The multiple functions of actin in apicomplexan parasites

The cytoskeletal protein actin is highly abundant and conserved in eukaryotic cells. It occurs in two different states‐ the globular (G‐actin) form, which can polymerise into the filamentous (F‐actin) form, fulfilling various critical functions including cytokinesis, cargo trafficking and cellular motility. In higher eukaryotes, there are several actin isoforms with nearly identical amino acid sequences. Despite the high level of amino acid identity, they display regulated expression patterns and unique non‐redundant roles. The number of actin isoforms together with conserved sequences may reflect the selective pressure exerted by scores of actin binding proteins (ABPs) in higher eukaryotes. In contrast, in many protozoans such as apicomplexan parasites which possess only a few ABPs, the regulatory control of actin and its multiple functions are still obscure. Here, we provide a summary of the regulation and biological functions of actin in higher eukaryotes and compare it with the current knowledge in apicomplexans. We discuss future experiments that will help us understand the multiple, critical roles of this fascinating system in apicomplexans.

acid identity, they display regulated expression patterns and unique non-redundant roles. The number of actin isoforms together with conserved sequences may reflect the selective pressure exerted by scores of actin binding proteins (ABPs) in higher eukaryotes. In contrast, in many protozoans such as apicomplexan parasites which possess only a few ABPs, the regulatory control of actin and its multiple functions are still obscure. Here, we provide a summary of the regulation and biological functions of actin in higher eukaryotes and compare it with the current knowledge in apicomplexans. We discuss future experiments that will help us understand the multiple, critical roles of this fascinating system in apicomplexans.

| ACTIN IN MOST EUKARYOTES
Actin is one of the most abundant proteins in eukaryotic cells and owing to its ability to polymerise into filaments (F-Actin) forming static or highly dynamic networks, plays an important function in many crucial cellular processes. The regulated dynamics of F-actin and crosslinking of individual filaments requires the integration of actin binding proteins (ABPs) with signalling cascades that ultimately regulate actin filament length, stability and anchorage (Hansen & Kwiatkowski, 2013). This control can be achieved at multiple levels, from direct binding of ABPs to post-translational modifications of actin and are based on influencing the basic polymerisation mechanism of this molecule (Dominguez & Holmes, 2011).
These arrangements of F-actin can result into different configurations and includes filaments orientated in the same direction, parallel and antiparallel bundles, and filaments forming actin lattices that enable it to fulfil different mechanical and functional roles (Chhabra & Higgs, 2007).
Actin polymerisation is a dynamic process that involves rapid changes in the assembly and disassembly of F-actin. One critical factor for the transition between G-actin and F-actin (actin treadmilling) is the hydrolysis of ATP (Korn, Carlier, & Pantaloni, 1987). The actin treadmilling cycle starts with the addition of G-actin-ATP to "barbed" (plus) end of the F-actin filament. During this polymerisation step, ATP is hydrolysed to ADP and inorganic phosphate (P i ). Hydrolysis of ATP results in stable F-actin filaments with bound ADP + P i (F-actin-ADP-P i ). Slow release of P i results in F-actin-ADP and destabilises the filament, resulting in the release of G-actin-ADP from the filament. In a growing F-actin strand, F-actin-ADP resides at the "pointed" (minus) end where the depolymerisation of G-actin-ADP occurs. Subsequently, ADP can be replaced with ATP in the monomer, generating a new G-actin-ATP molecule. The addition of a new G-actin-ATP molecule to the F-actin barbed end causes a conformational change in the adjacent actin molecule. Murakami and co-workers proposed this structural change to be responsible for the initiation of ATP hydrolysis (Murakami et al., 2010). Within subdomain 2 of the actin molecule lies the DNase I binding (D-) loop, which inserts itself into the hydrophobic cleft between subdomains 1 and 3 of the apposing monomer and is crucial for polymerisation (Fujii et al., 2010). Important differences in the amino acid residues of the D-loop between canonical and apicomplexan actins give rise to different filament polymerisation dynamics, which will be discussed in Section 2.2.
Of relevance is that the amount of G-actin has to be above a certain concentration threshold for polymerisation to occur. This threshold is referred to as the critical concentration, where the removal of G-actin occurs at the same rate as the addition of new monomers, but the net mass of the F-actin polymer does not change (Pollard & Borisy, 2003). Importantly, the critical concentration for polymerisation to happen at the barbed end is lower than the concentration needed for polymerisation at the pointed end or for the formation of an actin dimer (Pollard & Borisy, 2003;Pollard, 2016). This results in the typical behaviour of F-actin, where monomers at the barbed end are added 5-10 times faster than to the slow-growing pointed end.
