https://orcid.org/0000-0001-9778-7337
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Figures
Abstract
The phylum Apicomplexa contains ~ 6000 known species of unicellular eukaryotic parasites. A unifying feature among the apicomplexans is the apical complex, which varies in complexity in different lineages, but always contains an annulus (a.k.a. the apical polar ring) into which the minus ends of an array of cortical microtubules are embedded. In Toxoplasma gondii, the apical complex also includes the conoid, which contains several signaling and structural proteins critical for parasite motility. The conoid extends and retracts through the apical polar ring in a calcium-dependent manner. Here we report the identification of several new apical polar ring components, including APR9, which is highly conserved among the apicomplexans and their free-living relative Chromera velia. The loss of APR9 alone has only a moderate impact on the parasite lytic cycle. However, the knockout of both APR9 and KinesinA (another apical polar ring component) paralyzes the parasite and drastically impairs invasion, egress and the lytic cycle. The double-knockout displays multiple subcellular abnormalities, including the formation of an apical actin concentration, impaired conoid extension, and significantly reduced secretion of a major adhesin (MIC2) upon stimulation with a calcium ionophore. These findings reveal that the apical polar ring plays a critical role in parasite motility and contributes to multiple subcellular processes.
Author summary
A large family of single-cell organisms, the apicomplexans, parasitize a wide range of multiceullar organisms (the Metazoa). Many of them are important human pathogens, such as Toxoplasma, Cryptosporidium, and Plasmodium. These parasites invade into a host cell with their apex making first contact. One major apical feature shared by the apicomplexans is the apical polar ring. In this work, we identified several new apical polar ring components in Toxoplasma. One of them is APR9, which is present not only in the apicomplexans, but also in a free-living alga related to these parasites. We found that APR9 is functionally connected to KinesinA, another apical polar ring component, because the removal of both proteins severely compromises the ability of the parasite to actively move into and disseminate from its host cell, though removal of either protein alone has little effect. In addition, upon the loss of both APR9 and KinesinA, several cellular activities at the parasite apex are perturbed, including actin kinetics, the protrusion of an apical cytoskeletal structure, and the secretion of a major adhesin. These results indicate that the apical polar ring is important for parasite movement, coordinating several distinct processes at the parasite apex.
Citation: Munera Lopez J, Arias Padilla LF, Tengganu IF, Hao Y, Zhang Y, Florens L, et al. (2026) The human parasite, Toxoplasma gondii, is paralyzed without two components of the apical polar ring. PLoS Pathog 22(6): e1014378. https://doi.org/10.1371/journal.ppat.1014378
Editor: Michael L. Reese, UTSW: The University of Texas Southwestern Medical Center, UNITED STATES OF AMERICA
Received: March 7, 2026; Accepted: June 12, 2026; Published: June 26, 2026
Copyright: © 2026 Munera Lopez et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: Raw data and search results files for the proteomics analysis have been deposited to the Proteome Xchange (accession: PXD074785) via the MassIVE repository and may be accessed at http://massive.ucsd.edu with MSV000100958 as the Dataset Identifier. Original mass spectrometry data underlying this manuscript can be accessed after publication from the Stowers Original Data Repository at http://www.stowers.org/research/publications/libpb-2610. All other relevant data and details of sources can be found within the article and its supplementary information.
Funding: Research reported in this publication was supported by the National Institute of Allergy and Infectious Diseases of the National Institutes of Health under Award Number 1R01AI184433 awarded to KH. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
The apicomplexans are parasitic protists that infect many different types of metazoans [1]. Notable human pathogens among the apicomplexans include Cryptosporidium, Toxoplasma, and Plasmodium [1–4]. While the host range differs greatly among the apicomplexans, they are unified by key genetic and cellular architectures, the most iconic of which is the apical complex (Fig 1A), the eponym of the phylum.
A. Diagrams (modified from [28]) illustrating the organization of the tubulin-containing cytoskeleton and associated structures in T. gondii. IMC: Inner membrane complex. B. Diagram of the workflow for enrichment and identification of candidate apical polar ring proteins from the mEmeraldFP-APR2 knock-in parasite line. mEmeraldFP-APR2 was used as the bait in immunoprecipitation (IP) with the high-affinity llama anti-FP antibody conjugated to agarose beads (“GFP-trap”). C. Table of peptide counts, unique spectral counts, and fitness scores for known apical polar ring proteins and for TgGT1_223790, 289990, and 295420 identified by MudPIT in the IP. No spectra were detected for these proteins in the negative control (untagged WT parental parasite), except for APR6, for which three spectra were detected. See S1 Table for the complete list of proteins identified.
The apical complex contains a cytoskeletal scaffold as well as associated membrane-bound secretory organelles [5]. In Toxoplasma, the cytoskeletal apical complex is well characterized structurally for mature parasites. The major elements are the conoid, the preconoidal rings, and intra-conoid microtubules (MTs), as well as the apical polar ring, which associates with the minus ends of 22 evenly-spaced cortical MTs [6–12]. Components of the cytoskeletal apical complex are involved in signaling and the mechanics of parasite invasion into the host cell [13–22]. The conoid is made of ribbon-like fibers, polymerized from the same tubulin subunits as the cortical MTs despite their radical structural differences [8,11,12,23,24]. It is a mobile structure. In response to changes in the intra-parasite calcium concentration, the conoid, together with the preconoidal rings and intra-conoid MTs, protrudes or retracts through the apical polar ring [6–8]. A number of apical polar ring proteins have been identified and characterized previously [16,17,25–31]. These proteins individually have varying degrees of impact on different aspects of the parasite lytic cycle. Some proteins, such as KinesinA and APR1, also have a notable structural role [17]. Without KinesinA, the electron dense, well-defined annulus conventionally noted as the apical polar ring becomes undetectable by negative staining electron microscopy (EM). The removal of APR1 results in frequent conoid detachment. Interestingly, the cortical MT array appears to be normal in parasites lacking only KinesinA or only APR1. However, when both KinesinA and APR1 are knocked out, cortical MTs often detach in groups from the apex in adult parasites as well as in large daughter parasites [17,32].
