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Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

We have utilized a highly sensitive approach based on fluorescence resonance energy transfer (FRET) and β-lactamase (BLA), which we adapted for the detection of Toxoplasma gondii secreted proteins. This assay revealed that the actin-binding protein toxofilin appears to be secreted into host cells during invasion. To determine the function of toxofilin during infection, we engineered a type I (RH strain) parasite with a targeted deletion of the toxofilin gene and compared the phenotypes of control and toxofilin knockout (Δtxf) parasites in several in vitro assays, including invasion, growth, gliding motility, and egress of the Δtxf parasites, as well as F-actin staining, phagocytosis and migration of cells infected with Δtxf parasites or wild-type controls. Despite its apparent secretion into host cells and its ability to bind to and modulate host actin, we observed that toxofilin does not appear to play a role in these processes, under the conditions we examined, and we report these findings here.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Toxoplasma gondii is a protozoan parasite of the phylum Apicomplexa, a group of globally distributed pathogens that includes the causative agents of malaria (Plasmodium), gastrointestinal infections (Cryptosporidium) and coccidiosis (Eimeria). These obligate intracellular parasites are characterized by the presence of specialized secretory organelles, called micronemes, rhoptries and dense granules, at their apical ends. As the parasite invades a host cell, a subset of rhoptry and dense granule proteins are secreted into the host cell and/or parasitophorous vacuole (PV). Some of these secreted effectors are known to contribute to parasite virulence by trafficking to distinct intracellular compartments, altering gene transcription and modifying cytokine production (Saeij et al., 2006; 2007; Taylor et al., 2006). Ultimately, these processes allow the parasite to establish its intracellular niche.

Biochemical isolation and proteomic analysis of the rhoptries have identified a large number of additional proteins that localize to this organelle: 29 proteins have been confirmed as rhoptry proteins by immunofluorescence assays (IFA), and an additional 28 proteins have been isolated from the rhoptries and await confirmation of their localization (Bradley et al., 2005). Based on sequence-prediction analysis, the rhoptry proteins are comprised of putative kinases, phosphatases and proteases, as well as proteins without known motifs or homologies. Understanding the mechanisms by which these proteins function in the host cell will shed light on the molecular interactions between parasite and host cell factors that underpin the intracellular life of the parasite.

One protein found in the rhoptry proteome, called toxofilin, was initially identified by its ability to bind to parasite and mammalian actin and to modulate actin dynamics in vitro (Poupel et al., 2000; Delorme et al., 2003). Recently, toxofilin was crystallized in a complex with mammalian actin (Lee et al., 2007), confirming the potential for a direct interaction between the parasite and host proteins. Because toxofilin has a predicted signal peptide and localizes to a secretory organelle (Bradley et al., 2005), we hypothesized that toxofilin may be secreted into the host cell cytosol during parasite invasion, where it might then interact with and somehow modulate host actin function.

A limitation of the examination of T. gondii-secreted proteins has been the ability to detect them in host cells. A standard method for visualizing secreted proteins, which involves antibody staining and IFA, requires that the protein of interest be secreted in high enough quantities or be concentrated on a membrane or in an intracellular compartment for visualization. Soluble or low-abundance proteins that are secreted by the parasite into the host cell cytoplasm, however, may be undetectable using these assays, due to the large volume of the host cell into which they diffuse. As described below, this appears to be the case for toxofilin. In order to examine T. gondii secretion of low-abundance proteins, we have utilized a highly sensitive assay based on fluorescence resonance energy transfer (FRET) and β-lactamase (BLA) (Zlokarnik et al., 1998; Marketon et al., 2005). Using this approach, we show here that toxofilin does appear to be secreted into host cells during invasion. We have investigated the function of toxofilin by engineering knockout (Δtxf) parasites. Using a variety of assays, we show that the absence of this protein has surprisingly little impact on the growth of T. gondii in vitro.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

The toxofilin–BLA fusion protein is secreted by the parasite

Toxofilin is a 26.7 kDa protein that localizes to the rhoptries (Poupel et al., 2000). Given its predicted signal peptide and localization to a secretory organelle, we hypothesized that toxofilin is secreted into host cells during invasion. We attempted to detect endogenous toxofilin and a haemagglutinin (HA) epitope-tagged form of toxofilin in infected host cells by fluorescence microscopy imaging using anti-toxofilin or anti-HA antibodies respectively. Although these antibodies strongly stained toxofilin in the parasite, we were unable to visualize toxofilin in the cytoplasm of infected host cells (data not shown). Considering the limited quantities of rhoptry proteins that are secreted during invasion and, unless concentrated in a particular location, the enormous dilution of such proteins within the host cell cytosol, we supposed that the amount of toxofilin released into the host cell might be below the threshold of detection of antibody staining and microscopy. To circumvent this issue, we utilized a highly sensitive FRET-based BLA assay for the detection of secreted proteins (Zlokarnik et al., 1998; Marketon et al., 2005). The assay detects the presence of BLA fusion proteins in host cells treated with a BLA substrate linking two fluorescent molecules: coumarin cephalosporin fluorescein (CCF2). The substrate is administered to host cells in an esterified form with an acetoxymethyl group (CCF2-AM), and cleavage of this group by host esterases results in retention of the substrate in the host cell cytosol. When the substrate is intact and the fluorophores are in close proximity, excitation of the coumarin at 407 nm results in FRET to the fluorescein and emission at 520 nm, a green fluorescent signal. The presence of BLA, however, results in cleavage of the substrate and dissociation of the fluorophores, such that excitation of the coumarin at 407 nm results in emission at 447 nm, a blue fluorescent signal. A change in fluorescence from green to blue therefore indicates the presence of BLA in host cells treated with the BLA substrate, CCF2-AM.

