Pivotal and distinct role for Plasmodium actin capping protein alpha during blood infection of the malaria parasite.

Accurate regulation of microfilament dynamics is central to cell growth, motility and response to environmental stimuli. Stabilizing and depolymerizing proteins control the steady-state levels of filamentous (F-) actin. Capping protein (CP) binds to free barbed ends, thereby arresting microfilament growth and restraining elongation to remaining free barbed ends. In all CPs characterized to date, alpha and beta subunits form the active heterodimer. Here, we show in a eukaryotic parasitic cell that the two CP subunits can be functionally separated. Unlike the beta subunit, the CP alpha subunit of the apicomplexan parasite Plasmodium is refractory to targeted gene deletion during blood infection in the mammalian host. Combinatorial complementation of Plasmodium berghei CP genes with the orthologs from Plasmodium falciparum verified distinct activities of CP alpha and CP alpha/beta during parasite life cycle progression. Recombinant Plasmodium CP alpha could be produced in Escherichia coli in the absence of the beta subunit and the protein displayed F-actin capping activity. Thus, the functional separation of two CP subunits in a parasitic eukaryotic cell and the F-actin capping activity of CP alpha expand the repertoire of microfilament regulatory mechanisms assigned to CPs.

Introduction

Pathogenic eukaryotes of the genus Plasmodium, the etiological agent of malaria, rely on active motility to infect their host cells (Frénal and Soldati-Favre, 2009; Sibley, 2011; Montagna et al., 2012). Progress through their life cycle is characterized by continuous stage conversion and a programmed sequence of proliferating and invasive stages (Sattler et al., 2011; Montagna et al., 2012). During the brief extracellular phases, parasites adopt distinct motile forms, termed merozoites, ookinetes and sporozoites. Unlike the slow, amoeboid motility displayed by other eukaryotic cells, gliding motility of Plasmodium, and other apicomplexan parasites, is fast (1–3 μm s−1) and does not involve apparent changes in cell shape (Vanderberg, 1974; Münter et al., 2009). The force that drives the parasite forward is provided by the interaction of myosins (class XIV) with filamentous (F-) actin (Dobrowolski et al., 1997a; Meissner et al., 2002; Morrissette and Sibley, 2002; Siden-Kiamos et al., 2011). Although most Plasmodium actin seems to be in the monomeric, globular (G-) actin conformation (Field et al., 1993; Dobrowolski et al., 1997b), gliding motility apparently requires F-actin, as drug inhibition of F-actin dynamics translates into defective motility and impaired host cell invasion (Dobrowolski et al., 1997a; Münter et al., 2009; Hegge et al., 2010).

Until recently, F-actin could not be visualized in apicomplexan parasites under physiological conditions (Gantt et al., 2000; Kudryashev et al., 2010). This virtual absence of F-actin in highly motile parasites is supported by consistent in vitro polymerization assays using recombinant and parasite-derived Plasmodium actin (Schmitz et al., 2005; 2010,; Schüler et al., 2005; Sahoo et al., 2006; Skillman et al., 2011; 2013,). These studies revealed that actin1 forms exceptionally short and unstable filaments, which could only be detected in the presence of stabilizing drugs, such as phalloidin. The recent generation of conformation-specific anti-Plasmodium actin1 antibodies further supported the notion of a transient actin “ring” and actin rods in invading merozoites and gliding ookinetes respectively (Riglar et al., 2011; Wong et al., 2011; Siden-Kiamos et al., 2012). Similar short F-actin structures were identified in mutant Toxoplasma gondii tachyzoites, which were depleted of the G-actin-binding protein cofilin/actin depolymerizing factor (Mehta and Sibley, 2011).