Together, this cooperative polymerisation process results in the so called treadmillling of actin filaments (Figure 1a). It should also be mentioned that the critical concentration required for G-actin-ATP to polymerise was reported to be lower than the concentration required for G-actin-ADP (Cooke, 1975;Pollard, 1984). Consequently, actin polymerisation is more likely to occur at the barbed end of an F-actin strand by attaching G-actin-ATP monomers.
The first step of de novo F-actin filament formation is called actin nucleation, where a new F-actin filament is assembled from G-actin monomers (Pollard, Blanchoin, & Mullins, 2000). Inevitably, this process requires the formation of actin dimers and trimers, a process that is kinetically unfavourable (Pollard, 2016;Sept & McCammon, 2001).
From this point onwards, G-actin monomer assembly to the actin trimer occurs with the same rate as G-actin polymerisation on existing F-actin filaments (Sept & McCammon, 2001). The assembly of this polymerisation nucleus therefore represents the critical step that has to be overcome for de novo F-actin formation (Sept & McCammon, 2001). To efficiently control kinetically unfavourable steps of actin dynamics, a vast number of actin binding proteins (ABPs) has been described, some highly conserved in most eukaryotes and others unique to certain species (see [Pollard, 2016] and Table 1).
The formation of actin polymers is a tightly regulated process that involves several steps: Nucleation, polymerisation and regulation of the polymer size by actin treadmilling and filament stabilisation.
Nucleation of canonical actins depends on a set of ABPs to overcome the activation energy barrier of de novo filament formation. Three different types of actin nucleators have been described: the Arp2/3 complex, spire and the formin protein family (Goode & Eck, 2007). The Arp2/3 complex consists of seven subunits and promotes branching and formation of novel daughter F-actin filaments at an angle of 70 from an already existing filament (Mullins, Heuser, & Pollard, 1998). The Arp2/3 complex is an important nucleator of Factin in lamellipodia and is involved in cell migration (Suraneni et al., 2012). Spire, first discovered in Drosophila, possesses four WH2 domains used for attracting four G-actin monomers to create a nucleation complex, and can collaborate with formin to build essential Take Away • Disease-causing apicomplexan parasites such as Toxoplasma and Plasmodium possess significantly divergent actin genes compared to other eukaryotes. Differences in key amino acid residues contribute to important structural differences and actin filament instability in these parasites.
• Despite having only a basic set of actin binding proteins (ABPs), an extensive and dynamic formin-dependent Factin network has been visualised in live parasites using actin-binding nanobodies fused to fluorescent tags (chromobodies).
• Actin dynamics control central biological processes such as invasion, gliding motility, vesicular transport, and apicoplast inheritance in apicomplexan parasites. Genusspecific functions such as haemoglobin uptake and completion of cytokinesis has been additionally observed in Plasmodium falciparum.
• Use of actin-binding chromobodies combined with stateof-the-art reverse genetics and microscopy will be immensely useful in mechanistic dissection of the actin network and uncovering functions of known and novel ABPs.
The FH2 domain nucleates and elongates unbranched actin filaments by "processive capping" at the barbed end, while the FH1 domain can interact with the ABP profilin, release actin monomers sequested by profilin and incorporate them into the growing filament (Courtemanche, 2018). In apicomplexans, only formin like proteins have been identified and are thought to represent the only actin nucleators in these parasites (Tosetti, Dos Santos Pacheco, Soldati-Favre, & Jacot, 2019).
Actin treadmilling is the continuous removal of monomers from the pointed ends of filaments and their simultaneous incorporation at the barbed end, a process in which the ABPs formin, actin depolymerizing factor (ADF)/cofilin and profilin are involved.