To employ the BLA assay for T. gondii infection, we generated transgenic parasites in the type I (RHΔhpt) background expressing toxofilin fused to the HA epitope tag and BLA (named toxofilin–HA–BLA). We also generated control parasites expressing toxofilin–HA without the fusion to BLA. Examination of the resulting transgenic parasite clones by IFA demonstrated that both toxofilin–HA and toxofilin–HA–BLA localized predominantly to the rhoptries. Importantly, we did observe a low level of anti-HA staining detectable in the cytosol of some transgenic parasites, which is consistent with a previously reported localization of toxofilin (Poupel et al., 2000). The potential impact of this cytosolic localization is discussed further below. Western blot analysis of lysates from the transgenic parasites demonstrated that the toxofilin–HA and toxofilin–HA–BLA fusion proteins were of the predicted size (∼27 and ∼53 kDa respectively), and that the fusion proteins were expressed at very similar levels to endogenous toxofilin (Fig. 1B).

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Figure 1. Toxofilin–BLA is secreted by the parasite. A. HFF were infected with toxofilin–HA (clone 7.7) or toxofilin–HA–BLA (clone 3.3) for 30 min, fixed and stained with anti-ROP2/4 to detect the rhoptries or with anti-HA to detect the fusion protein. B. Lysates were generated from HFF cells infected with RHΔhpt, toxofilin–HA (clone 7.7) or toxofilin–HA–BLA (clone 3.3) for 24 h and separated by SDS-PAGE. The membranes were blotted with anti-HA, anti-toxofilin or anti-SAG1 antibodies. C. HFF were infected with RHΔhpt, toxofilin–HA (clone 7.7) or toxofilin–HA–BLA (clone 3.3) for 1 h and then loaded with CCF2-AM for 1 h. The live cells were imaged using a Leica SP2 AOBS Confocal Laser Scanning Microscope with a blue diode 405 nm laser for excitation and with detection filters set at 410–450 nm for coumarin and 493–550 nm for fluorescein. White arrowheads indicate the location of intracellular parasites, and red arrowheads indicate a few of the many extracellular parasites.

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To examine the secretion of toxofilin in infected cells, the parasites expressing toxofilin–BLA fusion proteins were then used in the in vitro BLA assay. Human foreskin fibroblasts (HFF) were infected with the transgenic parasites for 1 h, exposed to CCF2-AM for an additional hour, and subsequently visualized by fluorescence microscopy and by flow cytometry for blue or green fluorescent signal. We observed that uninfected cells, cells infected with the parental RHΔhpt parasites, or cells infected with parasites expressing toxofilin–HA remained green after treatment with CCF2-AM (Fig. 1C). HFF infected with parasites expressing the toxofilin–HA–BLA fusion, however, were blue, indicating the presence of BLA in the PV space and/or host cell cytosol and cleavage of the CCF2 substrate (Fig. 1C). Importantly, uninfected HFF in the same culture remained green. Moreover, intracellular parasites, designated by the white arrowheads, were not fluorescent and appeared dark in infected host cells, indicating that the substrate does not reach parasites that are inside the host cell. This would be expected, since the removal of the acetoxymethyl modification on CCF2-AM by host esterases would prevent further diffusion across the parasite membrane. It does appear though that CCF2 can cross into the vacuole (data not shown), and this is discussed below in more detail.

Since the toxofilin–HA–BLA fusion protein appeared by Western blotting to be intact (Fig. 1B), it appears that toxofilin–HA–BLA, and not simply BLA alone, is secreted into the host cells and/or PV space at some point during or soon after invasion. We have added the substrate CCF2-AM at 30 min post infection, but because substrate loading requires an additional 30 min, the earliest time we have examined is 1 h post infection. We detected the presence of toxofilin–HA–BLA in host cells at this early time point and when adding the substrate as late as 32 h post infection (data not shown), beyond which time lysis of the host cells precluded further analysis. Toxofilin–HA–BLA secreted during a single invasion event was sufficient to cause cleavage of the substrate, as we observed blue cells containing a single intracellular parasite.

Interestingly, most of the extracellular toxofilin–HA–BLA parasites in the above experiment showed blue fluorescence (Fig. 1C, red arrowheads), indicating that in addition to diffusing into host cells, the substrate is taken up by extracellular parasites and cleaved. This would not be expected if the fusion protein is exclusively located within the rhoptries, as the CCF2-AM substrate should be de-esterified upon entry into the parasites and thus unable to cross into the rhoptries to be cleaved. The blue signal may result from the fact that some of the toxofilin–HA–BLA appears to be in the parasite cytosol, where it should have easy access to de-esterified CCF2. We cannot exclude, however, the possibility that CCF2 may also access toxofilin in the rhoptries.

Detection of toxofilin in host cells requires parasite secretion but not invasion

To confirm that the cleavage of CCF2 within the host cells was due to secretion from the rhoptries, we pre-treated parasites with a chemical inhibitor of invasion and examined whether toxofilin–HA–BLA was still introduced into host cells. Cytochalasin D, an actin depolymerizating agent, can be used to pre-treat parasites prior to contact with host cells. Because the parasite actin-myosin machinery is required for invasion, but not for attachment to host cells, cytochalasin D pre-treatment allows parasites to attach to host cells and to secrete their rhoptry contents, but not to invade (Hakansson et al., 2001). Treatment of toxofilin–HA–BLA parasites with cytochalasin D before infection, followed by CCF2-AM addition to the culture, resulted in cleavage of the BLA substrate and blue host cells (Fig. 2). These data suggest that parasite secretion, but not invasion, is required for the translocation of toxofilin–HA–BLA into the host cell.

