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Keywords:

  • malaria;
  • Plasmodium;
  • actin;
  • microfilaments;
  • cofilin;
  • actin-depolymerising factor;
  • cyclase-associated protein;
  • profilin;
  • coronin;
  • formin;
  • crystallography;
  • protein structure

Abstract

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. ACTIN AND ITS DYNAMICS
  5. ADF/COFILIN
  6. CYCLASE-ASSOCIATED PROTEIN
  7. PROFILIN
  8. FUTURE DIRECTIONS
  9. Acknowledgements
  10. REFERENCES

Parasites from the phylum Apicomplexa are responsible for several major diseases of man, including malaria and toxoplasmosis. These highly motile protozoa use a conserved actomyosin-based mode of movement to power tissue traversal and host cell invasion. The mode termed as ‘gliding motility’ relies on the dynamic turnover of actin, whose polymerisation state is controlled by a markedly limited number of identifiable regulators when compared with other eukaryotic cells. Recent studies of apicomplexan actin regulator structure—in particular those of the core triad of monomer-binding proteins, actin-depolymerising factor/cofilin, cyclase-associated protein/Srv2, and profilin—have provided new insights into possible mechanisms of actin regulation in parasite cells, highlighting divergent structural features and functions to regulators from other cellular systems. Furthermore, the unusual nature of apicomplexan actin itself is increasingly coming into the spotlight. Here, we review recent advances in understanding of the structure and function of actin and its regulators in apicomplexan parasites. In particular we explore the paradox between there being an abundance of unpolymerised actin, its having a seemingly increased potential to form filaments relative to vertebrate actin, and the apparent lack of visible, stable filaments in parasite cells. 2012 IUBMB IUBMB Life, 2012


INTRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. ACTIN AND ITS DYNAMICS
  5. ADF/COFILIN
  6. CYCLASE-ASSOCIATED PROTEIN
  7. PROFILIN
  8. FUTURE DIRECTIONS
  9. Acknowledgements
  10. REFERENCES

Apicomplexa are an ancient eukaryotic phylum of obligate intracellular parasites that are the causative agents of diseases such as malaria and toxoplasmosis. Their ability to traverse and invade host cells is central to development and is mediated by a unique and conserved mode of movement, termed ‘gliding motility’, based on an actomyosin motor complex located in the pellicle of the parasite (1). Biochemical studies have demonstrated that, at its core, parasite motor function relies on a dynamic microfilament of actin. Actin is a conserved ubiquitous eukaryotic protein that lies at the heart of many cellular processes via its ability to cycle between a monomeric and polymerised form (reviewed in ref.2). Indeed, it is this cycling that is thought to constitute a ‘molecular clutch’, critically controlling the ability of parasites to move and invade host cells (3, 4). However, despite extensive effort to visualise filaments in parasite cells, no study to date has definitively seen microfilaments of any form under native conditions. Furthermore, the signals that govern the key changes in actin filament state in motile parasite cells are still poorly understood.

Whilst in most eukaryotic cells the polymerisation state of actin is tightly governed by 100 or more regulators (2), a drastically reduced repertoire appears to carry out this key function in Apicomplexa (5, 6). This minimal set includes formins (7, 8), actin-depolymerising factor (ADF)/cofilin 1 and 2 (9–11), cyclase-associated protein (CAP) (12), profilin (13–15), coronin (16) and capping protein (CP) (17). The crystal structures of three of these apicomplexan regulators have recently been determined, providing unique insights into the molecular basis of actin regulation (9, 10, 12, 14). Furthermore, new revelations regarding apicomplexan actin are beginning to shed light on our understanding of the molecular basis for parasite cell movement. In this review, we aim to summarise the recent advances in apicomplexan actin regulation with a particular focus on structural and biochemical insights, exploring fundamental properties of actin and the role of its regulators in holding back the actin monomer pool from forming filaments.

