GABA releasing or transportation at the initial phase might be more relevant given the rapid response of cell motility to infection. The expression levels of those components might affect the interpretation of the contribution of GABA on cellular motility at different phases of post-infection.

We have expanded our analysis to include additional time points for hMoDCs and Monocytes and these are provided in the new Figure 1H (for GABA synthesis and catabolism enzymes) and new Figure 5D (for GABA transporters).

Additionally, more detailed kinetics (1-3-6-16-24 h) of GABA secretion by hMoDCs from three human donors are provided in the new Figure 1I. The data show an increase of GABA in the supernatant shortly after infection and an increase over time, indicating that GABA production starts shortly after parasite invasion and production/secretion is maintained during infection. Jointly, the data reinforce the conclusions and is consistent with the observed maintained hypermotility of infected phagocytes over time (Results, new Figure 1I).

6) In the proposed model (Figure 10), NKCC1 mediated influx of chloride, which became the substrate of GABA-A R, and regulated GABA signaling. By pharmacological inhibition and gene silencing of NKCC1, the results in Figure 6 suggested that NKCC1 played a role in regulating hypermotility of parasitized phagocytes. However, there were no experiments indicating that NKCC1 functioned through regulating activities of GABA-A R. The authors should establish the connections between NKCC1 and GABA-A R in parasitized phagocytes.

The review raises an important question related to the role of NKCC1 in GABA-A R function and hypermotility. While well-characterized in neuronal cells, the expression of NKCC1 has remained uncharacterized in leukocytes and the present data represent the first evidence of expression by phagocytes, to our knowledge.

To address a direct connection between NKCC1 function and GABA-R we have now performed reconstitution experiments in NKCC1-inhibited cells (bumetanide) using GABA R activation by GABA and muscimol. Contrasting with results on cells with abrogated GABA synthesis (SC; shown in Figure 5G), the results show that GABA or the GABA A receptor agonist muscimol do not reconstitute hypermotility in NKCC1-inhibited cells (new Figure 6O). To confirm this, we have performed stimulation experiments with GABA in shNKCC1 and shGAD67-transduced cells. The data show that upon silencing of GABA production (shGAD67), addition of exogenous GABA reconstitutes hypermotility. In contrast, upon silencing of NKCC1, GABA fails to reconstitute hypermotility (new Figure 6P and Q, Figure 6—figure supplement 1).

Jointly, the data indicate that NKCC1 is needed for optimal GABA-A R function mBMDCs and hMoDCs, reinforcing the signaling model (Figure 10) and in line with regulatory functions on GABA signaling attributed to NKCCs/KCCs in neurons (Bortone and Polleux, 2009, Kaila et al., 2014). The data indicates a connection between NKCC1 and GABA-A R function in parasitized phagocytes (Results, new Figure 6O, P, Q) and we now bring up this aspect in the Discussion.

7) To demonstrate that calcium channels were the downstream mediator of GABA signaling, the author showed that perfusion of GABA generated transient cytosolic Ca2+ elevations (Figure 8A). The calcium responses in phagocytes infected by coccidian parasites should be measured to support that calcium influx did happen in parasitized phagocytes. In addition, the calcium influx of parasitized phagocytes treated with the GABA-A R blocker should also be shown.

We have added this data. Indeed, parasitized DCs respond with calcium fluxes to GABA-stimulation in a GABA-A R dependent manner. Moreover, perfusion of SCS (that inhibits β-subunit-containing GABA-A Rs) or picrotoxin (broad inhibitor that blocks all known subtypes of GABA-A Rs) yielded decreased or abolished calcium fluxes in parasitized cells, reinforcing our conclusions on GABA-A Rs and VDCCs. The data have been added to Figure 8H, I, J (Results).

8) SNAP was used as blockade of GABA transporters (Figure 3E). The concentration of GABA in the supernatant should be measured to support that SNAP did block the transport of GABA. The motility and velocities of unchallenged mBMDCs treated with SNAP and other modulators should be measured as negative controls (Figure 3E).

We have performed the suggested experiments with SNAP. The data show that in presence of SNAP, at concentrations applied in neuronal cells, the amount of GABA in the supernatant is consistently reduced by ∼ 60% even after 24 h, supporting the notion that SNAP blocks the transport of GABA. This reinforces the conclusions on the role of GATs. It also provides indications on the involved GABA transporters as SNAP has different affinity for different GATs. SNAP has been shown to have highest selectivity for GAT-2 and GAT-3 and given IC50 values are 5, 21 and 388 μM for hGAT-3, rGAT-2 and hGAT-1, respectively. Note also that, in mice, GAT2 corresponds to A12/BGT1, GAT3 corresponds to A13/GAT2 and GAT4 corresponds to A11/GAT3, while GAT1 carries the same name as in humans and rats (A1) (Nelson, 1998; Cohen-Kfir, 2005). Thus, a complete inhibition of GABA transportation by SNAP is not expected. The data has been added to the new Figure 5F (Results).

Data including velocities of all negative controls has now been added in the new Figure 3E and F. The data clarify and reinforce the conclusions (Results).