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Figure 2. Secretion of toxofilin requires active rhoptry secretion but does not require parasite invasion. HFF cells were infected with RHΔhpt or toxofilin–HA–BLA (clone 3.3) for 1 h followed by loading with CCF2-AM for 1 h. The cells were then washed, trypsinized and examined with a 407 nm krypton laser on a modified FACStar flow cytometer for side scatter (SSC) and for the detection of coumarin (in the cascade blue channel).

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To address the possibility that the detection of BLA in host cells was due to leakage of toxofilin–BLA from the parasite into the host cell, rather than rhoptry secretion, we generated transgenic parasites expressing a mutant form of toxofilin–HA–BLA that lacks a secretory signal peptide [named toxofilin(-SP)-HA–BLA] and localizes to the parasite cytoplasm. The expression level of the mutated toxofilin fusion protein in independent clonal lines was about 10% of the expression of the full-length toxofilin–HA–BLA, presumably due to toxicity issues associated with having large amounts of actin-binding protein in the parasite cytosol. We also generated parasites expressing cytosolic HA–BLA, without a signal peptide or any toxofilin coding sequence. In host cells infected with these two negative control parasites and treated with the BLA substrate, the cells remain green, even after infection with a 10-fold higher multiplicity of infection than used for parasites expressing the original, secreted version of toxofilin–HA–BLA (Fig. S1), and even after exposing the infected cells to the substrate at 24 h post infection (data not shown). Collectively, these findings indicate that toxofilin–HA–BLA appears to be introduced into the host cell via active secretion, rather than by passive leakage of cytosolic material.

Toxofilin knockout parasites exhibit normal growth, invasion, gliding motility and egress

Since toxofilin appears to be secreted into host cells and was initially characterized by its ability to bind to actin and modulate actin dynamics, its function during the infection of host cells was examined. We engineered a parasite line lacking the toxofilin gene (Δtxf) by using homologous recombination in the type I RH strain (RHΔhpt) background (Fig. 3A). The resulting Δtxf parasites, which are GFP+, were deficient in the toxofilin gene, as determined by PCR with primers for the toxofilin open reading frame (ORF) (Fig. 3B). We confirmed that the toxofilin protein was absent from the Δtxf parasites by staining the parental RH strain parasites and the Δtxf parasites with an anti-toxofilin antibody, and there was an absence of staining in the Δtxf parasites (Fig. 3C).

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Figure 3. Δtxf parasites are deficient in toxofilin and lack toxofilin protein. A. A schematic representation of the strategy for replacing the toxofilin gene with the selectable marker HXGPRT. The 5′- and 3′-targeting sequences were each ∼2 kb of genomic DNA from just upstream and just downstream of the toxofilin ORF. The promoters are represented by the bent arrows. B. Genomic DNA was generated from HFF cells infected with RHΔhpt or Δtxf parasites, amplified with primers for the toxofilin ORF or the SAG1 ORF, and separated by agarose gel electrophoresis. C. HFF cells infected with RHΔhpt or Δtxf parasites were fixed and permeabilized with 100% ethanol to quench the GFP expression in the parasites. They were subsequently stained with anti-SAG1 or anti-toxofilin antibodies.

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The role of toxofilin in parasite invasion of host cells and in intracellular growth was assessed by comparing the Δtxf and control parasites in human fibroblasts. To examine parasite invasion, HFF were infected with Δtxf or control parasites in a synchronized invasion assay using a potassium buffer shift (Kafsack et al., 2004). Parasite invasion of HFF cells was allowed to proceed for 30–180 s. Figure 4A depicts the per cent of intracellular, invaded parasites at each time point. Invasion of Δtxf parasites appeared normal when compared with control parasites. Similarly, no difference in invasion efficiency was observed when Δtxf and control parasites were compared using temperature shift to synchronize invasion (data not shown).

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Figure 4. Δtxf parasites have similar invasion efficiency and in vitro growth as control parasites. A. The invasion of Δtxf or control parasites in HFF was synchronized by treatment with high potassium buffer and a shift to DMEM containing lower potassium for the indicated time periods to allow parasite invasion. As the Δtxf and control parasites are GFP+, the invasion efficiency was scored by counting the number of intracellular parasites (GFP+SAG1-) relative to the total number of parasites in each field. This experiment was performed six times with comparable results; a representative experiment is shown. The error bars represent the standard deviation of intracellular parasites in 10 independent fields. B. HFF were infected with Δtxf or control parasites for 2 h and washed to remove uninvaded parasites. Twenty-four hours later the cells were fixed and the number of parasites per vacuole in eight fields was counted and expressed as a percentage of the total number of parasites in the field. The error bars represent the standard deviation from four biological replicate wells. This experiment was performed three times as a time-course, and the data shown are a representative result. C. HFF were infected with control or Δtxf parasites, and at 30 h post infection, the calcium ionophore A23187 was added to the cells for 15, 30, 60 or 90 s. For each time point, the number of lysed vacuoles was counted in five fields and expressed as a percentage of the total number of vacuoles in the field. This experiment was performed five times on different days, and all of the data are shown. An adjusted Wald F-test was used to calculate the P-values (= 0.3 at 15 s, P = 0.74 at 30 s, P = 0.41 at 60 s, P = 0.54 at 90 s).