Surprisingly, and in contrast to other eukaryotes, near-complete genome data sets (Gardner et al., 2002) suggested that the dynamics of these short actin filaments are regulated only by a minimal set of binding proteins (Baum et al., 2006; Schüler and Matuschewski, 2006; Sattler et al., 2011). Several classes of canonical actin regulators, including gelsolin, the Arp2/3 complex and WASP homology domain-containing proteins, are simply absent in Plasmodium genomes. One of the few conserved actin-binding proteins of Plasmodium parasites is the F-actin capping protein (CP), which is found in all eukaryotic organisms and metazoan cell types (Casella et al., 1987; Wear and Cooper, 2004; Cooper and Sept, 2008). In muscle cells, for instance, CP anchors actin filaments to the Z-disc, leading to the name CapZ (Casella et al., 1987; Schafer et al., 1993). CP binds in a calcium-independent manner to the fast-growing (barbed) ends of F-actin, thereby blocking subunit exchange (Cooper and Sept, 2008). Importantly, CP also belongs to the defined set of proteins that are needed to reconstitute actin-based motility in vitro (Loisel et al., 1999), and CP depletion results in reduced cell motility in vitro and in vivo (Hug et al., 1995; Sinnar et al., 2014). In addition to capping F-actin filaments, CP also stabilizes minifilaments composed of actin-related protein-1 (Arp1) (Cooper and Sept, 2008). In both cases, CP prevents addition as well as removal of actin protomers at the barbed end of the filaments. Active CP is composed of two subunits, CPα and CPβ (Cooper and Sept, 2008), and production of recombinant active CP in Escherichia coli is typically only achieved by co-expression of both subunits (Soeno et al., 1998), whereas recombinant expression of individual subunits was reported to produce insoluble protein (Remmert et al., 2000).

Plasmodium CPβ is encoded by a single open reading frame (Ganter et al., 2009), whereas CPα (PBANKA_124310 and PF3D7_0528500 for Plasmodium berghei and Plasmodium falciparum, respectively) is composed of nine small exons (Supporting Information Fig. S1A). Overall, Plasmodium CPα subunits share approximately 19% amino acid sequence identity with other eukaryotic CPα subunits, and 50–90% identity across different Plasmodium species (Supporting Information Fig. S1B and C). Most importantly, the residues that contribute to actin binding and heterodimer formation (Yamashita et al., 2003) are conserved (Supporting Information Fig. S1B).

We previously identified the CPβ subunit of the rodent malaria parasite P. berghei (PbCPβ) as an essential regulator of sporozoite motility and malaria transmission (Ganter et al., 2009). Deletion of PbCPβ did not influence asexual and sexual blood-stage development in the mammalian host. In the insect vector, Anopheles mosquitoes, mutant parasites displayed defective motility, which completely arrested life cycle progression at the sporozoite stage. Our study also established that recombinant P. berghei CPα/β heterodimers display capping activity on heterologous non-muscle actin (Ganter et al., 2009). The stage-specific function of CPβ in sporozoites implies that CPα alone might be functional during blood infection of cpβ(−) parasites. Given that independent functions of CP subunits have not been described, this notion was unexpected and prompted us to investigate the cellular role(s) of Plasmodium CPα for parasite life cycle progression.

Here, we show that the CPα subunit has a distinct and important in vivo role during Plasmodium blood infection, the exclusive cause of malaria-related pathology. We also show that recombinant PbCPα is stable in solution and displays actin filament capping activity in the absence of the partner β subunit.

Results

Expression profiling of Plasmodium CP

We first quantified CPα steady-state transcript levels by quantitative real-time PCR (qPCR) in three parasite stages: (i) asynchronous blood stages, (ii) schizonts and (iii) ookinetes, the stage that infects the Anopheles mosquito (Fig. 1A). We normalized transcript levels to actin1 (PBANKA_145930) and compared them with the class XIV myosin A (MyoA, PBANKA_135570) and the reporter GFP. Although expressed in all parasite stages (Ganter et al., 2009), levels of both CP subunits were at least 30-fold lower than MyoA or GFP, in accordance with the model that only a few CP units are needed to regulate actin filaments composed of multiple actin protomers (Cooper and Sept, 2008; Sattler et al., 2011). We also noted a minor increase of CPα, but not CPβ, in schizonts, which is then subdivided into multiple merozoites that invade new target cells (Fig. 1A). Our qPCR data are also supported by recent RNA-seq data (Otto et al., 2014; Supporting Information Table S1).

Fig 1

Plasmodium CPα subunit is soluble and binds to F-actin.A. Expression profiling of Plasmodium berghei CPα. Shown is a quantitative real-time PCR transcription analysis of P. berghei CPα, CPβ, myosin A (myoA) and the reporter protein GFP in mixed blood stages, schizonts and ookinetes. Relative mRNA levels were normalized to actin1 transcript levels and are shown as the mean of at least two independent experiments, each done in triplicate (± standard deviation).B. Recombinant expression and purification of soluble PbCPα protein. Shown is a Coomassie-stained SDS-PAGE of recombinant purified PbCPα protein.C. Size exclusion chromatography of recombinant PbCPα revealed an apparent molecular mass of ∼75 kDa (black). Molecular weight standards used for this analysis are shown as gray dots.D. PbCPα specifically co-sediments with actin polymers upon ultracentrifugation. Shown is Coomassie-stained SDS-PAGE using three different recombinant purified CPα proteins. Gelsolin, denoted with an asterisk, is included as positive control.