ADF/cofilin family bind and destabilise F-actin filaments, thus increasing the amount of available G-actin monomers (Moon & Drubin, 1995;Nishida, Maekawa, & Sakai, 1984;Yonezawa, Nishida, & Sakai, 1985;Lappalainen & Drubin, 1997). Cofilin and ADF have higher affinity to G-Actin-ADP than to G-Actin-ATP, thereby increasing the depolymerisation rate of F-actin. While depolymerisation occurs at the pointed end during the treadmilling process, F-actin filament elongation takes place at the barbed end. One of the proteins involved in mediating filament assembly is the polymerisation factor profilin, which binds monomeric G-actin (Baum et al., 2006;Pollard, 2016;Carlsson, Nystrom, Sundkvist, Markey, & Lindberg, 1977;Pantaloni & Carlier, 1993). Profilin binds to G-actin-ATP and G-actin-ADP with similar affinity while drastically increasing the exchange rate of actin-bound ADP for ATP (Selden, Kinosian, Estes, & Gershman, 1999). Actin treadmilling occurs by formins assembling a pool of regenerated G-actin-ATP above the critical concentration at the barbed end of an actin polymer (Romero et al., 2004;Kovar, 2006). Cyclase associated protein (CAP) is essential for most eukaryotes and can work in synergy with ADF/cofilin to increase F-actin depolymerization by almost 100-fold, and furthermore, can exchange ADP on depolymerized monomers with ATP to enable another round of F-actin assembly (Kotila et al., 2019). Coronins have been described as a "double-edged sword," promoting F-actin disassembly in coordination with ADF/cofilin at ADP-rich pointed ends of networks, while promoting rapid F-actin growth at the ATP-rich barbed ends by recruiting the Arp2/3 complex for expansion of branches, thereby functioning during rapid actin-mediated processes auch as endocytosis and cell migration (Gandhi & Goode, 2008). Capping proteins (CPs) are a heretodimer composed of the α and β subunits, bind to barbed ends of F-actin and prevent the addition or removal of monomers, thereby stabilising the filament (Edwards et al., 2014;Pollard, 2016). Interestingly, they also stabilise short filaments produced by the actin related protein 1 (ARP1) (Cooper & Sept, 2008 (Baum et al., 2006;Dobrowolski, Niesman, & Sibley, 1997). There are notable differences in apicomplexan actin, in both the number of isoforms and conservation in amino acid sequence. While Toxoplasma (and most other apicomplexans) possesses only a single gene for actin , Plasmodium species possess two isoforms, act-1 and act-2 (Wesseling, Smits, & Schoenmakers, 1988). Plasmodium falciparum ACT1 (PfACT1) F I G U R E 1 Host cell invasion and intracellular F-actin flow. (a) F-actin polymer growth depends on a critical concentration of G-actin in the cytoplasm. The balance between incorporation of monomers during elongation and depolymerisation is controlled by multiple regulators. At the barbed end, profilin delivers ATP-rich monomers to formins for nucleation and polymer elongation; at the pointed end, ADF-1 binds ADP-rich actin and contributes to depolymerisation, polymer fragmentation and ATP hydrolysis. Nucleotide exchange on released ADP-actin to form ATPactin for a new round of polymerisation could be brought about by CAP and profilin. Stability of F-actin bundles is maintained by several actin binding proteins including CPs, coronins and formins. (b) The linear model for parasite motility proposes that a motor complex consisting of actin, myosins and associated proteins is located within the narrow space of 20 nm between the plasma membrane and the underneath saccule-like structure, the IMC. (c) Other alternative models can be proposed in which cytosolic myosins and IMC permeability contribute to transport of microneme adhesins and exchange of actin between the cytosol and space underneath the plasma membrane. (d) Cytoplasmic Actin and cell invasion. Initial attachment to the surface of a host cell depends on F-actin accumulation at the posterior pole and at the apical end. During invasion, the tight junction complex (TJ) is formed that stabilises the attachment of the parasite to the host cell. F-actin at the TJ is involved in force generation and parasite stability to facilitate nuclear entry. F-actin at the posterior end produces an actin lattice that provides plasticity and a "pushing" contraction force to allow nuclear deformation and entry through the narrow space of the tight junction. Deformation and contraction of the parasite region lacking the rigid structure of microtubules might facilitate nuclear entry by "pulling." We propose that the nucleus is squeezed through the TJ by a push-pull mechanism controlled by actin and potentially microtubules. (e) F-actin flow model in the parasitophorous vacuole during endodyogeny. There are three formins identified that have specific localisations: Formin-1 is localized at the apical end, formin-2 is closely associated to apicoplasts and formin-3 is at the parasite posterior end. Flow analysis combined with formin conditional knock-out studies suggest that F-actin flow is controlled by the formins. Formin-1 controls the apical posterior flow direction, formin-2 may be responsible for bidirectional flow of actin, and formin-3 may be responsible for controlling the flow in the RB and retrograde flow in the parasite. (f) Actin flow is involved in apicoplast inheritance in Apicomplexa. In Plasmodium, the actin network additionally regulates the transport of endocytosed haemoglobin to the food vacuole/fusion of vesicles, and the completion of cytokinesis is expressed in all life cycle stages, while ACT2 was found to be solely expressed in the sexual stages (Wesseling et al., 1989). Toxoplasma gondii actin (TgACT1) shares 93% amino acid sequence identity with PfACT1.