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To assess intracellular parasite growth, HFF were infected with Δtxf or control parasites for 26 h, the cells were fixed, and the number of parasites per vacuole was counted. Figure 4B shows that the Δtxf and control parasites formed vacuoles containing similar numbers of parasites, indicating no difference in their ability to replicate in vitro. These results were also observed at 32 and 40 h post infection (data not shown). Since toxofilin was initially reported to be capable of interacting in vitro with both host and parasite actin, and since there appears to be a small amount of toxofilin in the parasite cytosol, we examined the role of toxofilin in parasite gliding motility. As parasites glide, they deposit proteins, such as SAG1, as a trail behind the gliding parasite. The control and Δtxf parasites both formed SAG1 trails when gliding on glass coverslips, and the trails were of equivalent length and number (data not shown), suggesting that a lack of toxofilin in the parasite does not impact parasite gliding motility, at least as determined by the presence and quality of SAG1 trails. Egress from host cells is another process that is integral to the parasite life cycle and that requires parasite motility. To determine if toxofilin plays a role in parasite egress, we infected HFF with the Δtxf or control parasites and induced egress by treating the infected cells with the calcium ionophore A23187 (Endo et al., 1982). We observed that the Δtxf parasites egressed slightly faster than the control parasites at the 15 s time point, but over five independent experiments, this slight acceleration was not statistically significant (Fig. 4C).

Toxofilin does not impact macrophage phagocytosis or migration

Since toxofilin appears to be secreted into host cells, but does not seem to impact parasite invasion, growth or gliding motility in vitro, we examined whether toxofilin affects host cell processes that require actin cytoskeleton remodelling. Macrophages are phagocytic cells that play an important role in innate immunity to T. gondii infection, both as a reservoir for the parasite and as an early inducer of cytokine production and activation of cellular immunity (Serbina et al., 2008). One important function of macrophages is their ability to take up or phagocytose infectious particles and apoptotic cells, a process that requires the macrophage actin cytoskeleton. To determine if the presence of toxofilin in infected macrophages impacts macrophage phagocytosis, we infected mouse bone marrow-derived macrophages (BMdM) with control or Δtxf parasites, and then measured their ability to phagocytose rhodamine-labelled Escherichia coli. As shown in Fig. 5, macrophages infected with either control or Δtxf parasites were able to take up E. coli in equivalent numbers, as indicated by the per cent of infected (GFP+) cells that are also positive for rhodamine. These data indicate that the presence or absence of toxofilin in infected macrophages does not seem to affect macrophage phagocytic ability.

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Figure 5. Mouse bone marrow-derived macrophages (BMdM) infected with Δtxf or control parasites have similar phagocytic ability. BMdM were infected with control or Δtxf parasites for 2 h and exposed to rhodamine-labelled (‘pHrodo’) E. coli Bioparticles at 37°C for 20 min. The cells were washed, removed from the plate by scraping, fixed, and examined on a modified FACStar flow cytometer to detect GFP and rhodamine. The numbers on each plot represent the percentage of cells that fall within the GFP+ gate, indicating infection efficiency. The histogram plots depict the rhodamine intensity of the GFP+ macrophage populations infected with the control (black lines) or Δtxf (grey lines) parasites.

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Cell migration is another host cell process that relies on actin cytoskeleton dynamics. To determine whether the secretion of toxofilin into host cells during parasite invasion impacts the ability of infected cells to migrate, we performed macrophage chemotaxis assays in transwell plates. BMdM were infected with Δtxf or control parasites in the upper chamber of a transwell plate for 2 h, and then conditioned medium from T. gondii-infected 3T3 cells was added to the lower chamber for 3 h. The migration of infected BMdM was assessed by scoring the percentage of cells that had migrated from the top to the bottom side of the filter separating the upper and lower chambers. As shown in Fig. 6, BMdM infected with the Δtxf or control parasites displayed a similar level of migration in this assay. Thus, toxofilin does not appear to play a role in migration of BMdM towards conditioned medium from infected cells.

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Figure 6. Toxofilin does not affect the migration of mouse macrophages. BMdM were plated on transwell filters, left uninfected or infected with control or Δtxf parasites, and then incubated at 37°C for 2 h. Complete DMEM (black bars: − chemoattractant) or conditioned medium from 3T3 cells infected for 24 h with RHΔhpt parasites (grey bars: + chemoattractant) was added to the bottom portion of the transwells, and the cells were incubated for an additional 3 h at 37°C to allow migration. The filters were mounted on slides with mounting medium containing DAPI. The percentage migration was determined by calculating the number of cells on the bottom of the filter in each field relative to the total number of cells in the field using Image J software. This experiment was performed four times with comparable results; a representative experiment is shown. The error bars represent the standard deviation of the percentage migration from three biological replicate wells.