ACTIN AND ITS DYNAMICS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. ACTIN AND ITS DYNAMICS
  5. ADF/COFILIN
  6. CYCLASE-ASSOCIATED PROTEIN
  7. PROFILIN
  8. FUTURE DIRECTIONS
  9. Acknowledgements
  10. REFERENCES

As one of the most abundant and conserved proteins in eukaryotic cells, actin has been co-opted for a wide range of cellular functions (reviewed in ref.2). It exists in two distinct states, monomeric or globular actin (G-actin) and filamentous actin (F-actin), with transition between the two states governed by its binding to divalent cations and ATP (reviewed in ref.2). Apicomplexan actins are highly divergent from other eukaryotic actins, sharing approximately 75–80% sequence identity with their mammalian counterparts (18, 19) (Fig. 1A). Toxoplasma gondii has one actin gene, TgACT (19, 20), whereas Plasmodium spp. have two genes (18). ACTI is expressed throughout the Plasmodium lifecycle, whereas ACTII is expressed predominantly in the sexual stages. The bulk of actin (∼98%) in apicomplexan cells is thought to exist in a soluble form, presumably in a monomeric state (19, 21). This imbalance may partly explain the elusive nature of actin filaments in motile parasite cells when visualized under native conditions (20, 22) and suggests an important role for cellular factors that sequester monomers from forming filaments. However, the failure to detect apicomplexan microfilaments is also certainly affected by their short length, rapid turnover and apparent instability (23–26).

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Figure 1. Divergent properties of PfACTI. A: PfACTI monomer model (based on actin PDB accession number 1ATN). Key residues implicated in filament stability are coloured: D180 (orange, conserved), G200 (green) and K270 (blue). B: Filament model of PfACTI (based on cryoEM structure of RbActin filament, 3G37.pdb). C: Close-up of protomer interface of RbACT filament model displaying stabilising residues: S199 (green) forms a hydrogen bond (dashed line) with D179 (orange), M269 (blue) on hydrophobic loop. D: Close-up of protomer contact interface of modelled PfACTI filament demonstrates loss of hydrogen bonding between G200 in red protomer with D180 in orange protomer and loss of hydrophobicity in loop containing K270, likely leading to filament instability (based on data from ref.26). [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

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Biochemical studies with purified recombinant TgACT have shown that it polymerises rapidly in vitro at a very low critical concentration (Cc), defined as the concentration of G-actin monomers in equilibrium with actin filaments above which filaments form and below which no polymerisation occurs (2). Whilst Cc values of between 0.1 and 0.6 μM are seen at the filament fast-growing or barbed end for mammalian actins, values of ∼0.03–0.04 μM were estimated for TgACT filaments (23). Even with this increased propensity to polymerise, TgACT forms exceptionally short filaments of ∼100 nm or less (23, 26), with similar observations seen for recombinant actins (PfACTI and II) and parasite-cell-extracted actin (PfACTI) of P. falciparum (24–26). As these filaments still move over rabbit skeletal myosin during in vitro motility assays and at similar velocities to mammalian actin, they are clearly functional and likely retain the canonical filament structure (24, 27). However, their short length suggests that apicomplexan actin filaments are inherently unstable.

Several lines of evidences have suggested possible reasons for this apparent filament instability. First, methylation of H73, a typical modification of other actins, is absent in PfACTI (24). This lack of methylation would potentially cause a conformational change in the monomer, giving it a more ‘open’ conformation (as seen in yeast actin), leading to a decrease in the stability of the filament (28). A recent thorough mutational study of apicomplexan actins identified several additional key residues that affect the stability of actin filaments (26). Modelling of the actin filament (Fig. 1B) suggests mammalian actin residue S199, located at the top of sub-domain four, forms a hydrogen bond with D179 in sub-domain 3 of the adjacent longitudinal protomer in the filament (Fig. 1C). In Apicomplexa, the aspartic acid residue (D180) is conserved but S199 is replaced with G200 (Fig. 1D), abolishing the hydrogen bond. In the same protomer contact interface, a hydrophobic loop containing M269 in mammalian actin is replaced by K270, causing a loss of hydrophobicity. Without these interactions, the stability of apicomplexan actin filaments is expected to be reduced. Indeed, substitutions of the conserved residues (G200S and K270M) in recombinant TgACT dramatically restore filament stability (26). Furthermore, the expression of mutant actins that recover filament stability in vivo greatly retard parasite motility and egress (26). These results indicate that the inherent instability of apicomplexan actins is a critical adaptation for the effective functioning of gliding motility.

The overall instability of the filament population combined with a predominance of G-actin over F-actin in malaria and Toxoplasma cells suggests that the apicomplexan actin regulatory machinery is heavily biased towards actin in a monomeric state. This is supported by the apparent lack of most known filament-binding proteins in apicomplexan genomes (5, 6). Retaining this imbalance implicates the G-actin binding proteins, in particular ADF/cofilin, CAP and profilin, as critical regulators for sustaining the monomer-filament status quo.