| Dynamics of apicomplexan Actin in vitro
While in most eukaryotes F-actin can form long filaments in vitro, apicomplexan actin forms only short filaments of less then 100 nm in the absence of filament-stabilising drugs such as jasplakinolide (Pospich et al., 2017;Schmitz et al., 2005). Apicomplexan actin has also been notoriously hard to visualise and characterize, both in vitro and in vivo, leading to conflicting interpretations regarding polymerisation mechanisms and functions. Early studies by Dobrowolski and colleagues used ultra centrifugation methods to investigate the state of the actin polymer ; the findings failed to detect filaments, leading the authors to propose that actin is mainly in the monomeric G-actin state in Toxoplasma. A comparative study with recombinant actin in vitro showed that Toxoplasma actin formed short unstable filaments (Sahoo, Beatty, Heuser, Sept, & Sibley, 2006).
Intriguingly, the critical concentration required for actin polymerisation in Toxoplasma was suggested to be lower compared to conventional actins, while F-actin assembly and turnover was suggested to occur very rapidly. It was proposed that amino acid residues on the Toxoplasma actin monomer surface differ from conventional actins and these differences contribute to filament instability, which could be an adaptation that enables fast parasite motility (Sahoo et al., 2006;Skillman et al., 2011). These results were supported by findings in Plasmodium falciparum (Schmitz et al., 2005), where short filaments ($100 nm) were observed in vitro when compared to rabbit actin ($350 nm). Further experiments using actin sedimentation assays led Skillman and co-workers to speculate on an isodesmic model for polymerisation in apicomplexan parasites, which would be a unique and surprising mechanism, only found in apicomplexans (Skillman et al., 2013 were responsible for filament instability (Pospich et al., 2017). Further crystallography studies of PfACT1 identified the Arg178/ Asp180-containing A-loop to be one of the factors responsible, which acts as a switch governing the relative stability of F-actin (Kumpula, Lopez, Tajedin, Han, & Kursula, 2019). Finally, a recent study used actin chromobodies and TIRF microscopy to visualise dynamics of PfACT1 (Lu, Fagnant, & Trybus, 2019) and a significantly higher critical concentration was determined. Here, the instability of F-actin was attributed to rapid filament shrinkage at the pointed end (Lu et al., 2019). The PfACT1 D-loop has important differences in amino acid residues compared to canonical actins, which contributes to natural filament instability essential for the parasite (Lu et al., 2019). A chimeric P. berghei actin-1 with a "canonical" Dloop could produce long filaments in vitro and restore gametocytogenesis in parasites lacking actin-2, indicating that the differential functional needs of the two actins rely heavily on differential filament stability (Vahokoski et al., 2014). In another study, a single point mutation N41H within the PfACT1 D-loop allowed PfACT1 incorporation into mammalian F-actin in a skin cell line (Douglas et al., 2018). Taken together, data obtained from recent in vitro experiments demonstrate that apicomplexan F-actin is more unstable than canonical actins. Differences are found in key residues in multiple regions of the G-actin monomers that appear to critically contribute to this phenomenon. Nonetheless, while different critical concentrations have been determined, a cooperative nucleation-elongation mechanism seems to be in place for F-actin polymerisation in apicomplexan parasites, as seen in all other eukaryotes studied thus far.
Interestingly, Theileria annulata parasites possess an actin isoform that has retained the amino acid residues Ser200, Met270 normally seen in canonical actins, which are mutated to Gly200, Lys270 in Plasmodium and Toxoplasma. Perhaps as a consequence, Theileria parasites make more stable F-actin structures, which were detected by cryoelectron tomography (Kuhni-Boghenbor et al., 2012). Indeed, TgACT1 produced more stable filaments with the reverse mutations Gly200Ser and Lys270Met (Skillman et al., 2011).