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Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

We have investigated the secretion and function of toxofilin, a rhoptry protein that was identified by its biochemical ability to bind to host and parasite actin in vitro. Using a highly sensitive BLA-based assay, we show that this protein appears to be introduced into host cells, probably during invasion. An important feature of the BLA assay is the ability to determine whether BLA fusion proteins are present at different time points post infection. We found that toxofilin was detectable in host cells within an hour and for as long as 32 h after infection. Given that pre-treatment of parasites with cytochalasin D resulted in toxofilin–HA–BLA secretion, we believe that toxofilin is introduced into host cells at an early time point during infection when rhoptry contents are secreted. The detection of toxofilin–HA–BLA in host cells as late as 32 h post infection most likely indicates that toxofilin is relatively stable and that the amount secreted during invasion persists in sufficient quantities to be detected after this amount of time. We cannot exclude the possibility, however, that a new pool of toxofilin is introduced into the host cell long after invasion. There is some precedent for rhoptry secretion into the PV late in infection: the bradyzoite-specific rhoptry protein BRP1 is secreted into the PV as bradyzoites develop from intracellular tachyzoites (Schwarz et al., 2005). While there is no precedent for the secretion of rhoptry proteins into the host cell cytosol once the parasite is contained within the PV, it remains possible that sufficiently sensitive means to detect such an event have not been previously available.

It is possible that the BLA substrate CCF2 may diffuse into the PV. When HFF infected with the parental RHΔhpt parasites for 24 h were exposed to CCF2-AM, fluorescent signal from the intact substrate could be visualized outlining the parasites within the PV (data not shown). This signal may be due to detection of the substrate on the host cytosol side of the PVM in the membranous network that interweaves between the parasites. It may also be due to diffusion of the substrate into the PV, since CCF2 is ∼1 kDa, and molecules of up to 1300–1900 Da can passively diffuse across the PVM (Schwab et al., 1994). In this situation, the substrate could be cleaved by parasite proteins on the interior of the PV. It is, thus, possible that the toxofilin fusion protein is secreted into the PV where CCF2 is cleaved, and the released coumarin subsequently diffuses back into the host cell cytosol. We do not believe that this is the case, however, because we cannot visualize toxofilin inside the PV of infected cells in the manner that other PV resident proteins are readily detected. The anti-HA antibody provides an easily detected signal in the rhoptries, and if the toxofilin fusion protein were released into the PV space, the dilution effect would be minimal. Nevertheless, the potential for CCF2 to diffuse into the PV will be an important consideration when examining BLA fusions to PV proteins, whether or not such proteins also reach the host cell cytosol.

The enzymatic activity of BLA is both an asset and a limitation. On the one hand, it allows for a dramatic amplification of the fluorescent signal, such that the secretion of relatively small amounts of protein into host cells will result in a detectable signal. On the other hand, it is difficult to quantify the amount of protein introduced because the assay quickly reaches saturation and establishing a standard curve for absolute amounts of protein is difficult. As a result, we cannot determine the precise number of toxofilin molecules secreted into a host cell during infection. However, to exploit the secretion of toxofilin during invasion, we have engineered parasites expressing toxofilin fused to the Cre recombinase with a nuclear localization signal (NLS). In host cells infected with these transgenic parasites, toxofilin–Cre is secreted in sufficient amounts to reach the host cell nucleus and to catalyse Cre-mediated recombination at loxP sites (A. Koshy and J. Boothroyd, unpubl. data).

The amount of toxofilin secreted by the parasite may have important implications for its activity in host cells, as we and others have observed that the introduction of high levels of toxofilin may have deleterious effects on the host cell. In the initial report on toxofilin from Tardieux and colleagues, the ectopic overexpression of a toxofilin–GFP fusion in mammalian HeLa cells (in the absence of parasite infection) resulted in a dysregulation of HeLa cell actin stress fibres and perturbed cell morphology (Poupel et al., 2000), and we have replicated these findings (data not shown). These data suggest that as an obligate intracellular parasite, T. gondii must limit the amount of toxofilin secreted into host cells. This would allow toxofilin to function appropriately but not impair the host cell to the extent that the parasite is unable to grow and replicate.

Throughout our examination of toxofilin activity, a lingering question has been the localization of toxofilin within infected host cells. The BLA assay effectively demonstrates that the toxofilin–BLA fusion protein gains access to the host cell cytosol during parasite rhoptry secretion. However, the indirect nature of the assay precludes a precise determination of the intracellular localization of toxofilin after it has been secreted. It is possible that toxofilin (1) functions locally, at the site of parasite invasion, (2) acts more broadly in the host cell cytosol or (3) traffics to a distinct subcellular compartment. Our failure to detect the protein by immunofluorescence argues against explanations 1 and 3, since in these cases, the protein should be concentrated enough to be seen. Further work will be required, however, to determine exactly where toxofilin traffics in infected cells.

To specifically address the function of toxofilin in infected host cells, we generated a toxofilin knockout (Δtxf) parasite. These parasites exhibited normal growth in vitro, indicating that toxofilin is not essential for parasite growth or replication. Although the absence of toxofilin did not appear to affect the staining of the host actin cytoskeleton in infected cells (data not shown), we did observe a very slight, but not statistically significant, acceleration in ionophore-induced egress with the Δtxf parasites compared with control parasites (Fig. 4C). Additionally, we examined whether toxofilin might have a functional effect on host cell processes that require actin dynamics. It has been demonstrated that T. gondii infection of dendritic cells induces a state of hypermotility that facilitates transmigration across endothelial cells (Lambert et al., 2006). Interestingly, we found that T. gondii-infected macrophages migrated similarly to uninfected macrophages, suggesting that there may be a difference in the effects of T. gondii infection on macrophage versus dendritic cell motility, or that migration across an endothelial cell monolayer involves distinct signals relative to migration in a transwell system. In the transwell system, however, it appears that toxofilin does not affect the migration of macrophages (Fig. 6) or of bone marrow-derived dendritic cells (data not shown). Finally, we examined the effect of toxofilin on inhibition of host cell apoptosis by infecting macrophages with the Δtxf or control parasites and assaying for Annexin V and propidium iodide staining of the infected cells by flow cytometry, as indicators of apoptotic cells. We did not observe a difference in cells infected with the Δtxf or control parasites in this assay (data not shown).