The structural integration of the apical polar ring and cortical MTs suggests a role of the apical polar ring as the organizing center for the cortical MTs. Using APR2—an early component of the apical polar ring—as the marker, we examined how the apical polar ring is constructed with respect to the cortical MT array [28]. We established that the assembly of the apical polar ring initiates prior to that of the MT array, starting from an arc with its open side facing the centrioles. As the arc grows into a ring, the MT array begins to assemble on the closed side. This observation supports the hypothesis that the apical polar ring initiates and template new MT arrays by directional, stepwise assembly coupled with depositing or recruiting factors for MT polymerization. Two other studies also showed that the MT nucleator γ-tubulin associates with the apical polar ring transiently during early stage of daughter assembly, further supporting the role of the apical polar ring in initiating the new MT array [33,34].
The removal of APR2 has only a minor impact on MT organization [28]. However, due to its early recruitment to the apical polar ring, we decided to use it as the bait in immunoprecipitation. Several new components of the apical polar ring were identified. Among them is APR9, a protein found not only in apicomplexans, but also in Chromera velia, a free-living relative of these parasites. The knockout of APR9 alone results in a mild phenotype in the parasite lytic cycle. In contrast, when APR9 is removed together with KinesinA, the lytic cycle is greatly inhibited. The ΔkinesinAΔapr9 parasite is nearly completely paralyzed, which severely impairs egress and invasion. Even though both APR9 and KinesinA are early components of the apical polar ring, MT organization is perturbed only in a small fraction of the ΔkinesinAΔapr9 parasites. However, the ΔkinesinAΔapr9 parasite does display abnormalities in multiple subcellular processes, including the formation of an apical actin concentration, impaired conoid extension, and significantly reduced secretion of a major micronemal adhesin (MIC2) upon stimulation with a calcium ionophore. These findings reveal that, in addition to its role as an MT organizing center, the apical polar ring coordinates distinct subcellular processes at the parasite apex and plays a critical role in parasite motility. They also highlight the importance of assessing the function of the components of the apical polar ring not only individually but also in combination with others.
Results
Enrichment and identification of new apical polar ring proteins using mEmeraldFP-APR2 as the bait
We previously showed that APR2 (TgGT1_227000) is an early component of the apical polar ring [28]. To enrich and identify new apical polar ring components, we carried out immunoprecipitation (IP) using an anti-GFP nanobody and mEmeraldFP (mE)-APR2 knock-in parasites. The parental parasite having untagged APR2 was used as the negative control. Parasites were homogenized after Triton X-100 (TX-100) extraction, and a high-affinity llama anti-GFP antibody conjugated to agarose beads (“GFP-trap”) was used for the pull-down. The IPs were analyzed by Multidimensional Protein Identification Technology (MudPIT) to identify enriched proteins [13,35–37] (Fig 1B). Known proteins among the top 15 hits include APR2, seven other components of the apical polar ring (KinesinA, APR1, APR3–5, APR7, MLC3), a preconoidal ring protein (Pcr7), two proteins previously localized to the parasite apex and other cytoskeletal structures (CDPK6 and IMC15), and the cytochrome c oxidase subunit ApiCOX35 [20,27,29,38–41] (S1 Table, Fig 1C). Three new hypothetical proteins were also identified in the IP: TgGT1_223790 (20 Peptides), 289990 (5 P), and 295420 (10 P). To examine the localization of these proteins, we generated knock-in lines, in which the genomic locus of the target gene is replaced with a DNA fragment including the CDS for mEmeraldFP (mE)-geneX with a 3’UTR and an expression cassette for the HXGPRT selectable marker (Fig 2A-2B). Expansion microscopy revealed that these three proteins are localized to an apical annulus immediately above the conoid in intracellular parasites, which is the expected location for the apical polar ring (Figs 2C-2E, S1). They are thus named APR9 (223790), APR10 (289990) and APR11 (295420).
A-B. Generation of mEmeraldFP (mE) tagged APR9(223790), APR10(289990), and APR11(295420) knock-in parasites. A: Schematic illustrating the strategy used to generate the mE-tagged knock-in parasites. Positions for the primers in the diagnostic genomic PCRs shown in B and expected DNA fragment sizes are indicated. G-3: GRA 3’UTR, HXGPRT Cas: HXGPRT expression cassette. B: Diagnostic genomic PCRs of the RHΔku80 parental (WT) and the knock-in lines confirming the homologous integration of the mE fusion in the knock-in lines. C-E. Projections of expansion microscopy (ExM) images of mE-tagged APR9, APR10, APR11 knock-in parasites labeled with anti-GFP and anti-tubulin antibodies. Insets (2X) correspond to the regions indicated by the arrows, which show that APR9, 10, and 11 are localized to an apical annulus positioned anterior to the retracted conoid (“C”, bracket), consistent with the localization of the apical polar ring. Arrowhead indicates intra-conoid MTs (“ICM”). Scale bar indicates an estimated length of ~ 1 µm before expansion and is based on an estimated expansion ratio of ~5.4 [28,32]. Image contrast was adjusted to optimize display. See S1 Fig for magenta-green version of the images.
Of the three new APR proteins, we decided to first focus on APR9. Aside from weak homology with the ENBA-3A domain at its C-terminus, APR9 does not have any readily recognizable protein domains. However, APR9 is highly conserved within Apicomplexa, and is found in all sequenced apicomplexan lineages. Furthermore, a well-conserved ortholog (Cvel_27126) is also found in Chromera, a free-living relative of the apicomplexans (Fig 3A-3B). In T. gondii, APR9 shares strong sequence similarities with Tg219500, which was previously localized to the apical polar ring and named as APR4 in [29]. Multiple APR4/9 homologs are found in the genomes of members of the Sarcocystidae family (e.g., T. gondii, Neospora caninum and Sarcocystis neurona), likely a result of a gene duplication event in their common ancestor. Alpha-fold 3 [42] predicts that both APR4 and APR9 have a core of multiple alpha-helices, although TgAPR9 and orthologs in Plasmodium berghei and Chromera velia are predicted to contain extended unstructured regions (S2 Fig).