Recombinant Plasmodium CPα is stable and binds to F-actin in the absence of CPβ

Given the results, we hypothesized that E. coli-expressed PbCPα alone might be soluble in vitro. Indeed, PbCPα could be readily expressed and purified as soluble protein (Fig. 1B). When we determined the approximate molecular weight by size exclusion chromatography, we observed that the ∼37 kDa PbCPα protein did not elute as a monomer but at an apparent molecular weight of 82 kDa. This value is consistent with a homodimer of PbCPα (Fig. 1C). To confirm this result, we performed crosslinking experiments under increasing salt concentrations (Weiss et al., 2000). Monomers of pure recombinant CPα disappear in favor of the crosslinked putative dimer, which remains even at high salt concentration (Supporting Information Fig. S2). This finding permitted us to examine the biochemical activities of recombinant P. berghei CPα without its cognate β subunit.

To test whether recombinant PbCPα protein could interact with actin filaments, solutions of actin in the presence of three recombinant PbCPα proteins were subjected to ultracentrifugation, and the pellet and supernatant fractions were analyzed by SDS-PAGE (Fig. 1D). All three proteins were recovered in the F-actin-containing pellet fraction indicative of their ability to bind to and co-sediment with F-actin. A similar analysis using varying concentrations of PbCPα confirmed that presence of the protein in the pellet fraction depends on the presence of F-actin (Supporting Information Fig. S3). Canonical CPs, by interacting with the barbed end of F-actin, cause a shift of filament length distribution toward shorter polymers (Xu et al., 1999). This is illustrated by a visible shift of actin from the pellet to the supernatant fraction in the presence of gelsolin (Supporting Information Fig. S2). We did not observe such an effect with PbCPα, most likely because of the narrow regime of polymer lengths for which this effect can be detected by sedimentation velocity.

Recombinant Plasmodium CPα exhibits capping activity in vitro

As an alternative quantitative approach to test capping activity of recombinant PbCPα, we employed fluorescent microscopy of actin polymers (Xu et al., 1999). The presence of CPs shortens actin filaments by blocking the fast-growing (barbed) end. We determined the length distribution of fluorescently labeled actin polymers in presence or absence of recombinant PbCPα (Fig. 2A). Bovine gelsolin, which not only binds to the barbed end but also severs filaments, served as control (Harris and Weeds, 1984; Bearer, 1991; Sun et al., 1994). Upon addition of recombinant PbCPα to a fourfold molar excess of heterologous bovine actin, the median actin filament length significantly decreased from 8.5 μm to 4.1 μm (P < 0.0001). Gelsolin reduced the median filament length to 2.9 μm at a 47-fold molar excess of actin (P < 0.0001) (Fig. 2B).

Fig 2

PbCPα exhibits capping activity in vitro.A. Representative images of Alexa Fluor® 546 phalloidin-stained filaments of non-muscle bovine actin alone (Bt actin; upper panel), in the presence of recombinant Plasmodium berghei CPα (+ 55 nM PbCPα; middle panel) or in the presence of bovine gelsolin (+ 5 nM Bt gelsolin; lower panel); bars = 10 μm.B. Box plot of filament length distribution of bovine actin alone (Bt actin) or in presence of 55 nM PbCPα or 5 nM bovine gelsolin respectively. The length of filaments was determined by employing ImageJ software; polymers shorter than 0.5 μm and longer than 40 μm were omitted from the analysis (n = 1038 for Bt actin, 853 for Bt actin + 55 nM PbCPα and 820 for Bt actin + 5 nM Bt gelsolin). Boxes contain 50% of the data distributed around the median (bar); whiskers extend to the extreme observations within the 1st and 99th percentiles. Bonferroni-corrected P values were determined by Mann–Whitney test.C. Filament length distribution from B. Polymer lengths were sorted into 2 μm bins and the percental distribution plotted.

In the presence of PbCPα, the percentage of filaments between 20 and 40 μm substantially decreased (from 19% to 7%), while the percentage of filaments between 0.5 and 2 μm increased (from 20% to 31%). Upon addition of gelsolin, only 1% of filaments were 20–40 μm and 36% were 0.5–2 μm (Fig. 2C). Together, these findings show that PbCPα alone displays capping activity in vitro.