| Apicomplexan Actin function and distribution in vivo
Until recently it was believed that actin is required primarily for parasite motility. This assumption was based on early inhibitor studies which suggested that microtubules, but not actin, are required for parasite replication (Shaw, He, Roos, & Tilney, 2000) and conversely, actin is required for motility and host cell invasion (Dobrowolski & Sibley, 1996). In the case of Plasmodium, a potential role for PfACT1 in haemoglobin uptake was suggested using inhibitors of actin dynamics (Smythe, Joiner, & Hoppe, 2008). In this study it was 2.4 | The role of F-Actin during gliding motility and invasion of the host cell Apicomplexan gliding motility and host cell invasion was believed to be a purely parasite actin-driven process (Dobrowolski & Sibley, 1996). According to the linear motor model (Figure 1b), short F-actin filaments are polymerised between the plasma membrane (PM) and the inner membrane complex (IMC, a specialised structure found in apicomplexan parasites that consists of membranous cisternae and structural components located 20-30 nm beneath the PM).
These short filaments interact with transmembrane proteins derived from secretion of micronemes (invasion related apical organelles) via the glideosome associated connector (GAC) and the myosin A motor complex that is anchored within the IMC (Jacot et al., 2016). Furthermore, it is believed that during gliding motility and invasion, F-actin is formed at the apical tip of the parasite, where formin-1 is localized (Baum et al., 2008;Jacot et al., 2016). While this model has been supported by several lines of evidence, it cannot explain recent findings that used reverse genetic and biophysical approaches to determine force production and transmission during gliding motility and invasion: • Motility and invasiveness of conditional mutants for core compo-  , raising the question whether the acto-myosin system is required as a molecular clutch in order to initiate motility and to transmit the force generated by retrograde membrane flow or other mechanisms (Bretscher, 2014;Whitelaw et al., 2017). Indeed, a recent study used a combination of traction force microscopy, quantitative RICM (reflection interference contrast microscopy), micropatterning and expansion microscopy to determine the forces and mechanisms involved in parasite gliding. Together, the data suggest a mechanism, where the MyoA motor directs the traction force, allowing transient energy storage by the subpellicular microtubule cytoskeleton and therefore sets the thrust force required for gliding (Pavlou et al., 2020). Interestingly, using expansion microscopy, it was also demonstrated that MyoA, the central motor of the glideosome is coaligned with subpellicular microtubules, arguing for a direct or indirect connection between the acto-myosin system and subpellicular microtubules. This leads to provocative questions regarding the exact location and orientation of the actomyosin system at the IMC (Tardieux & Baum, 2016).
Current models (Soldati, Foth, & Cowman, 2004;Tardieux & Baum, 2016) suggest that the acto-myosin system is localized between the IMC and the PM of the parasite. This space is very narrow (20-30 nm) and surprisingly electron lucid, indicating a low density of proteins. In contrast, just below the IMC is the so called sub- a ring-like junction through which they actively invade the host cell (Besteiro, Dubremetz, & Lebrun, 2011;Riglar et al., 2011). During this process the parasite is deformed and a recent study suggests that the host cell exerts counter-pressure on the junction, which can result in abortive invasion events, especially when components of the actomyosin system are disrupted (Bichet et al., 2014;Bichet et al., 2016).
This situation is akin to other eukaryotes, where the nucleus represents a major obstacle for the migration through a constricted environment (McGregor, Hsia, & Lammerding, 2016). When F-actin dynamics of invading parasites were analysed, a meshwork surrounding the nucleus could be detected, leading to the hypothesis that Factin, potentially in concert with the subpellicular microtubules facilitates nuclear entry through the junction in a push-and-pull mechanism, as observed for other motile eukaryotic cells when moving through constricted environments  ( Figure 1d). Indeed, an integration of the nucleus with the cytoskeleton is observed in most eukaryotes and this is facilitated by the socalled LINC-complex. However, to date, the components of the LINC complex in apicomlexan parasites (similar to many nuclear envelope proteins) remain unknown (Rout, Obado, Schenkman, & Field, 2017).
On the other hand, the requirement of the actomyosin machinery for invasion into erythrocytes by P. falciparum seems to be absolute.
Three different components of the glideosome were conditionally knocked out in three independent studiesnamely PfACT1 (Das et al., 2017), PfMyoA (Blake, Haase, & Baum, 2020) and PfGAP45 (Perrin et al., 2018), and in each of these cases, a complete abrogation of invasion was observed. These observations may be explained by the relatively larger size of the nucleus in comparison to the merozoite, differences in shape of the merozoite and tachyzoite, differences in stiffness of the erythrocyte membrane compared to other mammalian cell membranes, or by different requirements of the Plasmodium invasion machinery compared to Toxoplasma.