A. Presence (filled circle) or absence (open circle) of predicted homologs for TgAPR1, APR2, APR9, APR10 and APR11 and KinesinA in representative members of the phylum Apicomplexa with available genome sequences, as well as their photosynthetic relatives, Chromera velia and Vitrella brassicaformis. Protein sequences of T. gondii homologs were used as queries for the BLAST searches against the predicted protein databases in VEupathDB (http://veupathdb.org). An E-value cut-off of 1e-5 was used to identify APR1, APR2, APR9, APR10, and APR11 orthologs. Grey circle indicates a hit with 1e-5 > E-value > 1e-10. Due to the conserved nature of the kinesin-fold, only hits with an E-value less than 1e-16 were considered tor KinesinA orthologs, and were manually curated. B. Phylogenetic analysis of APR4/9 homologs found in Apicomplexa and Chromera velia constructed using T-coffee (http://www.ebi.ac.uk/jdispatcher/msa/tcoffee). Bar indicates bootstrap values of 0.5 and 1. Except for TgAPR4 and TgAPR9, all proteins are listed by their gene IDs in VeupathDB.org. C-H. Projections of ExM images of mE-APR9 knock-in parasites through daughter development. Insets (2X) in H show that the APR9 labeling is apical to the retracted daughter conoid (single section),but posterior to the protruded conoid in the mother parasite (projection of a substack). Cen: centrioles. Scale bar ≈ 1 µm prior to expansion based on an estimated expansion ratio of ~5.4 [28,32]. Image contrast was adjusted to optimize display. See S3 Fig for magenta-green version of the images.
APR9 is an early component of the apical polar ring and its localization is independent of APR2, APR4 and KinesinA
To determine the recruitment schedule of APR9 in developing daughters, we used expansion microscopy and labeled the FP-knock-in parasite with anti-GFP and anti-tubulin antibodies. The timing of recruitment of APR9 was determined by using the number and length of cortical MTs to estimate daughter developmental stages. Similar to APR2, APR9 is assembled into the precursor of the apical polar ring prior to the appearance of the daughter MTs. As reported before [28], the apical polar ring extends towards the centriole region from an arc to a closed ring. The APR9 signal in the apical polar ring persists throughout the daughter development and in the mature parasite (Figs 3C-Fig 3H, S3). In addition to the prominent localization to the apical polar ring, there is also notable mE-APR9 signal in the daughter cortex (Fig 3G). To determine how the localization of APR9 is affected by other known apical polar ring components, we generated mE-APR9 knock-in lines in the Δapr2, Δapr4, and ΔkinesinA parasites (Figs 4A-4E, S4-S5). APR9 is targeted to an apical annulus in these lines, indicating that the localization of APR9 is independent of APR2, APR4, and KinesinA (Figs 4C-4E, S5). Interestingly, while the positioning of the APR9 ring appears to be normal in the Δapr2 and Δapr4 parasites, it occasionally appears to be tilted in the ΔkinesinA parasite (Fig 4E, insets).
A-B. Southern blot analysis of the apr9 locus in the WT, mEmeraldFP- APR9 (mE-APR9 KI), Δapr9, ΔkinesinA:mE-APR9 knock-in (ΔkinA:mE-APR9 KI), ΔkinesinAΔapr9 (ΔkinAΔapr9), ΔkinesinAΔapr9:mE-APR9 complement clone 1 and clone 2,Δapr4:mE-APR9 knock-in, Δapr4Δapr9, Δapr2:mE-APR9 knock-in, and Δapr2Δapr9 parasites. A: Schematic for the predicted APR9 locus in WT, mE-APR9 knock-in, and Δapr9 lines. Restriction sites, hybridization targets of the Southern blot probes for the apr9 coding region (CDS probe 1 and 2, orange bars), regions upstream (5’ UTR probe, blue bar) and downstream (3’ UTR probe, purple bar) of the CDS, and the expected DNA fragment sizes are indicated. B: Southern blot analysis confirming the homologous integration of the mE-APR9 fusion in the knock-in lines, and the deletion of the apr9 locus in the Δapr9 lines. See S4 Fig for Southern blot analysis of the apr4 locus in the APR4 knock-in and knockout lines. C-E. Projections of expansion microscopy (ExM) images of mE-APR9 knock-in in Δapr2 (C), Δapr4 (D) and ΔkinesinA (E) parasites. Inset (2X) in E indicates a daughter APR9 ring tilted relative to the parasite cortex. See S5 Fig for magenta-green version of the images. F. Plaque assays of WT, Δapr9, Δapr4Δapr9, Δapr2Δapr9, ΔkinesinA:mE-APR9 knock-in (ΔkinA:mE-APR9 KI), ΔkinesinAΔapr9 (ΔkinAΔapr9), and ΔkinesinAΔapr9:mE-APR9 complement clone 1 and clone 2 (ΔkinAΔapr9:mE-APR9 Comp 1 and 2) parasites. All cultures were infected with 100 parasites and incubated for 7 days before crystal violet staining. Inset (10X) highlights the small plaque indicated by the red arrow. G. Bar graphs that quantify plaquing efficiency for the lines shown in F. Three to seven independent plaque assays were performed for each line. For each replicate, the total lysed area of three wells, each infected with 100 parasites for 7 days, was measured. Error bars represent standard error of the mean (SEM). Value indicated in each bar is average ± SEM for the corresponding line. P-values were calculated in KaleidaGraph using two-tailed unpaired Student’s t-tests with unequal variances.
Impact of APR9 deletion on the parasite lytic cycle
To determine the impact of APR9 on the parasite lytic cycle, we generated a knockout line, Δapr9, by transient expression of Cre-recombinase in the mE-APR9 knock-in parasite to excise the LoxP flanked region (Fig 4A-4B). Putative knockout clones were first identified by the loss of mE-APR9 fluorescence. The excision of the mE-APR9 + selectable marker in the LoxP-flanked region was then confirmed by Southern blotting. Parasite growth was assessed by the ability of parasites to generate “plaques” over 7 days of incubation with host cell monolayers. The plaques are lesions in the monolayer generated by continuing destruction of the host cells by the parasite via cycles of invasion, replication, and egress. The Δapr9 parasite shows only moderate growth defects in plaque assays (Fig 4F-4G). This is consistent with the prediction from a previous genome-wide CRISPR-Cas9 screen, which assigned a phenotype score of 0 for APR9 (ToxoDB.org, [43], Fig 1C).