P. berghei CPα is refractory to targeted gene deletion

Viability of cpβ(−) parasites during blood infection in mice (Ganter et al., 2009) prompted us to design an experimental genetics strategy to select a similar cpα(−) mutant. To this end, we generated a targeting plasmid (pCPαREP) to replace the P. berghei CPα coding region with the positive selectable marker T. gondii DHFR/TS [Fig. 3A (i)]. In striking contrast to CPβ (Ganter et al., 2009), we failed to generate viable recombinant cpα(−) parasites in three independent transfection experiments, each conducted in duplicate (Supporting Information Fig. S4). Failure to select cpα(−) parasites could be either due to an essential function of PbCPα in asexual blood-stage parasites, the stage where transfection is performed, or refractoriness of the PbCPα gene locus to gene targeting.

Fig 3

Plasmodium berghei CPα is essential for asexual blood stages and can only be targeted in the presence of functional Plasmodium falciparum CPα.A. Replacement and complementation strategies to generate the potential cpα(−) and transgenic PfCPα parasites. The CPα ORF is targeted with a replacement vector that either contains (i) the selection cassette only, (ii) the orthologous P. falciparum (PfCPα) gene or (iii) a truncated version lacking the C-terminal 27 amino acids (PfCPαΔtail). Numbers indicate the length of the transgene in base pairs. Replacement-specific test primer combinations are indicated by arrows and expected fragments are shown as lines. Capped lines indicate the probe for diagnostic Southern blot; dotted capped lines indicate the expected genomic fragments upon XhoI (X) and SpeI (Sp) digest. Constructs were linearized using SacII (S) and KpnI (K) respectively.B. Genotyping of two clonal cpα(−)::PfCPα parasite lines (II1 and III1) obtained by in vivo cloning via limiting dilution. Genomic DNA from parental (par) and wild-type (WT) parasites serves as controls.C. Asexual blood stage development of cpα(−)::PfCPα II1 and III1, as well as WT parasites upon intravenous injection of 1000 infected red blood cells. Shown are mean parasitemias (± standard deviation) from groups of five mice per parasite line.

To distinguish between these two possibilities, we generated a targeting vector for trans-species complementation of the PbCPα gene, employing the ortholog from the human malaria parasite P. falciparum (PfCPα) [Fig. 3A (ii)]. Upon successful transfection, PfCPα is expressed under the control of the endogenous P. berghei promoter and the heterologous P. berghei DHFR/TS 3′ untranslated region. The first transfection of the functional P. falciparum CPα copy readily resulted in recombinant parasites, termed cpα(−)::PfCPα (Supporting Information Fig. S4), indicative of an essential function of CPα in Plasmodium blood infections.

We next asked whether the carboxy (C)-terminus of CPα, an important actin-binding site in the chicken ortholog (Narita et al., 2006; Takeda et al., 2010), also mediates vital functions in our experimental model. Truncation of the 28 C-terminal amino acids of chicken CPα led to a 5000-fold reduction in capping affinity (Wear et al., 2003). We, hence, generated a P. falciparum CPα replacement vector containing the corresponding C-terminal truncation (PfCPαΔtail) [Fig. 3A (iii)]. Two independent transfections of this targeting plasmid, each conducted in duplicate, did not result in recombinant parasites (Supporting Information Fig. S4). This result indicated that the presence of the predicted actin-binding motif is essential for CPα function in vivo.

Together, the experimental genetics data revealed that targeted gene deletion did not resemble the phenotype of cpβ(−) parasites (Ganter et al., 2009). This unexpected finding is strongly suggestive of independent vital role(s) of CPα during Plasmodium blood infection, where lack of CPβ is phenotypically silent.

Complete rescue of blood infection defects by P. falciparum CPα

To confirm and further analyze the P. falciparum CPα complementation, we generated two independent clonal parasite lines, termed cpα(−)::PfCPα II1 and III1, by limited in vivo dilution. Successful replacement of the endogenous PbCPα by the transgenic P. falciparum CPα was confirmed by diagnostic PCR in both lines (Fig. 3B). We tested whether the mutant parasites were able to proliferate normally during blood infection, utilizing wild-type (WT) parasites as controls (Fig. 3C). The blood stage parasitemia of both mutant parasite lines increased over time indistinguishably from WT parasites, confirming the successful trans-species complementation of CPα. This result also suggests that the vital CPα function(s) during blood infection are apparently conserved between the murine parasite P. berghei and the human pathogen P. falciparum. Together with our biochemical characterization of PbCPα and our previously reported successful generation of a cpβ(−) mutant parasite (Ganter et al., 2009), these genetic findings indicate that the α subunit can operate independently of the β subunit.