ADF/COFILIN

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. ACTIN AND ITS DYNAMICS
  5. ADF/COFILIN
  6. CYCLASE-ASSOCIATED PROTEIN
  7. PROFILIN
  8. FUTURE DIRECTIONS
  9. Acknowledgements
  10. REFERENCES

The ADF/cofilin family of regulators are G- and F-actin–binding proteins that sever and depolymerise actin filaments and sequester actin monomers (29). ADF/cofilins have a conserved fold (the ADF-H domain), which is also found in other classes of actin-binding proteins, such as twinfilins, consisting of a five-stranded β-sheet surrounded by four α-helices (Fig. 2A). The only structure solved to date of an ADF-H domain containing protein bound to G-actin is that of the C-terminal domain of mouse Twinfilin-1 (TWF-C; ref.30). The G-actin-binding site (G-site) is formed by the N-terminus, the long α3-helix containing several positively charged residues and a turn that connects the β6-strand to the C-terminal α4-helix (Fig. 2A). When bound to G-actin, the N-terminal region of TWF-C interacts with sub-domain 1 of G-actin, the α3-helix sits in the cleft between sub-domains 1 and 3 and the turn connecting β6-strand and the α4-helix interacts with sub-domain 3 (30).

thumbnail image

Figure 2. Comparison of apicomplexan G-actin-binding proteins and their yeast/human counterparts. A: ADF/cofilin family. Residues implicated in filament severing are highlighted in red, and residues implicated in G-actin binding are highlighted in yellow on the α3 helix and green on the β6 strand. Structures shown are for PfADF1 (PDB accession number 3Q2B), PbADF2 (2XFA), TgADF (2L72) and ScCOF (1QPV). G-actin-binding site is conserved, whereas PfADF1/TgADF lacks the F-loop. B: CAP family. Dimerisation involves domain swapping, hydrophobic residues (highlighted in green) and stacking residue interactions (highlighted in yellow). Structures shown are for CpCAP (2B0R) and HsCAP (1K8F). C: Profilin family. Basic residues implicated in G-actin binding are highlighted in blue, acidic residues in the apicomplexan-specific acidic loop and unique mini-domain are highlighted in red and aromatic residues implicated in polyproline binding are highlighted in yellow. Structures shown are for PfPfn (2JKF), TgPfn (3NEC) and HsPfnII (1D1J). [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

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ADF/cofilins bind preferentially to ADP-Pi-actin on filaments (binding in the vicinity of the slow-growing pointed end) and stimulate the dissociation of Pi. The F-actin-binding site (F-site) of ADF/cofilin, mapped via mutagenesis studies, consists of a β4/5-strand protrusion [also known as the filament-binding loop (F-loop); Fig. 2A] and the C-terminal consists of a α4-helix (31, 32). Charged residues in these regions (K79, R80 and K82 on β5 of ScCOF, E134, R135 and R138 on α4 of ScCOF, K95 and K96 on HsCOF) have been implicated in mediating filament interactions. Cryoelectron microscopy of actin filaments decorated with HsCOF indicates that upon binding, substantial alterations in sub-domain 2 of the actin protomers are introduced, leading to a unique conformational change in the actin filament and consequently increased filament flexibility (33).

P. falciparum has two ADF/cofilin isoforms, PfADF1 and PfADF2. Both are highly divergent, sharing ∼30% sequence identity to ADF/cofilins from other organisms (34). PfADF1 is expressed throughout all motile and non-motile erythrocytic stages and of the two variants is the one implicated in cell motility (9, 34). Biochemical characterisation of PfADF1 suggests that it is able to form a complex with G-actin with a preference for ADP-G-actin (by gel shift assay) (34). The same study found that PfADF1 was also capable of stimulating nucleotide exchange on actin monomers, a function usually performed by profilins. Analyses from several groups have shown that PfADF1 has very low affinity for F-actin, an unusual characteristic within the ADF/cofilin family (34). Two recent studies solved the crystal structure of PfADF1 (9, 10), demonstrating that it displays the basics of the canonical ADF-H fold with a six-stranded β-sheet flanked by four α-helices. Structural comparison with ScCOF indicates that the G-site of PfADF1 is intact (Fig. 2A); however, the β6-strand is significantly shorter and lacks homology to other ADF/cofilins. Because residues in β6 of TWF-C (K294 and E296) are known to mediate G-actin binding as well as affecting nucleotide exchange (30), divergence in β6 may partly relate to the unusual ability of PfADF1 to promote nucleotide exchange. A crystal structure of PfADF1 in complex with G-actin will be vital for further understanding this activity.