Using a variety of in vitro assays, we have not determined a role for toxofilin in host cells, suggesting that the function of toxofilin may only be revealed during an in vivo infection. In preliminary experiments using intraperitoneal infection of C57BL/6 mice, the Δtxf parasite had similar virulence to the control parasite (data not shown). Because the background strain for the Δtxf parasite is the hypervirulent type I RH strain (LD100 is a single parasite), however, any attenuation in virulence due to the absence of toxofilin would have to be very dramatic to be detectable as a difference in survival. It is possible that more sensitive read-outs may be necessary to uncover a role for toxofilin during in vivo infection.

Although we have observed that toxofilin is predominantly a rhoptry protein and appears to be secreted into infected host cells, it may also function inside the parasite. In the initial report on toxofilin, some weak staining of the parasite cytosol was observed (Poupel et al., 2000). In a subsequent report, toxofilin was observed only in the rhoptries (Bradley et al., 2005). We have found that toxofilin fusion proteins localize predominantly to the rhoptries but we also observe that in a minority of parasites (∼25%), we can detect a small amount of toxofilin in the parasite cytosol. Because the toxofilin fusion protein is expressed at similar levels to the endogenous copy of toxofilin (Fig. 1B), we do not believe that this cytosolic localization is due to overexpression, but this possibility cannot be excluded. Toxofilin was demonstrated to bind to both parasite and host actin, and was proposed to contribute to the relatively low level of F-actin in gliding parasites. Our results with the knockout parasites do not indicate a crucial role in gliding motility or parasite replication but other, more subtle roles cannot be excluded.

Overall, our data show that toxofilin appears to be injected into the host cell, but its role there, as well as in the parasite itself, remains a mystery. Nevertheless, the approach used – BLA fusions – represents a sensitive and efficient method for identifying rhoptry and other Apicomplexan proteins that are introduced into host cells.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Host cell culture and parasites

Human foreskin fibroblasts and 3T3 cells were maintained in complete DMEM: Dulbecco's modified Eagle's medium (Invitrogen, Carlsbad, CA) supplemented with 10% heat-inactivated fetal calf serum (FCS; Hyclone, Logan, UT), 2 mM l-glutamine, 100 U ml−1 penicillin and 100 μg ml−1 streptomycin.

Mouse bone marrow-derived macrophages were generated as previously described (Hamerman et al., 2005). Briefly, bone marrow was flushed from the femurs of C57BL/6 mice, and the red blood cells were lysed in ACK lysis buffer (Invitrogen, Carlsbad, CA). The cells were cultured in complete DMEM supplemented with 10% M-CSF. After 3 days, half the medium was replaced with fresh medium, and the cells were cultured for an additional 3–4 days before T. gondii infection assays.

Toxoplasma gondii tachyzoites were maintained by serial passage in confluent monolayers of HFF grown in complete DMEM at 37°C with 5% CO2.

Generation of toxofilin knockout and transgenic parasites

The toxofilin gene was targeted in the RH strain of T. gondii by using homologous recombination to replace the toxofilin ORF with the hypoxanthine-xanthine-guanine phosphoribosyltransferase (hxgprt) gene. To do this, ∼2 kb of genomic sequence flanking the toxofilin ORF in the RH strain was amplified by PCR using the following primers: (5′-targeting sequence) 5′-ATTGATGCGGCCGCTGGAGTGCGTCCTTGCTTCAT-3′; 5′-CCATTGGATATCGTTGACCGGTTACAAGTGGAA-3′; (3′-targeting sequence) 5′-CCTGACAAGCTTAAACGACGACGAACCCGCG-3′; 5′-TTATCAGTTAACCATTTCCGGGCAATGGACAAAG-3′. The 5′-targeting sequence was cloned into the NotI and EcoRV restriction sites of the pTKO vector (kindly provided by G. Zeiner, Stanford University, Stanford, CA), and the 3′-targeting sequence was cloned into the HindIII and HpaI restriction sites of pTKO to generate the targeting plasmid. The plasmid was linearized by restriction digest with NotI, and 30 μg of plasmid was transfected into the RHΔhpt strain of parasites by electroporation, as described previously (Soldati and Boothroyd, 1993). The parasites were added to monolayers of HFF cells and 24 h after transfection, the medium was replaced with complete DMEM supplemented with 50 μg/ml of mycophenolic acid and 50 μg/ml of xanthine (MPAX) to select for HXGPRT activity. Fourteen days after transfection, the parasites were single-cell cloned by limiting dilution, and the genomic DNA from individual clones was analysed by PCR for the presence of the toxofilin gene using primers specific for the toxofilin ORF and for specific recombination of the targeting construct at the toxofilin locus. Two parasite clones were used in subsequent analyses: Δtxf clone 11.1, in which the toxofilin ORF was specifically deleted, and control clone 11.4, in which the targeting construct integrated non-specifically in the T. gondii genome. In control clone 11.4, the toxofilin ORF remains intact, but the presence of the targeting construct confers HXGPRT activity and MPAX resistance, as in Δtxf clone 11.1. Both the Δtxf clone 11.1 and the control clone 11.4 are GFP+.