Combinatorial complementation reveals distinct roles for CP subunits during Plasmodium life cycle progression

In marked contrast to the observed normal blood stage growth, cpα(−)::PfCPα parasites displayed defects in colonization of the Anopheles midgut and a complete block of salivary gland invasion (Fig. 4). Because these defects are very similar to the phenotype of cpβ(−) parasites (Ganter et al., 2009), we generated a parasite line that harbors the P. falciparum CPβ in place of the endogenous CPβ (Supporting Information Fig. S5). Strikingly, these parasites remained defective in life cycle progression in the Anopheles vector as well (Fig. 4).

Fig 4

Plasmodium falciparum CPα/CPβ complementation.A. Scheme of the procedure to select a cpα(−)::PfCPα × cpβ(−)::PfCPβ double complemented parasite by meiotic recombination in the Anopheles vector and transmission to susceptible C57Bl/6 mice followed by pyrimethamine selection to counter-select for WT parasites and in vivo cloning to obtain a clonal parasite line. For details of sporozoite haplotypes, see Supporting Information Fig. S6A.B. Southern blot analysis to validate the correct genotype of the selected clonal cpα(−)::PfCPαX cpβ(−)::PfCPβ double complemented parasite line.C. Quantification of oocysts per mosquito midgut, infected with Plasmodium berghei WT (white), cpα(−)::PfCPα (green), cpβ(−)::PfCPβ (orange) and cpα(−)::PfCPα × cpβ(−)::PfCPβ (blue) parasites. Data are from a minimum of three independent feeding experiments. ***P < 0.0001 (Kruskal–Wallis test followed by Dunn's post-test).D. Quantification of midgut-associated sporozoites on day 14 after infection. Data are from a minimum of three independent feeding experiments. *P < 0.05; **P < 0.01 (Mann–Whitney test).E. Quantification of mature sporozoites that have colonized mosquito salivary glands on day 17 after infection. Data are from a minimum of three independent feeding experiments. *P < 0.05; **P < 0.01 (Mann–Whitney test).

We hypothesized that for these stage-specific tasks, a functional CP heterodimer is necessary and that P. berghei and P. falciparum subunits are incompatible. To test this hypothesis experimentally, we generated a double complemented parasite line, cpα(−)::PfCPα × cpβ(−)::PfCPβ, by cross-fertilization and chromosomal recombination during meiosis (Fig. 4A and Supporting Information Fig. S6A). Selection of the rare recombination event, due to proximity of the two CP loci on chromosome XII, was augmented by counter-selection of recombined WT parasites with the antimalarial drug pyrimethamine. Natural transmission to susceptible C57Bl/6 mice and in vivo cloning of the resulting blood stage parasites yielded two clones, the desired cpα(−)::PfCPα × cpβ(−)::PfCPβ line and cpβ(−)::PfCPβ parasites (Fig. 4A). The latter is in good agreement with the previous finding that defective transmission of cpβ(−) parasites can be compensated for by presence of a WT allele during sporogony (Ganter et al., 2009). Southern blot analysis (Fig. 4B) and diagnostic PCR (Supporting Information Fig. S6B) confirmed successful generation of a double complemented parasite line by this strategy, already indicating that these parasites can complete the life cycle successfully.

Phenotyping of cpα(−)::PfCPα × cpβ(−)::PfCPβ parasites in the Anopheles vector revealed significantly more oocysts than in infections with parasites that contain single subunit complementations (Fig. 4C), indicative of partial recovery of the midgut colonization defects. Because of density dependence of oocyst-sporozoite transition (Sinden et al., 2007), the differences were compensated for in the production of oocyst sporozoites (Fig. 4D). Most importantly, we recovered sporozoites from salivary glands of cpα(−)::PfCPα × cpβ(−)::PfCPβ- infected Anopheles mosquitoes (Fig. 4E), corroborating partial recovery of the complete defect detected in infections with single subunit complementations. We interpret the finding that trans-species complementation of both subunits is needed to rescue the severe sporozoite defect as an indication that a CP heterodimer is required in mosquito stages, while CPα alone exerts vital functions during blood stage propagation.

