Toxoplasma gondii is a parasite that commonly infects most warm-blooded animals and is thought to affect over two billion people worldwide. In most cases, the infection does not cause any symptoms, although it can lead to serious complications in pregnant women or people with a weakened immune system.

T. gondii has a complex life cycle that involves different stages. During infection, the parasite invades the host cells and replicates inside a specialized cell structure called a ‘parasitophorous vacuole’ until the host cell bursts. The parasite then spreads and infects more host cells. The replication is synchronised, meaning all parasites in a host cell replicate at the same time. It was unclear how the parasites coordinated this process, but some researchers suggested that the parasites remained connected to each other to communicate by exchanging material and information. A good candidate to form such connections is the protein actin, which in many organisms forms filaments that guide the transport of cargo molecules in the cell. However, previous research indicated that actin in T. gondii is incapable of forming these stable filaments.

Periz et al. developed a new tool of fluorescence markers that specifically bind to actin in T. gondii and found extensive actin networks that connected parasites with each other and also to the membrane of the parasitophorous vacuole. Actin was needed to transport molecules between the parasites within a vacuole and was also found to enter the cells of the parasite. When the protein was depleted in the parasite, the network collapsed; the parasites started to replicate at different times and could no longer leave the host cell.

A next step will be to further investigate the role of actin in T. gondii and other parasites using the tools developed by Periz et al. A better understanding of replication of T. gondii could provide clues to new treatments for parasitic diseases that cause substantial economic losses worldwide.

Toxoplasma gondii is a wide-spread obligate intracellular parasite that is thought to infect over two billion people worldwide. T. gondii infection of healthy individuals causes no major complications, infection can cause severe disease in immunocompromised individuals and foetuses infected in utero. The pathological manifestation is caused in a large part by repeated rounds of the parasite’s lytic cycle, beginning with active invasion of the host cell by the parasite, replication within a specialized vacuole termed the parasitophorous vacuole (PV), followed by egress and lysis of the host cell. Replication occurs via a unique process of endodyogeny, where two daughter parasites are constructed within the mother, before elongating and budding, leading to the breakdown of the maternal parasite (Francia and Striepen, 2014). Remnants of the mother cell remain at the posterior end of the daughter parasites in a structure known as the residual body (RB), which has a role in organizing the parasites into their characteristic rosette pattern within the PV (Muñiz-Hernández et al., 2011; Hu et al., 2002). Membrane connections persist between the RB and the parasites until host cell lysis and parasite egress, presumably to allow inter-parasite communication (Muñiz-Hernández et al., 2011). To date little is known about the molecular mechanisms underlying these processes. Previous studies focused on the role of microtubules during parasite replication, since treatment with microtubule inhibitors leads to structural collapse of the parasite, while actin-modulating drugs are thought to cause only slight defects in replication (Shaw et al., 2000). In contrast, research on parasite actin focused on its crucial role during host cell invasion and egress (Dobrowolski and Sibley, 1996). However, recent studies have also implicated actin and myosins in intracellular processes including apicoplast division (Egarter et al., 2014; Andenmatten et al., 2013; Jacot et al., 2013), secretory organelle (dense granule) transport (Heaslip et al., 2016) and parasite replication (Haase et al., 2015).