| The role of F-Actin during intracellular replication
Visualisation of F-actin in T. gondii and P. falciparum using Cbs demonstrated that individual parasites within the PV are connected via an extensive intravacuolar network that appears to be critical for the organisation of parasites within the PV, for regulation of parasite replication and material exchange between parasites (Periz et al., 2017;Periz et al., 2019;Stortz et al., 2019). In good agreement, conditional mutagenesis of Tgact1 results in asynchronous replication, aberrant parasite organisation within the PV and a blockade in co-ordinated parasite egress (Periz et al., 2017). Furthermore, the intravacuolar network is highly dynamic and its formation and disassembly during egress appears to be tightly regulated, suggesting the presence of unknown regulatory mechanisms (Periz et al., 2017).

| The F-Actin network is connected with the apicoplast
The use of actin-chromobody revealed that individual parasites in Toxoplasma (and Plasmodium with notable variations) are connected by a highly dynamic filamentous actin network. In T. gondii, this network flows through the parasite cytosol and through the posterior end, thereby forming a dynamic network in the parasitophorous vacuole that connects daughter parasites via the residual body. This network is in close association with formin-2 and the apicoplast in both T. gondii and P. falciparum (Stortz et al., 2019), (Figure 1e,f). A deletion of formin-2 resulted in disappearance of the F-actin cytosolic network followed by accumulation of apicoplast(s) within the residual body (T. gondii) or near the food vacuole (P. falciparum), showing that formin-2 acts as a nucleator for the cytosolic actin network and F-actin is essential for apicoplast inheritance to the daughter cells in both apicomplexan genera (Stortz et al., 2019). While formin-2 appears to be the only formin required in formation of the network in the case of P.falciparum, in the case of T.gondii a third formin, formin-3 is also involved in formation and organisation of the network (Tosetti et al., 2019).
Complementary data also supports the role of F-actin in apicoplast inheritance. Perturbation of actin dynamics by depletion of profilin, ADF or overexpression of the formin-2 FH2 domain led to defects in apicoplast inheritance. Finally, a conditional mutant for myoF, the gene encoding the unconventional Myosin F that localizes in proximity to the apicoplast also resulted in loss of the apicoplast and parasite death (Heaslip et al., 2016;Jacot et al., 2013).  Together these results led us to propose that vesicle trafficking occurs via at least two inter-connected mechanisms (Figure 2a,b): one that is actin independent and is possibly associated to the microtubule network and a second mechanism that relies on F-actin in close association with the parasite membrane , (Figure 2c,d).

| F-actin network and vesicular transport
The F-actin network may facilitate vesicle transport in two distinct ways: material transport can occur on actin tracks in a myosindependent manner as described for myosin VI in other eukaryotes (Frank, Noguchi, & Miller, 2004). Second, actin bundles appear to be highly mobile themselves and able to associate transiently to each other and thus, can transport and exchange associated vesicles between two membrane sites. This observation resembles data in transport associated with F-actin polymerisation driven by actin comet like trails, as discussed previously (Khaitlina, 2014).
As circular actin flow was observed within the residual body (RB), the RB was suggested to be a major sorting station for recycling and distribution of material between parasites . Using SIM, a close proximity between cytosolic F-actin, parasite membrane compartments and microtubules was demonstrated, suggesting a physical linkage between cytoskeletal components regulating cytosolic vesicular transport . The connection between actin and microtubules has been investigated in higher eukaryotes and it is becoming increasingly clear that a large number of protein complexes including myosins, plus end tracking factor, formins or septins can crosslink actin and microtubules directly or indirectly. This extensive crosslinking of microtubules and actin enables the sharing and co-ordination of material transport depending on these structures (Dogterom & Koenderink, 2019).

| F-actin in egress from the host cell
The importance of F-actin in egress was shown by conditional gene deletion studies. The results of these experiments show a complete abrogation of egress in Toxoplasma gondii but not in Plasmodium falciparum. This could be in part due to the fact that osmotic pressure and outward curling of the erythrocyte membrane plays an important role in the explosive dissemination of P. falciparum merozoites (Das et al., 2017). Another observation suggests that the actin network is important for preserving the structure of the parasitphorous vacuole and maintaining communication between parasites, so that they can respond in a co-ordinated fashion to external stimuli during egress.
This idea is supported by experiments in which the collapse of the actin nanotubular network after treatment with calcium ionophore appears to be a prerequisite for the release of the tachyzoites from the parasitophorous vacuole (Periz et al., 2017).