Discussion

The most important finding of our study is that, at least in the parasitic protist Plasmodium, CPα exerts important functions in the absence of its cognate β subunit. This unexpected claim is supported by two complementary findings: (i) the CPα subunit of the malarial parasite has likely vital in vivo role(s) during asexual blood stage replication, which is not the case in cpβ(−) parasites, and (ii) recombinant PbCPα is functional in absence of CPβ. Ablation of CPα is incompatible with in vivo proliferation of asexual blood stage parasites, whereas CPβ is dispensable for parasite propagation in the mammalian host (Ganter et al., 2009). CPα is the only remaining subunit in the latter mutant, indicating that the yet unknown vital function of CPα in asexual blood stages is likely performed by CPα alone, perhaps as a homodimer. Together, our reverse genetics analysis of P. berghei CP suggests that both subunits perform multiple, independent tasks.

We produced recombinant P. berghei CPα subunit, for the first time in the absence of PbCPβ. The recombinant PbCPα protein displayed capping activity on heterologous non-muscle actin. Inspection of the crystal structure of the chicken CPα/β heterodimer along with sequence comparison also suggests that the existence of a PbCPα/α homodimer is more likely than that of a PbCPα monomer (Yamashita et al., 2003): Taking the subunits out of their dimer context would expose the dimerization interface, creating a large unfavorable solvent-accessible hydrophobic surface. However, we cannot formally exclude the existence of monomeric pools of PbCPα in vivo, possibly stabilized by a protein-binding partner.

When analyzing residues that make important contributions to the chicken α/β heterodimer (Yamashita et al., 2003), it is apparent that in most cases both bond donors and acceptors are conserved in their respective positions in PbCPα (and, in fact, in GgCPα). For instance, chicken CPα R259 is involved in a salt cluster with CPβ Y107, E221 and N222; all four positions are strictly conserved in the PbCPα sequence (R281, Y116, E258, N259; Supporting Information Fig. S1B). This suggests the intriguing possibility of a PbCPα/α homodimer that uses the same dimerization interface as the CPα/β heterodimer and a mechanism of regulation that is based on competition of available subunits, e.g. by transcriptional regulation. Based on our results, we cannot rule out PbCPα homodimers with a dimer interface that differs from the one in the heterodimer; these remain important topics for future studies. Crystals of PbCPα could not be produced to a quality sufficient for structure determination. Thus, the structure of Plasmodium CP to atomic detail remains elusive, as does the function of the 23-residue insertion. To the best of our knowledge, no evidence for CPα being active as individual subunits has been published yet, whereas a large body of literature demonstrates that CP acts as α/β heterodimers (Cooper and Sept, 2008; Pollard and Cooper, 2009).

CP stabilizes actin filaments, as well as Arp1 minifilaments, by preventing both gain and loss of monomers at the filament barbed end (Cooper and Sept, 2008; Pollard and Cooper, 2009). For this capping activity, the C-terminal tails of each subunit are crucial (Narita et al., 2006; Narita and Maéda, 2007). Basic residues of the CPα C-terminus initially interact with acidic amino acids of the penultimate and terminal actin protomers; the mobile CPβ C-terminus then occupies a hydrophobic pocket on the terminal protomer (Narita et al., 2006; Takeda et al., 2010). This two-step binding model assigns a central role to the CPα C-terminus, and deletion of CPα tail led to a 5000-fold decrease in capping affinity, in contrast to a 300-fold reduction upon deletion of the CPβ C-terminus (Wear et al., 2003). We note that recombinant PbCPα protein lacking the eight most C-terminal residues had actin capping activity and co-sedimented with actin polymers in ultracentrifugation experiments. There are several possible explanations for this. This function may not be conserved in the C-terminus of the parasite CP protein or may not come to bearing in the interaction with heterologous actin. These explanations would also be consistent with the high concentrations of PbCPα, as compared with chicken CP (Wear et al., 2003), needed to detect an effect. Alternatively, experiments at higher resolution might be required to detect a similar effect of the PbCPα C-terminal extension.

Capping activity appears to be essential throughout the Plasmodium life cycle. Intriguingly, in apicomplexan parasites, the two CP subunits apparently split the task of F-actin and/or Arp1 minifilament capping, as the parasite changes its cellular environment. One possible explanation is that the CPα/β heterodimer largely mediates microfilament capping at ambient temperatures in the mosquito vector, whereas the α homodimer might have evolved for the corresponding activity in the warm-blooded mammalian host. Alternatively, considering the divergent functional properties of parasite actin, F-actin and Arp1 minifilament capping may be divided between the α and β subunits. In such a scenario, the fast-moving sporozoites might rely on F-actin capping by CPα/β, while an intracellular lifestyle and host cell manipulation in general might require efficient vesicle uptake that involves the dynactin complex together with a CPα-capped Arp1 minifilament.