Actin is a highly conserved protein, which forms dynamic filaments in eukaryotic cells. Through an association with actin-binding proteins, these filaments are themselves organized into higher order structures that play important roles in a wide variety of cellular functions including muscle contraction, vesicle transport and cytokinesis. T. gondii actin is encoded by a single gene, act1 and has only ~80% sequence identity with mammalian actin isoforms but shares 93% similarity with Plasmodium ACT1 (Dobrowolski et al., 1997). Apicomplexan ACT1 is clearly essential, and compared to its counterparts in higher eukaryotes is believed to be intrinsically unstable, resulting in the formation of only short filaments (Skillman et al., 2011). Biochemical assays indicate that 97% of the parasites actin is present in the globular form (Dobrowolski et al., 1997; Skillman et al., 2011; Wetzel et al., 2003). It has been proposed that apicomplexan actin is unique amongst actins as it polymerizes in a highly unusual, isodesmic manner (Skillman et al., 2013). According to the isodesmic polymerisation model, monomer addition is governed by a single equilibrium constant, meaning that no (unfavourable) activation step is required to initiate the formation of the first dimer leading to polymerisation. In this instance, nucleation and elongation are equally favourable. This contrasts to cooperative polymerisation, where the activation step is the formation of the first dimer/trimer, which has a higher equilibrium constant than polymer elongation (Smulders et al., 2010). Therefore polymer formation can only occur above a critical concentration (Cc) of monomers (Pantaloni et al., 1985). It is this activation step that is regulated by actin nucleators, such as the Arp2/3 complex or formins (Carlier et al., 2015). Puzzlingly, formins and other actin nucleating proteins have been shown to have essential roles in Toxoplasma and Plasmodium, begging the question of their function if they are not required to initiate actin polymerisation or accelerate filament elongation (Baum et al., 2008; Jacot et al., 2013). A recent study suggested that the polymerization process of apicomplexan actin needs to be reinvestigated, as heterologously expressed apicomplexan actin, the basis for many of the previous studies, is incorrectly folded (Olshina et al., 2016). Furthermore, it was demonstrated that conditional deletion of act1 in T. gondii results in complete abrogation of known actin functions, long before G-actin levels are fully depleted, suggesting that in vivo the formation of F-actin depends on a critical monomer concentration (Whitelaw et al., 2017). This raises concerns about previous studies of actin polymerization kinetics based on mis-folded actin. Furthermore, imaging studies on different life cycle stages of the apicomplexan parasite, Plasmodium falciparum, demonstrated the formation of an extensive F-actin cytoskeleton in both gametocytes and ookinetes (Hliscs et al., 2015; Siden-Kiamos et al., 2012). Using 3D-SIM it was demonstrated in gametocytes that these filaments appear to be organized in the cytoplasm, below the Inner Membrane Complex (IMC; a specialised structure found in apicomplexans that consists of membranous vesicles and structural components located just beneath the plasma membrane) of the parasite rather than between the IMC and the plasma membrane (Hliscs et al., 2015), where it is thought to power parasite gliding motility (Meissner et al., 2013).

To further address the role of ACT1 in parasite growth, we characterized a recently generated conditional actin knockout parasite line (act1 cKO) during the intracellular portion of the parasites lytic cycle (Andenmatten et al., 2013). We demonstrate that act1 cKO parasites lack a residual body and grow in an asynchronous and disorganized manner within the PV. Further investigation of actin functions required imaging the filamentous actin cytoskeleton. Previous attempts to visualise F-actin within the parasites have largely been unsuccessful, since conventional actin labelling techniques such as Life-Act, Phalloidin and SiR-Act do not allow detection of F-actin within the parasites. GFP-actin shows a high signal to noise ratio in parasites causing the inability to differentiate actin filaments from the monomeric form Angrisano et al. (2012a). Thus we sought a new approach to image F-actin in intracellular parasites by expressing actin-chromobody, a single chain anti-actin camel antibody that has been successfully employed in diverse eukaryotes, including plants (Rocchetti et al., 2014) and animal cells (Plessner et al., 2015; Panza et al., 2015). Chromobodies (Cb) were found to have several advantages compared to other actin probes, such as lower toxicity, less influence on F-actin dynamics and a high signal to noise ratio (Panza et al., 2015). Using this probe, we were able to visualise filamentous actin in live parasites. While some filamentous structures were seen within the parasite cytosol as expected, we were surprised to observe extensive F-actin networks in the RB and F-actin-containing membranous tubules linking parasites within a vacuole. These tubules appear to be involved in the transport of material between parasites and recycling of the mother IMC at the end of the replication cycle. Collectively, these data identify new roles for actin in the intracellular phase of the parasites lytic cycle and provide a robust new tool for imaging parasitic F-actin dynamics.

Studies on Toxoplasma actin performed in the 90’s suggested that apicomplexan actin is highly divergent and incapable of forming stable filaments (Dobrowolski and Sibley, 1997; Dobrowolski et al., 1997; Dobrowolski and Sibley, 1996). Since then, polymerization kinetics of heterologously expressed apicomplexan actin showed that it polymerized in an unusual isodesmic process, in contrast to conventional eukaryotic actin (Skillman et al., 2013). These findings have been recently questioned, as heterologously expressed apicomplexan actin was shown to be misfolded (Olshina et al., 2016). In parallel, a number of recent genetic studies have demonstrated important functions of parasite actin during intracellular development, including maintenance of the apicoplast (Andenmatten et al., 2013; Egarter et al., 2014), daughter cell replication (Haase et al., 2015) and motility of secretory organelles (Heaslip et al., 2016). These findings cannot be easily reconciled with the current view of parasite actin being incapable of efficient polymerization, and would instead predict the presence of F-actin filaments within the parasite cytosol. Furthermore, there are 11 putative myosin motors within the parasite (Foth et al., 2006), driving diverse cellular processes from cell division to motility, which require actin filaments to function.