Given the broad conservation of APR9, which extends to a free-living relative of apicomplexans, the weak phenotype of the Δapr9 parasite is somewhat surprising. One possible explanation is the function and/or structural redundancy among the apical polar ring components. Because APR4 is a close homolog of APR9, we generated a double-knockout line (Δapr4Δapr9) by transiently expressing the Cre-recombinase in the Δapr4:mE-APR9 knock-in parasite to excise the mE-APR9 knock-in locus (Fig 4A-4B). We similarly generated two additional double knockout lines: a Δapr2Δapr9 line, because APR9 was identified in the APR2 IP, and a ΔkinesinAΔapr9 line, because KinesinA shows a synergistic effect with two other apical polar components, APR1 and APR2 [17,28]. We found that both Δapr4Δapr9 and Δapr2Δapr9 parasites show a moderate defect in plaque assays (Fig 4F-4G). In contrast, the phenotype of the ΔkinesinAΔapr9 parasite is severe. The ΔkinesinAΔapr9 clones were isolated with great difficulty. After 7 days of incubation, the parental lines formed many sizable plaques, but the few plaques in the ΔkinesinAΔapr9 culture were barely visible (Fig 4F). The plaquing efficiency of the ΔkinesinAΔapr9 parasite is ~ 0.03% of the wild-type parasite, ~ 0.05% of Δapr9, and ~ 0.06% of ΔkinesinA:mE-APR9 knock-in parasite, its immediate parental line (Fig 4G). We generated two complemented lines (ΔkinesinAΔapr9:mE-APR9 complement clone 1 and clone 2) in which the mE-APR9 expression cassette is integrated in different genomic loci in the ΔkinesinAΔapr9 parasite (Fig 4B). Plaquing efficiencies in both complemented lines are restored to a level similar to that of the ΔkinesinA:mE-APR9 knock-in parental line (Fig 4F-4G).
The loss of KinesinA and APR9 slows parasite replication and severely compromises parasite motility
To further determine the specific defects in the ΔkinesinAΔapr9 parasite, we examined its replication, invasion, and egress behaviors. We found that the ΔkinesinAΔapr9 parasites replicate with a doubling rate lower than that of the WT parasites (Fig 5A). The replication rate of the WT parasite between 12 hr and 36 hr is 2.95 doublings/24hr. The doubling of the ΔkinesinAΔapr9 parasite is 2.2 doublings/24hr. Although this difference is statistically significant (p = 0.0025), it translates to only ~32 fold fewer parasites over 7 days. This cannot fully explain the drastic difference in plaquing efficiency, which differs by more than 3,700 fold between the WT and the ΔkinesinAΔapr9 parasites. Therefore, other aspects of the lytic cycle must also be affected by the loss of KinesinA and APR9. Indeed, host cell invasion of the ΔkinesinAΔapr9 parasite is severely impaired when assessed using a dual-color invasion assay that distinguishes between intracellular and extracellular parasites based on antibody accessibility to a surface antigen, P30 [44,45] (Table 1). The ΔkinesinAΔapr9 parasite invades at ~ 11% (p < 0.0001) of the level of the WT parasites. Both invasion and egress rely on active gliding motility of the parasite. To determine if the motility of ΔkinesinAΔapr9 is affected, we carried out live egress assays induced by the calcium ionophore, A23187. The calcium ionophore treatment elevates the calcium concentration in the parasite cytoplasm. In the wild-type parasite, upon A23187 treatment, the combined effects of active parasite movement and host-cell-membrane permeabilization by pore-forming proteins secreted from the parasite enable a rapid dispersal of the parasites from the host cell (Fig 5B-5C). The ΔkinesinAΔapr9 parasites, however, were nearly completely paralyzed. Most parasites remained immotile. Only sporadic movement was observed in a small minority of the parasites (Fig 5B-5C, S1 Video). Compared with the WT, Δapr9, ΔkinesinA, and the complemented lines, it takes significantly longer for the first ΔkinesinAΔapr9 parasite to become mobile and initiate active egress, with a p-value of 0.0003, 0.002, 0.0005, and 0.0003, respectively. Furthermore, for the WT, Δapr9, ΔkinesinA, and complemented lines, the median time for ≥ 50% of the parasites in a vacuole to actively egress is around 50 seconds. In contrast, the ΔkinesinAΔapr9 line did not reach this threshold in any vacuole during the 300-second observation period (p < 0.0001). Notably, this phenotype is specific to motility but not due to a universal block of calcium sensing because the ΔkinesinAΔapr9 parasites still respond to the A23187 treatment by secreting factors to permeabilize the host cell membrane (Fig 5C, S1 Video). This is indicated by the abrupt change in morphology and contrast of the host cell in the DIC image as well as the rapid labeling of the host cell nucleus by the cell-impermeant dye DAPI upon A23187 treatment.
A. Replication assay of RHΔku80 parental (WT), Δapr9, ∆kinesinA:mE-APR9 knock-in (ΔkinA:mE-APR9 KI), ΔkinesinAΔapr9 (ΔkinAΔapr9), and ΔkinesinAΔapr9:mE-APR9 complement parasites. X-axis: time after the start of parasite-host cell incubation. Y-axis: number of parasite doublings. Error bars: SEM. B. Dot plots of time taken for the first parasite to initiate egress (left) and time taken to disperse (right) during A23187-induced egress for RHΔku80 parental (WT, N = 18), Δapr9 (N = 19), ∆kinesinA (N = 6), ΔkinesinAΔapr9 (N = 19), and ΔkinesinAΔapr9:mE-APR9 complement (comp, N = 20) parasites. The median for each dataset is indicated by a black bar. Time-taken-to-disperse for each vacuole was defined as the time point at which more than 50% parasites have actively egressed. ΔkinesinAΔapr9 parasites did not reach this threshold in any vacuole during the 300-second observation period (*). C. Time-lapse images of A23187-induced egress of RHΔku80 parental (WT), ΔkinesinAΔapr9 (ΔkinAΔapr9), and ΔkinesinAΔapr9:mE-APR9 complement (ΔΔ:mE-APR9 comp) parasites. The cell-impermeant nucleic acid dye DAPI was added to the medium prior to the A23187 treatment. Labeling of the host cell nuclear DNA indicates host-cell permeabilization. Note that although ΔkinAΔapr9 parasites display drastically reduced motility, they still secrete factors to lyse the host cell upon A23187 treatment, as indicated by the change in the host-cell morphology in the DIC images and by DAPI entering and binding to DNA in the host cell nucleus (S1 Video). Insets (1X) show DAPI images of the host cell nucleus, with contrast adjusted to clearly display labeling at the rim of the nucleus.
