ookinete) of apicomplexan parasites [5,6], the phylum to which malaria parasites belong. Furthermore, each retains a conserved way of moving and invading cells based on actin and myosin, termed gliding motility [7]. The current model for the gliding motor [8] is centred on a short single-headed myosin that is attached to a double membrane bound complex of organelles called the inner membrane complex (IMC), which lies directly under the plasma membrane [9]. Microfilaments of actin are then thought to form in the intervening space, called the supra-alveolar space, between IMC and plasma membrane [10]. On polymerisation these actin filaments are thought to provide the key rigid element upon which myosin bears to create the required rearward traction force for movement. The myosin stroke is then conveyed, via the actin filament, to surface bound adhesins from the thrombospondin-related anonymous protein (TRAP) family [11]. Coupling of TRAP to surface receptors in the extra-cellular milieu, transmits the internal rearward force, driving the parasite forwards [12]. The topology of this motor model is largely based on immunoprecipitation data along with immunofluorescence imaging of several core components in malaria parasites and the related apicomplexan parasite Toxoplasma gondii [8,11,13,14]. According to the current model for gliding motility, actin filaments only form transiently at sites where the gliding motor is engaged – in other words, sites where traction is being applied between the parasite motor and the substrate surface or host cell [15]. In parasites that are focussed primarily on movement in the absence of true invasion, for example the ookinete [4], this could be anywhere along the length of the cell where the stage-specific TRAP-like adhesin is linked to the extra-cellular environment. For parasites that actively invade host cells, such as the sporozoite and merozoite, this would presumably be restricted to the tight junction – an electron dense interface formed between the invading parasite and its host cell [16]. Whilst the molecular architecture of the junction has been elucidated in great detail recently [17], its linkage to the actomyosin motor is currently unknown. Indeed, to date, and despite its essential role in motility, no study has provided direct visual evidence for the placement of actin filaments in the parasite pellicle of any moving apicomplexan cell [18,19,20,21]. The evidence in support of actin’s role in motility comes largely from actin inhibitors that disrupt parasite gliding and host cell invasion [15,22,23,24]. Of these, the marine sponge cyclodepsipep- tide Jasplakinolide (JAS) has proven particularly useful, binding to and stabilising formed filaments preventing disassembly [25]. Use of JAS has facilitated the demonstration of high concentrations of dynamic actin at the apex of free Plasmodium merozoites, ookinetes and T. gondii tachyzoites [21,24,26,27]. Microfilament structures, presumed to be actin, have also been seen lying under the plasma membrane of motile tachyzoites following JAS treatment [28]. Complementing these studies, electron and cryo-electron micros- copy studies have observed structures with dimensions consistent with filamentous actin in this pellicular compartment under native conditions [19,20]. Beyond these encouraging observations, how- ever, no study has unambiguously demonstrated microfilament spatial organisation during zoite movement under native conditions. This likely derives from the intrinsic short length of apicomplexan actin filaments (,100 nm), their instability, dynamic and transient nature and the poor utility of conventional filament markers such as phalloidin with apicomplexan cells [29,30,31,32]. Aside from motility actin likely plays several additional roles in parasite development, including roles in haemoglobin uptake [33] and general vesicular trafficking [34] along with several possible functions in the nucleus [35]. However, like motility, these roles have remained incompletely explored because of difficulties in decisively localising actin and its microfilaments within parasite Actin Filaments in Plasmodium Development cells. To visualize the spatial dynamics of malaria parasite actin we generated mouse and rabbit parasite-specific antibodies towards actin I (the conserved isoform implicated in most actin-dependent processes across Apicomplexa [36]) that recognises filamentous actin in preference to monomeric actin. We employed these tools on mouse and human malaria parasites to gain access to the three major motile or invasive lifecycle forms (ookinete, sporozoite and merozoite) along with asexual blood stages and tachyzoites from T. gondii to provide a map for dynamic actin filament formation. We demonstrate actin concentrates in discrete zones in the nuclear compartment during development, within the supra-alveolar space during motility, and at sites predicted to be core regions of traction during host cell invasion. These results point to new functions for actin in parasite development and refine current understanding of the role of microfilaments during key stages of parasite infection. Results Generation of a malaria parasite actin-specific antibody Conventional antibodies against mammalian actin have been used successfully to label the entire actin pool in Toxoplasma gondii tachyzoites [37] and Plasmodium merozoites and ookinetes [22,38]. However, these antibodies cannot differentiate monomeric (G)- from filamentous (F)- actin and have the added drawback of also recognising host cell actin with equal or greater affinity. Serum generated against a short peptide corresponding to amino acids 237–251 of non-muscle mammalian actin, anti-Gly245 [39] (Fig. 1A), has been reported to preferentially recognise short actin filament ends associated with vesicle transport in human fibroblasts [40]. This epitope, on sub-domain 4 of the actin monomer, is exposed in free actin monomers and at the end of the filamentous form (Fig. 1B). The specificity for short filament ends is thought to result from the epitope being hidden in the body of filaments (from subunit contact), long filament ends (as a result of capping) and in free monomers either by virtue of the topology of the epitope in monomers versus filaments (Fig. 1B) or because of association with actin binding proteins in the cell cytosol [40]. We raised antiserum in rabbits and mice to the homologous epitope of P. falciparum actin I (PFL2215w, amino acids 239–253), which is conserved across most Apicomplexa (Plasmodium, Toxoplasma, Theileria and Babesia spp.) but not outside of the apicomplexan phylum. Of note, this sequence diverges at three residues from mammalian beta-actin (Fig. 1A,B). We recently reported that rabbit serum against this peptide, which we refer to as anti-Act239–253, reacted specifically with cell lysate from P. falciparum asexual stages, but showed poor reactivity with erythrocyte actin (reported in [41]). Immunoblots with rabbit and mouse antisera confirmed the specificity of this reactivity against human parasite lysate, and extended the observation to lysates of mouse malaria parasites and T. gondii (recognising a specific product of ,40 kD consistent with the predicted masses of the respective actins: 41.8, 41.9 and 41.7 kD (Fig. 1C). When compared to conventional vertebrate actin antibodies the antiserum showed minimal cross-reactivity with mouse erythrocyte, human erythro- cyte or human fibroblast actin (Fig. 1D). Thus, based on only a few divergent residues, an antibody that differentiates between human and parasite actin has been generated. Actin dynamics localise to pellicular and apical regions of motile or invasive zoites To explore the general utility of anti-Act239–253 against all motile or invasive lifecycle stages and beyond P. falciparum [41,42], we used the mouse malaria parasite P. berghei, which greatly facilitates generation of each zoite form: merozoite, ookinete and sporozoite. Actin labelling by immunofluorescence was seen to PLoS ONE | www.plosone.org 2 February 2012 | Volume 7 | Issue 2 | e32188