A major impediment to resolving this controversy has been the lack of appropriate reagents for specifically labelling F-actin. Parasite actin does not bind phalloidin, the gold standard reagent in other eukaryotic systems (Skillman et al., 2011) and, attempts to use genetically encoded actin sensors, such a LifeAct (Riedl et al., 2008) or Utrophin-CH (Burkel et al., 2007) have failed thus far (our and others unpublished data). Using a novel tool based on camelid nanobodies, Chromobody, we have visualized F-actin in T. gondii for the first time in live parasites and demonstrated it has important and previously unforeseen roles during the intracellular development of the parasite. While individual actin filament kinetics could not be easily defined, due to their highly dynamic nature within the parasite cytosol (Videos 5–7), a relatively stable F-actin network was clearly visible that linked individual parasites within the vacuole. Chromobodies have been used successfully in diverse eukaryotes to analyse actin dynamics (Panza et al., 2015; Plessner et al., 2015; Rocchetti et al., 2014) and appear to have less toxicity than other F-actin sensors. However, we were concerned that Cb expression in T. gondii would perturb actin polymerisation, especially given the proposed dynamic and unstable nature of T. gondii actin. Indeed, high levels of transiently expressed Cb inhibited dense granule motility, likely by stabilizing F-actin (Figure 3C). However, upon stable expression of Cb-Halo or Cb-Emerald, no effect on the actin-dependent processes of invasion, replication and egress were observed, while only small alterations in dense granule movement and parasite motility were detectable (Figure 4). The reason for these discrepancies is currently unknown, but may be related to differing sensitivities of actin dynamics for these processes. Additionally, we demonstrated that the total amount of F-actin in the parasite does not change upon expression of Cb (Figure 5), indicating that while actin dynamics may be modulated, the overall polymerization state of actin does not change. We show that parasite actin remains susceptible to depolymerising and stabilizing agents Cyt-D and Jas. Moreover, the localization of Cb to filamentous structures in the residual body and inter-parasite network critically depends on the presence of actin (Figure 6). These data confirm that genetically encoded Cb-Halo and Cb-Emerald does not significantly alter actin dynamics within the parasite and so is an appropriate and robust tool for examining F-actin in this system.

The presence of actin in the RB and inter-parasite tubules has not previously been observed or predicted, and so we sought to independently verify that this localization was not an artefact of Cb expression. Labelling parasites with α-actin that preferentially recognises F-actin (Angrisano et al., 2012b) allowed the detection of actin positive structures connecting individual parasites (Figure 6), though the stain appeared more diffuse. When actin filaments were stabilised by the presence of Jas (Figure 6) this stain becomes more prominent and this was not dependent on the expression of Cb. This suggests that F-actin is less accessible to exogenously added antibodies after fixation, perhaps due to epitope alterations during fixation or the association of actin binding proteins to filaments. Interestingly, while actin has never been previously associated with this membranous network in T. gondii, it appears very similar to structures recently described for the related parasite Theileria annulata, which was shown to contain F-actin in a similar configuration (Kühni-Boghenbor et al., 2012).

Using a combination of imaging and reverse genetics, we demonstrate that F-actin is required for the formation and/or maintenance of the residual body. Parasites lacking actin do not contain a residual body and are disorganized within the PV. In addition, we show that these extracellular actin-containing structures are required for the transport of material between parasites within a vacuole. The residual body and inter-parasite network allow both the free diffusion of cytoplasmic material between parasites, and transport of membrane bound vesicles between parasites. Further work will be required to determine if these membrane bound vesicles are then taken up by other parasites or are simply transported to the RB to remove them and if a myosin motor powers this for its transportation, as is the case of dense granule transport within the parasite cytosol (Heaslip et al., 2016).

The organization of F-actin within the parasites and vacuole is highly dynamic throughout the cell cycle (Figures 8 and 9). In interphase parasites, long inter-parasite contentions extend throughout the PV. At the beginning of the replication cycle this network collapses and is found concentrated in the residual body (or bodies). Within the parasites, F-actin is found concentrated at the IMC of growing daughter cells during elongation (Figures 8 and 9). As the daughters bud from the mother cell, the inter-parasite connections again extend throughout the PV. As would be expected, the network also collapses in response to calcium ionophore, freeing the parasites and allowing egress. As motile parasites leave the PV, actin appears concentrated at the basal end of parasites (Figures 8 and 9).