The loss of KinesinA and APR9 results in an accumulation of actin at the apical portion of the parasite upon stimulation with a calcium ionophore
Parasitic gliding motility is dependent on actin polymerization and associated myosins. The current model predicts an apical-basal actin flux powered by cortex-associated myosins [46]. In recent years, an mEmerald-tagged actin nanobody (“actin-chromobody”) that preferentially associates with F-actin has been used to assess actin kinetics in Toxoplasma [18,47–49]. The most notable observation is the buildup of a basal accumulation of mEmerald fluorescence, presumed to be actin-nanobody complexes when the parasite motility is stimulated. To determine how the loss of KinesinA and APR9 affects actin kinetics, we transiently expressed the mEmerald-actin-chromobody (mE-actin-Cb) in WT and ΔkinesinAΔapr9 parasites and treated intracellular parasites with A23187 to induce egress (Fig 6A-6C, S2-S4 Videos). In contrast to the WT parasite, where mE-actin-Cb builds up at the basal end of the parasite upon A23187 treatment, mE-actin-Cb accumulates in an apical cap in the ΔkinesinAΔapr9 parasite. Previously it was shown that the knockdown of APR2 results in apical mE-actin-Cb concentration when treated with BIPPO, an inhibitor of apicomplexan phosphodiesterases implicated in the Ca2+ signaling pathways [29,50]. We also observed the formation of a prominent apical mE-actin-Cb concentration when our APR2 knockout line (Δapr2) was treated with A23187 (Fig 6B). However, as we reported before [28], the Δapr2 parasites actively move out of the host cell during A23187-induced egress using gliding motility (S3 Video). This indicates that the buildup of an apical actin concentration cannot explain the motility defect in the ΔkinesinAΔapr9 parasites during parasite egress. To examine the parasite motile behavior in a host-cell-free environment, we carried out two-dimensional (2-D) gliding motility assay of WT, Δapr2, and ΔkinesinAΔapr9 parasites (Figs 6D, S6). We found that while long, non-circular trails were readily detectable in the WT samples (Fig 6D, yellow arrows), the longer trails deposited by the Δapr2 parasite tended to be circular in nature (yellow arrowheads). This might account for the modest invasion defect of the Δapr2 parasite [28]. The difference between the ΔkinesinAΔapr9 parasite and the other two lines was much more striking. Very few trails were detected in the ΔkinesinAΔapr9 parasite samples. Therefore, even though both the Δapr2 and ΔkinesinAΔapr9 parasites accumulate an apical actin cap upon A23187 treatment, their motile behaviors during egress and on a 2-D surface are qualitatively different.
A-C. Live imaging of RHΔku80 parental (WT, A), Δapr2 (B), and ΔkinesinAΔapr9 (C) parasites expressing actin-Cb-mE and treated with 5 µM A23187. Also see S2-S4 Videos. Yellow arrows: basal actin-Cb-ME accumulation. Cyan arrows: apical actin-Cb-mE accumulation. D. 2-D gliding motility assay of RHΔku80 parental (WT), Δapr2, and ΔkinesinAΔapr9 parasites. Long non-circular trails are readily detected in the WT sample (yellow arrows). The longer trails deposited by the Δapr2 parasite tend to be circular in nature (yellow arrowheads). The trail density of the ΔkinesinAΔapr9 parasite is markedly lower than those of both WT and Δapr2 parasites. See S6 Fig for two additional sets of images.
The loss of KinesinA and APR9 only has a minor impact on the organization of cortical MTs, but severely compromises conoid protrusion
Although APR9 and KinesinA are both early components of the apical polar ring, the loss of KinesinA and APR9 does not have a major impact on the organization of the cortical MTs (Fig 7A). Only ~15% of mature intracellular ΔkinesinAΔapr9 parasites have detached MTs or abnormal MT organization (total 177 parasites counted). To determine how APR9 and KinesinA impact the overall structure of the apical complex, we carried out negative staining EM analysis of TX-100 extracted parasites. Similar to the ΔkinesinA parasite [17], ΔkinesinAΔapr9 does not have a strongly stained annulus at the apical ends of the cortical MTs. There is no overt structural abnormality in the conoid of ΔkinesinAΔapr9 parasites (Fig 7B). The conoid is a motile organelle and extends upon calcium stimulation through the apical polar ring. To determine whether conoid protrusion is affected in the ΔkinesinAΔapr9 parasite and its parental lines, we treated extracellular parasites with A23187 for 1 min, fixed with formaldehyde and processed for expansion microscopy labeled with anti-tubulin antibody (Fig 7C). While over ~96% of the wild-type parasites were found with prominent, extended conoid (total 198 parasite counted), only ~ 31% of the ΔkinesinAΔapr9 parasites have a fully protruded conoid (total 249 parasites counted) (Fig 7C-7D). The conoid and the associated preconoidal rings contain multiple motility-related signal and structural proteins [14,15,18,19]. It is thus conceivable that blocking the movement of this structure loaded with motility-relevant factors could globally impact parasite motility. However, we found that only ~10% of the Δapr2 parasites fully extend the conoid (total 165 parasites counted) (Fig 7C- 7D), even though this knockout moved actively and persistently during A23187-induced egress (Fig 6B and [28]). The lack of conoid extension under this condition therefore cannot, by itself, explain the motility phenotype of the ΔkinesinAΔapr9 parasites.
A. Projections of ExM images of eight intracellular ΔkinesinAΔapr9 parasites labeled with an anti-tubulin antibody. Cortical MT organization appears normal in most parasites. The left inset (1X) is a projection of a substack close to the parasite surface and includes two detached cortical MTs. The right inset (2.5X) is a projection of a substack that includes the parasite apex, with the arrowhead indicating the gap between the cortical MTs and the conoid (“C”). B. Negative-staining EM images of TX-100 extracted RHΔku80 parental (WT), Δapr9, ΔkinesinA, and ΔkinesinAΔapr9 parasites. Arrowheads in the WT and Δapr9 images indicate the well-defined annulus, a readily recognizable feature of the apical polar ring. This annulus is undetectable in ΔkinesinA and ΔkinesinAΔapr9 parasites, although the cortical MTs remain converged at the parasite apex. Two retracted conoids are indicated by brackets. C. Projections of ExM images of extracellular RHΔku80 parental (WT), ΔkinesinAΔapr9, and Δapr2 parasites after treated with 5 µM A23187 for 1 min and labeled with an anti-tubulin antibody. Insets (2X) include a projection of a substack of the parasite apex to show the state of conoid extension. D. Bar graphs that quantify percentage of extended (blue), partially extended (green), and retracted (yellow) conoids in the extracellular RHΔku80 parental (WT), Δapr9, ΔkinesinAΔapr9, ΔkinesinAΔapr9:APR9 complement (comp) and Δapr2 parasites after treatment with 5 µM A23187 for 1 min.