| F-actin regulation: The role of ABPs in maintaining the Actin network
With the establishment of Cbs in apicomplexans, it is now possible to readdress the role of previously described ABPs and their influence on F-actin polymerisation and dynamics in vivo. Apicomplexan parasites, including Toxoplasma and Plasmodium species, possess a limited set of ABPs (Baum et al., 2006). For example, Toxoplasma encodes a single gene each for ADF and profilin, while three genes encode formins. Plasmodium falciparum encodes only two formins and two ADFs (Schuler, Mueller, & Matuschewski, 2005;Baum et al., 2006). In comparison, humans possess 5 profilin genes, 14 adf/cofilin genes and 16 formin genes (Baum et al., 2006). Most actin nucleation factors, such as spire (Baum et al., 2006) or Arp2/3 (Gordon & Sibley, 2005;Baum et al., 2006) are missing in apicomplexan parasites.
Apicomplexan parasites retain most of their cellular actin as monomeric G-actin. While structural features of the molecule contribute to physiologically relevant filament instability in vivo, ABPs such as ADF, profilin, CAP and CPs also play important roles (Figure 1a).
TgADF was first described in 1997 as a single copy gene (Allen, F I G U R E 2 Recycling of proteins during the replication cycle of Toxoplasma gondii. (a) After invasion, the newly formed PV contains microneme organelles containing unused micronemal proteins that have not been secreted and can be recycled (MICr) during replication. During endodyogeny, the mother cell is disassembled, and MICr and other proteins from the IMC accumulates at the RB. The RB is a sorting station for recycled material. Newly synthesised micronemal proteins (MICn) are transported towards the apical tip of daughter parasites. The microneme vesicles accumulated in the RB are redistributed to the final location at the apical tip of daughter parasites, where recycled MICr and newly synthesised MICn accumulate in the micronemes. This mechanism of recycling and sorting material is also applicable for components of the IMC. (b) This multidirectional transport of material is supported by a network of F-actin that links mother, daughter and the RB. There are two population of vesicles, recycled vesicles appear to be associated to F-actin bundles, and a second population of vesicles synthesised de novo that is not linked to F-actin. These observations raise the possibility of several models supporting vesicle transport. (c) Two sorting systems based mechanism: this model integrates microtubule based transport and membrane based sorting system with F-actin and contributes to recycling of vesicles. Vesicles can be transported on microtubules and actin may control the proximity and exchange of vesicles transported on microtubules and acts as a barrier for vesicle transport. (d) Actin dependent transport model: F-actin dynamics regulate vesicular transport and cluster formation. We speculate that two populations of micronemes can be observed, 1: formed by recycled vesicles (MICr) that are redistributed using a mechanism dependent on F-actin and 2: formed by micronemes synthesised de novo and associated with the secretory pathway. Recycled vesicles use F-actin tracks and mobile F-actin for transport and migrate to specific cellular locations using myosin based transport and/or comet like transport. Localisation and cluster density of the vesicles are controlled by flow, bundle compaction and mobility of the F-actin scaffold. Depolymerisation of the actin tracks results in an accumulation of vesicle clusters containing adhesins or other material in specific sites. Alternatively bundling and stabilisation of the F-actin network may also lead to a blockade in vesicle transport, resulting in specific localisation of vesicle clusters Dobrowolski, Muller, Sibley, & Mansour, 1997). Recombinant TgADF is capable of binding to G-actin and of depolymerising F-actin in vitro (Allen et al., 1997). In vivo, a cytosolic localisation was reported for TgADF by antibody staining and endogenous tagging (Allen et al., 1997;Haase et al., 2015;Mehta & Sibley, 2011). Depletion of TgADF resulted in accumulation of actin structures and compromised host cell invasion, egress and overall gliding motility, making TgADF essential for the lytic replication cycle (Mehta & Sibley, 2011;Periz et al., 2017). Plasmodium falciparum has two adf genes, of which only ADF1 is essential and expressed throughout all life-cycle stages. Differing structurally from mammalian ADF/Cofilins, PfADF1 was shown to be unable to bind to F-actin in vitro, but functioned in sequestering G-actin, and surprisingly similar to mammalian profilins, promoted nucleotide exchange on monomers (Schuler et al., 2005). In a later report, PfADF1 demonstrated F-actin severing activity without stable binding, but via a low-affinity binding interface (Wong et al., 2014).