Birds and mammals express at least three isoforms of CPα and two isoforms of CPβ, and all display tissue-specific expression patterns (Barron-Casella et al., 1995; Hart et al., 1997; Hart and Cooper, 1999). Similarly, we identified PbCPβ initially as a gene that is up-regulated during sporozoite maturation (Matuschewski et al., 2002). While we present the first description of separate cellular roles for CP subunits, differential and tissue-specific heterodimers as a result of functional requirements appear to be established in other organisms as well. For instance, in mouse muscle cells, the two CPβ isoforms perform distinct in vivo functions that cannot be compensated for by the other isoform (Hart and Cooper, 1999). Together with our findings, this warrants future studies to test whether CP subunits alone exert specific functions in other eukaryotes, as seen in Plasmodium with its morphologically and phenotypically distinct life cycle stages.

Experimental procedures

Experimental animals

Naval Medical Research Institute (NMRI) mice were purchased from Charles River Laboratories, Sulzfeld, Germany. All animal work was conducted in accordance with the German “Tierschutzgesetz in der Fassung vom 18. Mai 2006 (BGBl. I S. 1207)”, which implements directive 86/609/EEC from the European Union and the European Convention for the Protection of Vertebrate Animals used for experimental and other scientific purposes. The protocol was approved by the state authorities (LAGeSo Reg# G0469/09).

Reverse transcription and qPCR

Total RNA was purified employing the RNeasy kit (Qiagen), and reverse transcription was performed using the RETROscript kit (Ambion). qPCR was performed on cDNA preparations from mixed blood stages, purified late schizonts/merozoites and purified ookinetes using the ABI 7500 sequence detection system or StepOnePlus™ and Power SYBR® Gene PCR Master Mix (Applied Biosystems), according to the manufacturer's instructions. qPCR was performed in triplicates, with 1 cycle of 95°C for 15 min, followed by 40 cycles of 95°C for 15 s, 55°C for 15 s and 60°C for 45 s. Data were analyzed with the SDS1.3.1 software (Applied Biosystems). Relative transcript abundance was normalized to the expression of actin. The following primers were used for qPCR: CPα_for and CPα_rev; CPβ_for and CPβ_rev; MyoA_for and MyoA_rev; actin1_for and actin1_rev; and GFP_for and GFP_rev. Primer sequences are listed in Supporting Information Table S1.

Recombinant expression and purification of CPα

DNA encoding the CPα fragments PbCPα4–307 (L4-V307), PbCPα4–299 (L4-D299) and PbCPα4–292 (L4-Y292) were subcloned into pNIC-Bsa4 to add an N-terminal cleavable hexa-histidine tag. All proteins were expressed in E. coli BL21-CodonPlus cells (Stratagene) and purified using immobilized nickel ion affinity chromatography, followed by size exclusion chromatography in 20 mM HEPES (pH 7.5), 300 mM NaCl, 10% glycerol and 1 mM TCEP. The three highly similar purified proteins were used for biochemical assays as follows: PbCPα4–307 for cosedimentation with actin polymers (Supporting Information Fig. S3); PbCPα4–299 to represent expression of soluble protein (Fig. 1B), for cosedimentation with actin polymers (Fig. 1D), for capping activity of fluorescently labeled actin polymers (Fig. 2) and for the crosslinking assay (Supporting Information Fig. S2); and PbCPα4–292 for size exclusion chromatography (Fig. 1C). Two additional proteins containing a C-terminal hexa-histidine tag, PbCPα1–307 (M1-V307) and PbCPα1–299 (M1-D299), were included in the cosedimentation with actin polymers (Fig. 1D). All recombinant protein batches were verified using time-of-flight mass spectrometry analysis (data not shown).

Apparent molecular mass determination

Analytical size exclusion chromatography was carried out using a 16/10 Superdex-200 column attached to an Äkta Explorer FPLC system (GE Healthcare) and buffer containing 20 mM HEPES (pH 7.5), 300 mM NaCl, 10% glycerol and 1 mM TCEP. The column was calibrated using molecular mass standards, i.e. aprotinin, ribonuclease A, carbonic anhydrase, ovalbumin and conalbumin (GE Healthcare); as well as adenosylhomocysteinase-3 log(MW) was plotted against KAV = (V0-Vt)/(V0-Ve) for each protein, where V0 is the total column volume, Ve is the exclusion volume and Vt is the elution volume of each protein. The apparent molecular mass of PbCPα was calculated from its elution volume and the regression line of the above plot.