The loss of KinesinA and APR9 results in significantly reduced secretion of the major micronemal adesin, MIC2
In order for the parasite to glide, the internal force generated by myosins traveling on the polymerizing F-actin needs to be relayed by associated proteins through transmembrane adhesins. The resulting cortical force powers the parasite gliding on a surface. The major adhesin secreted by the parasite is the micronemal protein MIC2, which forms a complex with M2AP to mediate parasite gliding during invasion and egress [51–54]. The localization of MIC2 appears to be normal in intracellular ΔkinesinAΔapr9 parasites (Fig 8A). Interestingly, in extracellular parasites treated with A23187, MIC2-containing vesicles are found within the extended conoid of the WT parasites, as well as within the retracted and partially extended conoid in the ΔkinesinAΔapr9 parasites (Figs 8B, S7). We then examined MIC2 secretion with and without A23187 treatment by Western blot (Fig 8C-8E). We first tested MIC2 secretion in DMEM parasite growth media (Fig 8C-8D). Under this condition, the parental lines (WT, Δapr9, ΔkinesinA:mE-APR9 knock-in) and the ΔkinesinAΔapr9:mE-APR9 complement displayed robust MIC2 secretion upon A23187 treatment. In contrast, MIC2 secretion from the ΔkinesinAΔapr9 parasites was much reduced. Because the egress assays revealing the profound motility defect of the ΔkinesinAΔapr9 parasites was carried out in an L15-based imaging medium, we also examined MIC2 secretion of the ΔkinesinAΔapr9 and WT parasites in this medium. Although the secretion of ΔkinesinAΔapr9 parasites remained significantly lower than that of the WT parasite, both parasite lines secreted more robustly in the L15 imaging medium (Fig 8E, S2 Table). Previous reports showed that the complete removal of MIC2 results in a major defect in active dispersion during A23187-induced egress [54]. However, even ~ 5% of WT MIC2 expression can support active motility during induced egress [53]. Therefore, factors other than or in addition to reduced bulk MIC-secretion underlie the motility phenotype of the ΔkinesinAΔapr9 parasites during A23187-induced egress.
A. Anti-MIC2 immunofluorescence labeling of intracellular RHΔku80 parental (WT) and ΔkinesinAΔapr9 parasites. B. Projections of ExM images of extracellular RHΔku80 parental (WT) and ΔkinesinAΔapr9 parasites after treated with 5 µM A23187 for 1 min and labeled with an anti-tubulin (red) and anti-MIC2 (green) antibodies. Insets (3X) include a single section of the conoid region. See S7 Fig for magenta-green version of the images. C. Western blot analysis of the secreted and pellet fractions of RHΔku80 parental (WT), ∆kinesinA:mE-APR9 knock-in, Δapr9, ΔkinesinAΔapr9 (ΔkinAΔapr9), ΔkinesinAΔapr9:mE-APR9 complement (comp), and Δapr2 parasites with (+) or without (-) A23187 treatment in DMEM growth medium. MIC2 was detected using the mouse anti-MIC2 antibody 6D10 [69]. Tubulin in the pellet fraction, detected by an anti-tubulin antibody, served as the loading control. M: molecular weight markers (kDa). Image contrast was inverted and enhanced to facilitate visualization. D. Quantification of the experiments shown in C. Values represent background-subtracted intensities of the secreted MIC2 bands in the A23187-induced samples relative to WT, normalized to the corresponding tubulin loading control in the pellet fraction. Negative background-subtracted values were set to zero. Three independent biological replicates were carried out for Δapr2 parasite, and five for the other lines. E. Western blot analysis of the secreted and pellet fractions of RHΔku80 parental (WT) and ΔkinesinAΔapr9 parasites with (+) or without (-) A23187 treatment in DMEM growth medium (D) and L15 imaging medium (L). See S2 Table for quantification.
Discussion
The apical complex is an ancestral feature of the apicomplexans. It is shared by not only the thousands of apicomplexan parasites, but also their free-living relatives, such as the chromerids and colpodellids. Structural details of the cytoskeletal apical complex, such as the presence or absence of a complete conoid, varies across different apicomplexan lineages [5,55]. However, an apical polar ring coupled with cortical MTs is found in all apicomplexans for which ultrastructural data is available. The conserved nature of the apical polar ring and its relevance to multiple aspects of parasite biology underlie the need for a deeper understanding of its composition and function.
Work in Toxoplasma gondii so far has identified more than a dozen components of the apical polar ring [16,17,25–31]. The probable extensive and complex interaction among these components is for the most part still unknown. For instance, we used APR2 as the bait in our immunoprecipitation protocol that led to the identification of APR9, yet the localization of APR9 is not perturbed in parasite lacking APR2. No doubt many other examples of these second order association will be found as more apical polar ring components are identified.
Interestingly, the known components of the apical polar ring are often poorly conserved outside Coccidia (Fig 3A). APR9, on the other hand, is not only broadly conserved among the apicomplexans, but also found in the free-living Chromera velia. Additionally, it has a close homolog, APR4, in Toxoplasma. The P. berghei ortholog of APR4/9 is predicted to be highly fitness-conferring (denoted “essential” in [56] and PlasmoDB). However, in Toxoplasma, the Δapr9, Δapr4, Δapr4Δapr9 and Δapr2Δapr9 parasites all have only modest defects in the lytic cycle (this work and [29]). One might wonder under what conditions these conserved proteins perform functions sufficiently beneficial to the parasites to necessitate their retention across a wide range of apicomplexans and related lineages. One plausible explanation is a synergistic action between the apical polar ring components. In Toxoplasma, this effect is particularly pronounced when an additional apical polar ring component is removed from the ΔkinesinAparasite. So far we have generated ΔkinesinAΔapr1, ΔkinesinAΔapr2, and ΔkinesinAΔapr9 parasites. The defects of all of these double-knockouts are far more severe than if the impact of the target genes were simply additive ([17,28] and this work).