The lack of interaction of PfADF1 with F-actin, in terms of both filament binding and severing, would be expected given the divergence in the F-actin-binding domain, specifically the F-loop. Structural analysis reveals that the β4–β5 protrusion (the F-loop) and the C-terminal α4 helix are markedly reduced in PfADF1 when compared with ScCOF and lack most of its associated charge (Fig. 2A) (9). Despite this, it has now been convincingly demonstrated that PfADF1 and its orthologue in T. gondii (recently solved by NMR solution structure; ref.11) can sever F-actin at rates comparable with SpCOF and HsCOF1 (9, 35). Chimeric PfADF1 mutants established that, in fact, only a single-solvent-exposed basic residue, K72, in the shortened F-loop is required for filament severing. This minimal motif is conserved across all eukaryotic ADF/cofilins capable of filament severing, including K82 in ScCOF, K95 in HsCOF (9) and also K86 in Plasmodium ADF2, which unlike ADF1 shows a much more canonical structure (10). Importantly, this establishes that severing can occur without high-affinity filament binding. Authentication of PfADF1 as an actin filament severing protein and dramatic demonstration of actin deregulation (apparent filament accumulation) following TgADF knockdown (36) suggests that ADF1 likely plays a critical role in regulating actin filament state during the process of parasite motility. This essential role is further supported by the apparent inability to knockout PbADF1 in P. berghei (34).

Several mechanisms are used to regulate the cellular activity of ADF/cofilins, including binding to membrane phosphoinositides (in particular PI(4,5)P2) and phosphorylation (reviewed in ref.29). Neither PfADF1 or TgADF bind to PI(4,5)P2in vitro (9, 11) nor does the presence of PI(4,5)P2 affect TgADF binding to G-actin in vitro. Absence of this interaction is supported by the divergent overall surface charge of apicomplexan ADF/cofilins, which lack the large, positively charged residues required for PI(4,5)P2 binding in other systems (11). In its absence, phosphorylation is a likely mechanism behind ADF/cofilin control. Several ADF/cofilin proteins are regulated via phosphorylation of a serine residue near the protein N-terminus (S3 in mammalian ADF/cofilin), which inactivates function by disrupting the G-site (reviewed in ref.29). Of note, apicomplexan ADF/cofilins have a conserved serine (S3) phospho-site in the N-terminal tail (37). Indeed, recent proteomic analysis of Toxoplasma and Plasmodium cells identified two phosphopeptides, with strong support for phosphorylation at S3 (38). Remarkably though, analysis of apicomplexan genomes shows that there are no ADF/cofilin-specific kinases (LIMK, TESK) and only one chronophin homologue (29). This suggests that there may be an entirely novel ADF/cofilin phosphorylation system in Apicomplexa, possibly via calcium- or calmodulin-dependent protein kinases (39) as seen in plants (40).

CYCLASE-ASSOCIATED PROTEIN

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. ACTIN AND ITS DYNAMICS
  5. ADF/COFILIN
  6. CYCLASE-ASSOCIATED PROTEIN
  7. PROFILIN
  8. FUTURE DIRECTIONS
  9. Acknowledgements
  10. REFERENCES

CAP proteins and their relatives are among the most enigmatic actin regulators characterised to date. CAPs are potent G-actin-binding proteins found in all eukaryotes (41). Originally identified in Saccharomyces cerevisiae (Srv2), they consist of three domains: an N-terminal α-helical domain, a central proline-rich region incorporating a WASP homology 2 (WH2) domain and a C-terminal actin-binding domain. Of these three domains, the C-terminal domain has been shown to predominantly function in binding ADP-G-actin, which it does so with a 100-fold higher affinity than ATP-G-actin (42). As well as having higher affinity for actin in an ADP-bound state, CAP proteins are known to enhance nucleotide exchange of ADP-G-actin and to act cooperatively with ADF/cofilin and profilin to promote actin filament turnover (43, 44). As a result, it has been hypothesised that, post-nucleotide exchange, CAP may effectively pass ATP-G-actin to profilin for addition to a growing actin filament end. Thus, combined with its ability to block the addition of actin monomers to the filament barbed end, CAP may function as a revolving door, facilitating the cycling of actin from a depolymerised ADP-ADF/cofilin-bound state to a newly ready ATP state for profilin-mediated addition to the barbed end (41).