Actin assays

The ultracentrifugation assay was conducted as previously described (Schüler and Peti, 2007) using 4 μM non-muscle bovine β-actin, PbCPα and 200 nM bovine gelsolin (Sigma-Aldrich G8032). The microscopic analysis of capping activity of recombinant PbCPα was carried out as described (Xu et al., 1999). In brief, β-actin in F-actin buffer (20 mM Tris pH 7.5, 1 mM MgCl2, 100 mM KCl, 1 mM TCEP, 0.1 mM CaCl2, 0.5 mM ATP, 0.1 mg ml−1 BSA) was incubated at room temperature for 2–3 h in the presence of 175 nM Alexa Fluor® 546 phalloidin (Invitrogen). Prior to microscopy, phalloidin-stained F-actin was diluted 1:4 in F-actin buffer either without additional protein or with 55 nM PbCPα or 5 nM bovine gelsolin. Of these solutions, 3 μl was transferred onto 12 mm diameter poly-L-lysine-coated cover slips and subsequently imaged. To minimize filament severing due to shear forces, we used trimmed pipette tips at all times. Filaments were imaged on a Leica TCS SP-1 confocal microscope, and their length was determined using ImageJ software (U. S. Natl. Institutes of Health).

Parasite transfection and genotypic analysis

For targeted disruption of PbCPα, two fragments were amplified from P. berghei genomic DNA as template using primers PbCPα_forI and PbCPα_revII to amplify the 5′ flanking region, and PbCPα_forIII and PbCPα_revIV for amplification of the 3′ flanking region. Cloning into the P. berghei transfection plasmid b3D.DT∧H.∧D (Janse et al., 2006) resulted in the plasmid pPbCPαrep. The targeting plasmid was linearized with SacII and KpnI; parasite transfection, positive selection and parasite cloning were then performed as previously described (Janse et al., 2006). Integration-specific PCR amplification of the predicted cpα(−) locus was done using specific primer combinations: CPα_5′_test_for and b3D_revX, as well as b3D Tg_rev_Pro and CPα_3′ test_rev. Primer sequences are listed in Supporting Information Table S2.

Single complementations of cpα(−) and cpβ(−)

In order to complement cpα(−) parasites, we amplified the orthologous P. falciparum CPα gene using the primers PfCPα_compforV and PfCPα_comprevVI and P. falciparum cDNA as template (Supporting Information Table S2). Cloning into the plasmid pPbCPαrep resulted in the complementation plasmid pPfCPα. For complementation with a C-terminally deleted P. falciparum CPα gene that lacks the last 27 amino acid residues, primers PfCPα_compforV and PfCPα_comprevVII were used, resulting in the complementation plasmid pPfCPαΔtail. Transfection and genotyping were conducted as described above; parasites were cloned by limiting dilution and intravenous injection of a single parasite per naïve NMRI mouse in a total of 10 animals.

To complement cpβ(−) parasites, we amplified the orthologous P. falciparum CPβ gene (PF3D7_0517600) using the primers PfCPβ_compforV and PfCPβ_comprevVI and P. falciparum genomic DNA as template (Supporting Information Table S2). Cloning into the plasmid pPbCPβREP (Ganter et al., 2009) resulted in the plasmid pPfCPβ with PfCPβ under the control of the endogenous PbCPβ promoter and the 3′ untranslated region of PbDHFR/TS. Transfection and genotyping were conducted as described above; parasites were cloned by limiting dilution and intravenous injection of a single parasite per naïve NMRI mouse in a total of 10 animals. cpβ(−)::PfCPβ clones were genotyped by diagnostic PCR using primers CPβ_5′_test_for and PfCPβ_comprevVII as well as b3D Tg_rev_Pro and CPα_3′ test_rev (Supporting Information Table S2).

Double complementation of cpα(−) and cpβ(−)

cpα(−)::PfCPα × cpβ(−)::PfCPβ double complemented parasites were obtained by genetic cross of cpα(−)::PfCPα and cpβ(−)::PfCPβ clones. We fed clonal single complemented parasites to Anopheles stephensi mosquitoes. Upon sexual recombination and crossover, we expected to obtain a fraction of sporozoites with the desired double complementation. C57Bl/6 mice were infected by natural transmission and a clonal cpα(−)::PfCPα × cpβ(−)::PfCPβ double complemented line was isolated by limiting dilution. The genotype was confirmed by Southern blot as previously described (Ganter et al., 2009).