The ΔkinesinAΔapr9 parasite displays a pronounced motility defect. Most parasites appear to be paralyzed during A23187-induced egress. At the same time, the ΔkinesinAΔapr9 parasite develops an apical actin concentration, as visualized by mE-actin-Cb. This, however, cannot explain the motility defect in the ΔkinesinAΔapr9 parasite, since the Δapr2 parasites, which also exhibit the accumulation of an apical actin cap, are motile and egress normally upon A23187 treatment. The Δapr2 and the ΔkinesinAΔapr9 parasites also both display a defect in conoid protrusion. This suggests that the apical polar ring is important for the mobility of the conoid. Together with the previously reported role of the apical polar ring in conoid attachment and positioning [17,29], multiple aspects of the structural and function dependence of these two macromolecular complexes have now emerged. Conoid protrusion, however, is not tightly coupled with parasite motility, as the Δapr2 parasite is highly impaired in conoid extension but actively moves out of the host cell during A23187-induced egress.
To move by gliding, the parasite needs to adhere to the surface that it interacts with. MIC2 has been proposed to be the major adhesin that connects the internal actomyosin machinery with the surface that the parasite glides on. Reduced MIC2 secretion results in a significant defect in parasite invasion [53]. Previous studies revealed that several components of the apical polar ring contribute to the secretion of MIC2, including RNG2, and KinesinA and APR1, which act synergistically [16,17,57]. This work shows a large reduction of A23187-induced MIC2 secretion from the ΔkinesinAΔapr9 parasite, which likely contributes to the invasion defect of this double-knockout and reaffirms the role of the apical polar ring in MIC2 secretion. Interestingly, the three apex activities investigated in this study involve overlapping but distinct molecular contributions from the apical polar ring. For example, KinesinA + APR9 and APR2 are both involved in conoid protrusion and apical actin distribution. However, KinesinA + APR9, but not APR2, are important for induced MIC2 secretion. Notably, unlike in the ΔkinesinAΔapr1 parasite [17], the organization of cortical MTs is largely unaffected in the ΔkinesinAΔapr9 parasite. This indicates that the function of the apical polar ring in controlling MIC2 secretion is independent of its role as an MT organizing center. Ultrastructural studies show that the apical complex is structurally coupled to secretory organelles [8,23,58,59]. The mechanics of micronemal secretion, however, remains elusive. The apical polar ring might facilitate protein release by interacting with surface structures on micronemal vesicles. Further studies that explore if and how KinesinA and functionally connected proteins such as APR1 and APR9 interact with micronemal vesicles to control secretion will provide useful insights. We note that the bulk secretion of MIC2 from ΔkinesinAΔapr9 parasite, although considerably reduced, is still detectable, especially in L15 imaging medium, where this double-knockout displays profound motility defects during A23187-induced egress. Given that even a small residual level of MIC2 expression can support active parasite movement during induced egress [53], we propose that the motility defect in ΔkinesinAΔapr9 parasite is a consequence of combined effects of abnormalities in actin kinetics, conoid protrusion and micronemal secretion. Alternatively, the apical polar ring dictates another, so far unknown, aspect of motility control through KinesinA and APR9.
Interestingly, while both KinesinA and APR9 are early components of the apical polar ring, their loss perturbs microtubule organization in only a minority of the parasites. Nevertheless, the broad conservation of APR9 among the apicomplexans is of great interest, especially in the free-living Chromera velia. We believe that it is a useful probe for structurally defining the counterpart of the apical polar ring in Chromera and exploring its construction in conjunction with other apical complex-related structures, such as the pseudoconoid. The role of APR9 and other conserved motility-relevant factors (e.g., AKMT [14,60,61]) associated with the apical complex also pose the questions regarding how the apical complex was involved in the origination of gliding motility, whether this behavior predates the development of intracellular parasitism, and if so, how it was utilized in the life of the free-living ancestor of apicomplexans, the motile form of which was likely to be flagellated like Chromera.
Materials and methods
T. gondii cultures
As described in [17,19,62,63], T. gondii tachyzoites were maintained in confluent cultures of human foreskin fibroblasts (HFFs) in DMEM growth medium [Dulbecco’s Modified Eagle’s Medium (Corning, 15–013-CV) supplemented with 1% (v/v) heat-inactivated cosmic calf serum (SH30087.3; Hyclone, Logan, UT) and Glutamax (Life Technologies-Gibco, 35050061)].
Immunoprecipitation and Multidimensional Protein Identification Technology (MudPIT) analysis
Immunoprecipitation experiments were performed as described in [64] using the mEmerald-APR2 knock-in parasites [28]. Untagged RHΔhxΔku80 parasites (a kind gift from Dr. Vern Carruthers at the University of Michigan [65]) was used as the negative control. Parasites were processed as described in [64], except that the lysis buffer contained 1% (instead of 0.5%) TX-100. The lysate was clarified by centrifugation and then incubated with Chromotek-GFP-Trap agarose beads (AB_2631357, Proteintech) for 90 minutes at 4°C before elution for MudPIT analysis.
Protein samples were processed for MudPIT and analyzed as described in [19,64]. Raw data and search results files have been deposited to the Proteome Xchange (accession: PXD074785) via the MassIVE repository and may be accessed at http://massive.ucsd.edu with MSV000100958 as the Dataset Identifier. Original mass spectrometry data underlying this manuscript can be accessed after publication from the Stowers Original Data Repository at http://www.stowers.org/research/publications/libpb-2610.
Plasmid construction (See S3 Table for primers used in this study)
Genomic DNA (gDNA) and coding sequences (CDS) were prepared as described in [19].
pTKO2_II-mEmerald-APR4 (mE-APR4) knock-in plasmid: ∼1.9 kb sequences upstream (5’UTR) or downstream (3’UTR) of the APR4 (TgGT1_ 219500) genomic locus were amplified from the parasite gDNA by PCR using primer pairs S1/ AS1 and S2/AS2, respectively, and inserted at the NotI (5’UTR) or HindIII (3’UTR) site of plasmid pTKO2-II-mCherryFP [64] using the NEBuilder HiFi Assembly kit. The coding sequences for mEmerald fluorescent protein (mEmerald) and APR4 were amplified using primer pairs S3/AS3 and S4/AS4, respectively, and assembled into the AsiSI site to generate pTKO2_II-mE-APR4. A five-amino acid linker (SGLRS) was inserted between the APR4 and mEmerald coding sequences, and the Kozak sequence from the endogenous APR4 locus (CCAATGG) was added to the 5’ end of the mEmerald coding sequence.
pTKO2_II- mE-APR9, pTKO2_II- mE-APR10, and pTKO2_II- mE-APR11 knock-in plasmids: These plasmids were constructed using the same strategy as described for pTKO2_II-mE-APR4 knock-in using the primers listed in S3 Table. The endogenous Kozak sequences added to the 5’ end of the mEmerald coding sequence were ACCATGA, GCGATGG, and GCGATGA for APR9 (TgGT1_ 223790), APR10 (TgGT1_ 289990), and APR11 (TgGT1_ 295420), respectively.