Apicomplexa contain a single CAP homologue, although each strikingly lacks the N-terminal and WH2 domains found in most other eukaryotes (12). Apicomplexan CAPs are therefore expected to have a function limited to monomer sequestration. Characterisation of CAP from the apicomplexan parasite Cryptosporidium parvum (CpCAP; a distant relative of Plasmodium and Toxoplasma spp.) reveals that it is a potent actin-monomer-binding protein, preventing barbed end elongation of F-actin (12). CpCAP binds G-actin in a mutually exclusive manner to DNase I; however, it has yet to be determined whether this is due to direct binding-site competition or allosteric inhibition. The crystal structure of CpCAP reveals a homo-dimer with the canonical fold of the CAP C-terminal domain (Fig. 2B). The cylindrical formation of β-sheets is stabilised by hydrogen bonding between the β-sheets via conserved hydrophobic and S/C residues (not shown). Dimerisation is achieved by ‘domain swapping’ of the β-strand protrusion formed by residues Q162-S179 at the C-terminus of each monomer (strong versus pale protrusion; Fig. 2B). This interaction is stabilised by several hydrophobic and stacking interactions (Fig. 2B) (12). It is yet to be determined how apicomplexan CAP performs its functions at the molecular level. To this end, a structure of CAP bound to an actin monomer is essential.

In vivo analysis of P. berghei CAP indicates that it is expressed only in the motile stages of the parasite, with highest expression in the merozoite though it is surprisingly not essential to the asexual phase of the life cycle. P. berghei knockout studies demonstrated CAP is vital for oocyst development in the mosquito midgut (the precursor of motile sporozoite stages), although further research is required to explain this role (12).

PROFILIN

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. ACTIN AND ITS DYNAMICS
  5. ADF/COFILIN
  6. CYCLASE-ASSOCIATED PROTEIN
  7. PROFILIN
  8. FUTURE DIRECTIONS
  9. Acknowledgements
  10. REFERENCES

Profilins are small (∼15 kDa), structurally conserved proteins that are responsible for providing a readily polymerisable pool of ATP-G-actin. They have been identified in all eukaryotic organisms, with some containing several isoforms. Although profilin sequences are not well conserved, protein structures do share homology (45). The ‘profilin-like’ fold consists of a seven-stranded antiparallel β-sheet, surrounded by four α-helices, two on each face (Fig. 2C). Profilins participate in many cellular processes including membrane trafficking, mRNA processing and cell movement (reviewed in ref.45). They bind G-actin and stimulate nucleotide exchange on ADP-G-actin monomers, bind to polyproline stretches in proteins, such as those found in the formin-homology domain (FH-1) and bind to phosphoinositides PI(4,5)P2 and PI(3,4,5)P3. The PI(4,5)P2 binding site on profilin overlaps those for G-actin and polyproline, and as such, PI(4,5)P2 binding is thought to regulate profilin activity (46). Furthermore, PI(4,5)P2 binding may, like ADF/cofilins, mediate localisation of profilin to actin-rich regions near the plasma membrane (47).

Analysis of apicomplexan genomes has identified a single profilin gene in each species (14). P. falciparum profilin (PfPfn) is an essential protein expressed across the asexual life cycle and peaks during late schizogony where it has a largely cytosolic localisation (8). The crystal structure of PfPfn shows a conserved profilin-like fold but with the presence of several unique structural elements, such as a β-hairpin protrusion and an extended acidic loop (Fig. 2C) (14). The β-hairpin, which spans 18 residues, is located between β2 and α2 forming an arm-like extension. Homology modelling of an in silico PfPfn–actin complex suggests that this β-hairpin may have a role in providing additional contact sites via several acidic residues (E64, D65 and E66; Fig. 2C) for several basic side chains (K284, R290 and K291) in sub-domain 3 of actin (14). The acidic loop (E42, E43 and D45; Fig. 2C) is relatively mobile in the structure, and its charged residues are conserved across Plasmodium spp.; however, its function is not known. In the context of binding to actin monomers, β4 and β5, which form the putative actin-binding site, are extended (48) and may control PfPfn binding affinity for PfACT. Across apicomplexan profilins, key residues that mediate actin binding are conserved, with in vitro data suggesting that they are able to sequester heterologous actin monomers, albeit with a lower affinity than mammalian profilins (14).