Figure 9

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While Cb expression allowed us to demonstrate F-actin-containing membranous tubules for the first time, we were unable to assess the length of the individual F-actin filaments using these techniques. We speculate that this network consists of either short F-actin bundles that are cross-linked via unknown actin-binding proteins (based on the formation of short actin filaments in vitro) (Skillman et al., 2011) or that the presence of actin binding proteins such as formin, profilin, and coronin may coordinate their activities in vivo to produce longer actin filaments than those formed in vitro (Skillman et al., 2011; Olshina et al., 2016;Salamun et al., 2014). While this study focuses on the characterization of the membranous network within the PV, it is worth nothing that highly dynamic actin filaments have been also detected within the cytosol of the parasite, and we show that these dynamics are almost completely abolished upon depletion of ADF in good agreement with previous findings (Haase et al., 2015; Mehta and Sibley, 2011) (Figure 6; Videos 5–7). Defining the organization of cytosolic filaments will be critical to further elucidating the role of F-actin both during intracellular processes such as vesicle transport and extracellular processes such as motility and invasion, where the role of the parasites acto-myosin system is currently readdressed, since parasites devoid of detectable actin remain invasive (Whitelaw et al., 2017). Interestingly, it appears that host cell membrane dynamics, driven by host cell actin, appears to be modulated by the parasite, enabling invasion in the absence of a functional parasite acto-myosin system (Bichet et al., 2016).

It is possible that F-actin is organized into distinct, higher order structures, perhaps forming large stable bundles in the tubules while existing as single filaments or small bundles in the cytosol. Future work will be needed to further define the organization of the actin cytoskeleton and to identify the actin binding proteins which contribute to the formation of these structures.

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Author details

1. Javier Periz

   Wellcome Centre for Molecular Parasitology, Institute of Infection, Immunity & Inflammation, University of Glasgow, Glasgow, United Kingdom

   Contribution

   JP, Conceptualization, Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing

   Contributed equally with

   Jamie Whitelaw and Clare Harding

   Competing interests

   The authors declare that no competing interests exist.

2. Jamie Whitelaw

   Wellcome Centre for Molecular Parasitology, Institute of Infection, Immunity & Inflammation, University of Glasgow, Glasgow, United Kingdom

   Contribution

   JW, Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing—original draft

   Contributed equally with

   Javier Periz and Clare Harding

   Competing interests

   The authors declare that no competing interests exist.

3. Clare Harding

   Wellcome Centre for Molecular Parasitology, Institute of Infection, Immunity & Inflammation, University of Glasgow, Glasgow, United Kingdom

   Contribution

   CH, Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing

   Contributed equally with

   Javier Periz and Jamie Whitelaw

   Competing interests

   The authors declare that no competing interests exist.

4. Simon Gras

   Wellcome Centre for Molecular Parasitology, Institute of Infection, Immunity & Inflammation, University of Glasgow, Glasgow, United Kingdom

   Contribution

   SG, Investigation, Methodology

   Competing interests

   The authors declare that no competing interests exist.

5. Mario Igor Del Rosario Minina

   Wellcome Centre for Molecular Parasitology, Institute of Infection, Immunity & Inflammation, University of Glasgow, Glasgow, United Kingdom

   Contribution

   MIDRM, Data curation, Formal analysis, Investigation, Methodology

   Competing interests

   The authors declare that no competing interests exist.

6. Fernanda Latorre-Barragan

   Wellcome Centre for Molecular Parasitology, Institute of Infection, Immunity & Inflammation, University of Glasgow, Glasgow, United Kingdom

   Contribution

   FL-B, Formal analysis, Investigation, Methodology

   Competing interests

   The authors declare that no competing interests exist.

7. Leandro Lemgruber

   Wellcome Centre for Molecular Parasitology, Institute of Infection, Immunity & Inflammation, University of Glasgow, Glasgow, United Kingdom

   Contribution

   LL, Investigation, Visualization, Methodology, Writing—original draft

   Competing interests

   The authors declare that no competing interests exist.

8. Madita Alice Reimer

   Wellcome Centre for Molecular Parasitology, Institute of Infection, Immunity & Inflammation, University of Glasgow, Glasgow, United Kingdom

   Contribution

   MAR, Investigation, Methodology

   Competing interests

   The authors declare that no competing interests exist.

9. Robert Insall

   Cancer Research United Kingdom Beatson Institute, Bearsden, United Kingdom

   Contribution

   RI, Conceptualization, Methodology, Writing—original draft, Writing—review and editing

   Competing interests

   The authors declare that no competing interests exist.

10. Aoife Heaslip

    Department of Molecular Physiology and Biophysics, University of Vermont, Burlington, United States

    Present address

    Department of Molecular and Cell Biology, University of Connecticut, Storrs, United States

    Contribution

    AH, Conceptualization, Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing

    For correspondence

    aoife.heaslip@uconn.edu

    Competing interests

    The authors declare that no competing interests exist.

11. Markus Meissner

    Wellcome Centre for Molecular Parasitology, Institute of Infection, Immunity & Inflammation, University of Glasgow, Glasgow, United Kingdom

    Contribution

    MM, Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing

    For correspondence

    markus.meissner@glasgow.ac.uk

    Competing interests

    The authors declare that no competing interests exist.

    "This ORCID iD identifies the author of this article:" 0000-0002-4816-5221

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