Transgenic parasites expressing engineered forms of toxofilin were generated by transfection of RHΔhpt strain parasites (by electroporation) with plasmids expressing toxofilin–HA (SP7) or toxofilin–HA–BLA (SP3), selection in MPAX, as described above, and single-cell cloning by limiting dilution. The plasmid SP7 was generated by PCR amplification of ∼1.1 kb upstream of the toxofilin ORF and includes the toxofilin ORF from the RH strain with a mutated stop codon. The resulting ∼1.85 kb fragment was cloned into the HindIII and NcoI sites of the pGRA vector (kindly provided by J.D. Dunn, Stanford University, Stanford, CA), which incorporated a HA epitope tag sequence and a stop codon downstream of the NcoI site. This construct (SP7) contains the toxofilin ORF with a C-terminal HA tag. SP3 was generated by PCR amplification from the RH strain of ∼1.1 kb upstream of the toxofilin ORF, the toxofilin ORF with a mutated stop codon, and a sequence encoding the HA epitope tag at the 3′ end. The resulting fragment was cloned into the HindIII and SfoI sites of the pGRA vector. The β-lactamase (BLA) cDNA with a stop codon was PCR-amplified from the pGRA vector and cloned into the SfoI and PacI sites downstream of the toxofilin cDNA and HA. This construct (SP3) contains the toxofilin ORF and HA tag fused to BLA.

Western blot analysis

Lysates were generated from RHΔhpt, toxofilin–HA clone 7.7 or toxofilin–HA–BLA clone 3.3 parasites by the treatment of parasite pellets with SDS-PAGE loading dye containing 10% β-mercaptoethanol. The lysates were separated by SDS-PAGE, transferred to a PVDF membrane, and the membrane was blocked with TBST (TBS, 0.05% Tween-20) containing 5% milk for 1 h at room temperature. The membranes were incubated with horseradish peroxidase (HRP)-conjugated rat anti-HA (clone 3F10) monoclonal antibodies (Roche, Indianapolis, IN) at a dilution of 1:500, mouse anti-toxofilin polyclonal serum at a dilution of 1:2000, or rabbit anti-SAG1 polyclonal serum at a dilution of 1:10000 for 1 h at room temperature. For secondary antibodies, goat anti-mouse-HRP (for toxofilin) or goat anti-rabbit-HRP (for SAG1) were used for 1 h at room temperature. HRP activity was detected using SuperSignal West Pico Chemiluminescent Substrate (Pierce, Rockford, IL).

BLA assay

For microscopy, monolayers of HFF cells grown on glass chamber slides were infected with RHΔhpt, toxofilin–HA clone 7.7 or toxofilin–HA–BLA clone 3.3 parasites at a nominal multiplicity of infection (moi) of 10 and incubated at 37°C. At various times post infection, the infected HFF were incubated with the BLA substrate CCF2-AM (Invitrogen, Carlsbad, CA) at a 1× concentration in complete DMEM for 1 h at room temperature in the dark. Live, infected cells were then visualized using a Leica SP2 AOBS Confocal Laser Scanning Microscope (Cell Sciences Imaging Facility, Stanford University, Stanford, CA) with a blue diode 405 nm laser for excitation and with detection filters set at 410–450 nm for coumarin and 493–550 nm for fluorescein.

For flow cytometry, RHΔhpt or toxofilin–HA–BLA clone 3.3 parasites were treated with 1 μM cytochalasin D or with DMSO as a control for 10 min and then added to monolayers of HFF in six-well dishes at a nominal moi of 10. To confirm that the cytochalasin D treatment effectively inhibited parasite invasion, we titrated the amount of cytochalasin D used to pre-treat the parasites and determined 1 μM to be the optimal inhibitory concentration. The infected cells were incubated at 37°C for 1 h and loaded with the BLA substrate CCF2-AM at a 1× concentration for 1 h at room temperature in the dark. The cells were washed with 1× PBS and trypsinized. The re-suspended cells were examined on a modified FACStar flow cytometer (BD, San Jose, CA) with the 407 nm krypton laser for the detection of coumarin (in the cascade blue channel). FlowJo software (Tree Star, Ashland, OR) was used for analysis.

Immunofluorescence assays

Monolayers of HFF cells grown on glass coverslips were infected as described above with toxofilin–HA clone 7.7, toxofilin–HA–BLA clone 3.3, RHΔhpt or Δtxf clone 11.1. The cells were washed with PBS, and then either fixed with 2.5% formaldehye in PBS for 10 min and permeabilized with 0.2% Triton X-100 in PBS for 20 min, or fixed and permeabilized with 100% ethanol for 20 min. The cells were then blocked with 3% bovine serum albumin (BSA) in PBS for 1 h. Primary and secondary antibodies were diluted in blocking buffer and incubated on cells for 1 h. All incubations were performed at room temperature, and the cells were washed three times with 1× PBS after each antibody stain. For primary antibodies, rat anti-HA (clone 3F10; Roche, Indianapolis, IN) was diluted 1:500, mouse anti-ROP2/4 (clone T34A7) was diluted 1:1000, mouse anti-toxofilin polyclonal serum was diluted 1:1000, and rabbit anti-SAG1 polyclonal serum was diluted 1:10000. Alexa fluor 488-conjugated anti-rat Ig (1:2000) or anti-mouse Ig (1:2000) and Alexa fluor 594-conjugated anti-mouse Ig (1:2000) or anti-rabbit Ig (1:5000) (all from Invitrogen, Carlsbad, CA) were used as secondary antibodies. After the final wash, the coverslips were mounted on glass slides using Vectashield mounting medium (Vector Laboratories, Burlingame, CA) and visualized by using an Olympus BX60 microscope with a 100× oil objective.