An important working partner of eukaryotic profilins are the formins, facilitators of the processive elongation of actin filaments that recruit profilin-bound G-actin to the filament barbed end via a proline-rich FH-1 domain (49). It has been shown that PfPfn binds to polyproline peptides, with the crystal structure of the PfPfn-octaproline peptide showing the N-terminal aromatic side chains, Y5, W7, Y10 and Y35 at β3 are involved in coordinating this interaction (14). However, it is not known whether apicomplexan profilins and formins interact in the same manner in vivo as putative FH-1 domain of PfFormins and TgFormins appeared to be relatively divergent lacking obvious polyproline tracks (7, 8). Furthermore, it has not yet been determined whether PfPfn stimulates nucleotide exchange (14). Interestingly, PfPfn binds only to phosphatidylic acid and some phosphoinositol monophosphates but not PI(4,5)P2 or PI(3,4,5)P3 (14).

Studies from T. gondii have provided the first definitive evidence that TgPfn is essential for several important processes in the parasite life cycle, which includes gliding motility, host cell invasion and egress (15). The crystal structure of TgPfn is similar to that of PfPfn, including the apicomplexan-specific β-hairpin protrusion and acidic loop (Fig. 2C) (13). Functionally, TgPfn only binds to G-actin weakly, promotes actin polymerisation in vitro and interestingly, in contrast to other mammalian profilins, appears to reduce nucleotide exchange. The relevance of these divergent properties, like those of the other monomer-binding proteins, is still elusive.

FUTURE DIRECTIONS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. ACTIN AND ITS DYNAMICS
  5. ADF/COFILIN
  6. CYCLASE-ASSOCIATED PROTEIN
  7. PROFILIN
  8. FUTURE DIRECTIONS
  9. Acknowledgements
  10. REFERENCES

The marked reduction of actin regulators in Apicomplexa when compared with other vertebrate cells clearly points to the three key monomer-sequestering regulators having a central role in determining the cellular state of actin. However, significant differences (and overlap) in their expected biochemical properties cloud our understanding of how actin is regulated. One possibility is that all three regulators have converged on a general monomer-sequestering role with some redundancy of function between regulators. Alternatively, apicomplexan parasites may have radically re-written the rules on cellular control of actin, shifting the focus from control of filaments to management of the G-actin pool. Indeed, the inherent instability of actin filaments in Apicomplexa may have released some of the evolutionary constraints of actin-binding proteins to evolve divergent properties with the unstable nature of apicomplexan actin possibly indicative of a level of self-regulation. Key questions for the future will be not only how each regulator interacts with parasite actin in its ADP and ATP bound states (and structural determination of this) but also how they act in concert. For example, understanding the combined interplay between ADF/cofilin, CAP and profilin, already reported for other organisms (e.g., ref.50), will be important in dissecting how each regulator is recruited to harness the enormous potential of the G-actin pool in the parasite cell for its myriad roles.

Acknowledgements

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. ACTIN AND ITS DYNAMICS
  5. ADF/COFILIN
  6. CYCLASE-ASSOCIATED PROTEIN
  7. PROFILIN
  8. FUTURE DIRECTIONS
  9. Acknowledgements
  10. REFERENCES

The authors apologise to many researchers in this field whose work have not been able to cite directly because of space limitation. This work was made possible through the Victorian State Government Operational Infrastructure Support and the Australian Government NHMRC IRIISS. This research is supported by the National Health and Medical Research Council of Australia (NHMRC) Project Grant Scheme (APP1024678) and the Human Frontier Science Program (RGY0071/2011). M.A.O. is supported by an NHMRC Dora Lush Postgraduate Research Scholarship (APP1018002). J.B. is supported by a Future Fellowship (FT100100112) from the Australian Research Council.

REFERENCES

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. ACTIN AND ITS DYNAMICS
  5. ADF/COFILIN
  6. CYCLASE-ASSOCIATED PROTEIN
  7. PROFILIN
  8. FUTURE DIRECTIONS
  9. Acknowledgements
  10. REFERENCES