In divergence from classical profilins, apicomplexan profilins contain an additional β-hairpin loop, which is critically required for actin monomer binding, and a single point mutation in this loop region eliminates a hydrogen-bond, thereby abrogating fast motility seen in P. berghei sporozoites (Moreau et al., 2017). Recently, the same group produced mutations in another acidic loop in profilin, which did not affect actin polymerisation in vitro and yet affected gliding motility of P. berghei sporozoites, indicating that additional factors beyond actin polymerisation are at play during motility and invasion of apicomplexan parasites (Moreau et al., 2020). TgProfilin is critical for the completion of the lytic life cycle as depletion of TgProfilin rendered parasites defective in gliding motility, invasion and host cell egress (Plattner et al., 2008). Intracellular replication was not affected by TgProfilin loss. In a comparative study of yeast, mouse and P. falciparum cyclase associated protein (CAP), a common conserved mechanism of nucleotide exchange on G-actin-ADP monomers via a β-sheet domain was found. In comparison to PfCAP, higher CAPs contained additional domains which might have later evolved for more complex dynamics (Makkonen, Bertling, Chebotareva, Baum, & Lappalainen, 2013). Toxoplasma tachyzoites lacking CAP exhibited impaired motility, invasion and egress (Hunt et al., 2019). TgCAP also appears to play a role in dense granule trafficking. Depletion of TgCAP  TgFormin2 to the vicinity of the apicoplast (Stortz et al., 2019;Tosetti et al., 2019). Furthermore, Tosetti and colleagues reported colocalisation of TgFormin2 and the Golgi apparatus (Tosetti et al., 2019). In Toxoplasma, conditional depletion of TgFormin1 suggested that it is not important for intracellular replication and not critical for egress (Daher et al., 2010). Instead, TgFormin1 is involved in tachyzoite motility and host cell invasion. A recent publication confirmed the importance of TgFormin1 in gliding and invasion, but also indicated a critical role in parasite egress (Tosetti et al., 2019). In Plasmodium, PfFormin1 was reported to localize to the apical tip of free merozoites and to follow the moving junction during invasion, implicating it in the process (Baum et al., 2008).
TgFormin2 was reported to be involved in intracellular replication as its deletion caused an increase in aberrant daughter cell orientation (Tosetti et al., 2019). By obtaining a clonal TgFormin2-KO line, Tosetti and colleagues showed that TgFormin2 is not essential for the completion of the lytic cycle. However, the line was rapidly outgrown by wild-type parasites in growth competition assays (Tosetti et al., 2019).
PfFormin2 was initially reported to have a diffuse cytoplasmic distribution in trophozoites (Baum et al., 2008). A comparative study of Formin-2 function in Toxoplasma and P. falciparum, revealed a localisation at the apicoplast vicinity in both genera (Stortz et al., 2019). Plasmodium falciparum parasites lacking functional Formin-2 showed a complete abrogation of detectable F-actin within growing and replicating asexual Plasmodium stages, making it the primary actin nucleator during asexual growth stages (Stortz et al., 2019).
TgFormin3-KO tachyzoites do not display any replication defects, making TgFormin3 dispensable for the lytic cycle (Daher, Klages, Carlier, & Soldati-Favre, 2012). Initially, TgFormin3 was localized to the apical and the basal pole as well as around the mitochondrion (Daher et al., 2012). In a later study TgFormin3 was localized to the basal pole and the residual body based on endogenous tagging (Tosetti et al., 2019). Asynchronous replication was observed in parasites lacking TgFormin3 (Tosetti et al., 2019). In addition, recovery of fluorescence in photobleaching experiments was slower for TgFormin3-KO parasites when compared to wild-type parasites. Tosetti  Together this will allow us to further understand actin dynamics and its regulation in apicomplexan parasites. As a starting point, we are proposing an actin treadmilling model that combines in vitro and in vivo data obtained from Toxoplasma and Plasmodium (Figure 1a). In this model, ADF acts to sever and depolymerise F-actin which reintroduces G-actin-ADP into the cytoplasmic G-actin pool. While Profilin sequesters G-actin, CAP proteins mediate the exchange of ADP to ATP. Formin nucleation factors can then enhance actin polymerisation, resulting in the quick assembly of F-actin at various polymerisation centres, akin to other eukaryotes. More stable actin structures such as the filamentous structures connecting individual parasites within the PV might be less affected by impaired actin treadmilling. Further in vivo research is needed to elucidate the precise functions of Profilin, ADF, CAP and CPs. In the future, the chromobody technology will be immensely useful to uncover novel ABPs and to further unravel the nature and function of the apicomplexan actin network.

ACKNOWLEDGMENTS
We thank all the investigators who have contributed to this body of knowledge, some of whom were not cited due to space limitations.