Parasite invasion, growth and egress assays

For invasion assays, control or Δtxf parasites were allowed to settle onto monolayers of HFF cells on glass coverslips in the presence of high potassium Endo buffer [10 mM K2SO4, 2.5 mM Mg2SO4, 1 mM glucose, 5 mM Tris (pH 8.2) and 3.5 mg ml−1 BSA] at 37°C for 20 min. The Endo buffer was replaced with warm DMEM for the indicated time periods to allow parasite invasion (as in Kafsack et al., 2004), and the cells were fixed and stained for extracellular SAG1 (as described above). The control and Δtxf parasites were scored for invasion efficiency by counting the number of intracellular parasites (GFP+SAG1-) relative to the total number of parasites in each field.

For growth assays, monolayers of HFF on glass coverslips were infected with control or Δtxf parasites at a low moi and washed after 2 h to remove uninvaded parasites. Twenty-four hours later, the cells were fixed, and the coverslips were mounted on slides. For each slide, the number of parasites per vacuole in eight fields was counted and expressed as a percentage of the total number of parasites in the field.

For egress assays, monolayers of HFF were infected with control or Δtxf parasites at a low moi and washed after 2 h to remove uninvaded parasites. At 30 h post infection, the cells were washed three times with warm PBS. The calcium ionophore A23187 (Sigma-Aldrich) diluted to 1 μM in warm HBSSc [Hanks' Balanced Salt Solution (HBSS; Invitrogen, Carlsbad, CA) supplemented with 1 mM MgCl2, 1 mM CaCl2, 10 mM NaHCO3, 20 mM Hepes, pH 7.0] was then added to the cells for 15, 30, 60 or 90 s. The ionophore was removed, and the cells were immediately fixed. For each time point, the number of lysed vacuoles (indicating parasite egress) was counted in five fields and expressed as a percentage of the total number of vacuoles in the field. This experiment was performed five times on different days, and the graph shown includes the data from all five experiments. P-values reflect results from the Wald F-test, adjusted for clustering, under the null hypothesis of equivalent proportions of egress in the control and Δtxf groups.

Phagocytosis and migration assays

For phagocytosis assays, BMdM were infected with control or Δtxf parasites at a nominal moi of 2, or the medium was replaced for the mock infection control. The cells were incubated at 37°C for 2 h, washed three times with PBS, and the medium was replaced with phagocytosis uptake buffer: HBSS supplemented with 20 mM Hepes, pH 7.4. pHrodo E. coli Bioparticles (Invitrogen, Carlsbad, CA) re-suspended in uptake buffer were added to the infected macrophages and incubated in a 37°C water bath for 20 min. The cells were washed three times with 1× PBS and removed from the plate by scraping. The re-suspended cells were examined on a modified FACStar flow cytometer (BD, San Jose, CA) for the detection of GFP and rhodamine. FlowJo software (Tree Star, Ashland, OR) was used for analysis.

Migration assays were performed by plating 105 BMdM on transwell filters with 5 μm pores in 24-well transwell plates (Corning, Lowell, MA) for 24 h. The cells were then infected with control or Δtxf parasites at a nominal moi of 4 and incubated at 37°C for 2 h. Conditioned medium from 3T3 cells infected for 24 h with RHΔhpt parasites was used as the chemoattractant. The conditioned medium was added to the bottom portion of the transwells, and the cells were incubated for an additional 3 h at 37°C to allow migration. The filters were washed twice with PBS, fixed, and mounted on slides with mounting medium containing DAPI. The cells on the top and bottom of the filter were imaged using an Olympus BX60 microscope with a 60× oil objective and were counted using Image J cell counting software (NIH) to detect the DAPI staining in five fields. The percentage migration was determined by calculating the number of cells on the bottom of the filter in each field relative to the total number of cells in the top and bottom of each field.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

We would like to thank the entire Boothroyd lab for helpful discussion, in particular Sandeep Ravindran for suggestions on characterizing the Δtxf parasites and Dr Gusti Zeiner for help with the pTKO plasmid. We also thank Dr Isabelle Tardieux (INSERM) for discussion about toxofilin and for sharing unpublished data from her lab, and Dr Tim Bruckner for help with statistical analysis for the egress assay. This work was supported by NIH Grant AI21423 (to J.C.B.) and the A.P. Giannini Foundation Medical Research Fellowship (to M.B.L.).

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  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Fig. S1. Leakage of toxofilin–HA–BLA into the host cell is not detectable. Host cells were infected for 1 h with parasites expressing toxofilin–HA–BLA, toxofilin(-SP)–HA–BLA (the signal peptide mutant form of the toxofilin–BLA fusion protein) or HA–BLA. CCF2-AM was then added for an additional hour and the monolayers were subsequently imaged using a Leica SP2 AOBS Confocal Laser Scanning Microscope with a blue diode 405 nm laser for excitation and with detection filters set at 410–450 nm for coumarin and 493–550 nm for fluorescein. In the inset, the white arrowheads indicate parasites that did not take up the CCF2-AM (and, thus, are black) whereas the red arrowheads show those that did, presumably because they invaded after addition of the substrate to the cultures. The latter are blue indicating cleavage of CCF2-AM and thus contact between BLA and the substrate.

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