Generation of knock-in and knockout parasite lines
mE-APR4, mE-APR9, mE-APR10 and mE-APR11 knock-in lines: All knock-in and knockout lines were generated in the RHΔhxΔku80 strain, referred to as “RHΔku80”, “wild-type” or “WT” throughout this study. ~ 1 x 107 RHΔhxΔku80 parasites were electroporated with 40 μg of the knock-in plasmid linearized by NotI, using the settings described previously [17,19,62]. Transfected populations were then selected with 25 μg/mL mycophenolic acid and 50 μg/mL xanthine. Because the backbone of the pTKO2_II plasmid contains a cassette driving cytoplasmic expression of mCherryFP [64], clones were first selected by positive mEmeraldFP fluorescence at the apical polar ring as well as absence of cytoplasmic mCherry fluorescence, allowing exclusion of non-homologous or single crossover recombinants. Correct homologous integrations were subsequently verified by Southern blot (for the mE-APR4 and mE-APR9 knock-in lines) or by genomic PCRs (for the mE-APR10 and mE-APR11 knock-in lines).
Δapr4 and Δapr9 lines: mE-APR4 or mE-APR9 knock-in clones confirmed by Southern blot were electroporated with 30 μg of pmin-Cre-eGFP_Gra-mCherry, and selected with 6-thioxanthine at 80 μg/mL as described previously [19,62]. Clones that had lost the mEmerald fluorescence were first identified by microscopic screening. The deletion of the LoxP-flanked region in the selected clones was then confirmed by Southern blot.
Δapr4:mE-APR9 knock-in, Δapr2:mE-APR9 knock-in, andΔkinesinA:mE-APR9 knock-in lines: The Δapr4, Δapr2 [28], or ΔkinesinA parasites [17] were electroporated with the NotI-linearized pTKO2_II-mE-APR9 knock-in plasmid. Knock-in clones were selected and confirmed as described above for the mE-APR9 knock-in line.
Δapr4Δapr9, Δapr2Δapr9, and ΔkinesinAΔapr9 lines: Δapr4:mE-APR9 knock-in, Δapr2:mE-APR9 knock-in, or ΔkinesinA:mE-APR9 knock-in parasites were electroporated with pmin-Cre-eGFP_Gra-mCherry. Clones were selected as described above for the Δapr9 parasite.
Southern blotting
Southern blotting was carried out as described in [19,28,62,64]. To probe and detect changes at the target genomic locus in parental (RHΔku80), knock-in and knockout parasites, 5 µg of gDNA from each line was digested with the restriction enzymes indicated in Figs 4A-4B and S4 before hybridization with probes for the 5’UTR, CDS, or 3’UTR regions.
For the APR4-related probes, templates for probe synthesis were generated from the pTKO2_II-mE-APR4 knock-in plasmid by restriction digestions followed by gel purification. The template for the 5’UTR probe was released by NotI and AvrII digestion. The template for the CDS probe was released by MreI and RsrII digestion. The template for the 3’UTR probe was released by SgrDI and PspOMI digestion. After gel purification, probes were synthesized from the templates by nick translation in the presence of all four dNTPs plus biotin-dATP.
For the APR9-related probes, the following templates for probe synthesis were generated from the pTKO2_II-mE-APR9 knock-in plasmid by restriction digestions: the template for the 5’UTR probe was released by MfeI and EcoRV digestion; the template for CDS probe 1 was released by SmaI and NgoMIV digestion; the template for the 3’UTR probe was released by Bstz17I and NheI digestion. CDS probe 2 was amplified from the pTKO2_II-mE-APR9 knock-in plasmid by PCR using the primer pair S17/AS17.
Sample preparation and imaging for Expansion Microscopy (ExM)
Expanded samples of intracellular and extracellular parasites were prepared, labeled with anti-tubulin and anti-GFP antibodies, and imaged with a DeltaVision OMX Flex imaging station (GE Healthcare-Applied Precision) as previously described [28,32,66].
For assessing conoid protrusion, freshly egressed extracellular parasites were treated with 5 μM A23187 for 1 minute at room temperature while settling on a poly-L-lysine-coated coverslip. Samples were then fixed and processed for expansion microscopy as previously described [28,32,66].
Sum or maximum projections were presented in the figures with contrast levels adjusted for optimization of display. The expansion ratio of ~ 5.4 was estimated based on [32].
Plaque assay
Freshly harvested parasites (100 or 200 per well) were used to infect confluent HFF monolayers in 6-well plates. After incubation at 37°C for seven or twelve days, the cultures were rinsed, fixed, stained, and scanned as described in [19]. Three to seven independent experiments were performed for each parasite line.
Invasion assays
Immunofluorescence-based invasion assays were carried out as described in [19,28] with five biological replicates. For each strain, parasites in 10 fields were counted per biological replicate. P-values for “%WT” were calculated in KaleidaGraph using two-tailed unpaired Student’s t-tests with unequal variance.
Replication assay
Intracellular replication assays were performed as described in [14], with replication rates calculated as described in [67]. Three or six independent experiments were performed for each parasite line. For each strain and time point, parasites in ~ 100 vacuoles were counted for each replicate.
A23187-induced egress
Calcium ionophore-induced egress assays were performed as described in [19] using 5 μM A23187 in L15 imaging medium [Leibovitz’s L-15 (21083–027, Gibco- Life Technologies, Grand Island, NY) supplemented with 1% (vol/vol) cosmic calf serum].
2-D gliding motility assay
Gliding motility assays were performed as previously described in [38] with modifications. Briefly, glass-bottom dishes (P35G-1.5-14-C; MatTek, Ashland, MA) were coated with poly-L-lysine (Sigma-Aldrich, P4707) for 30 min at room temperature, washed 3 times with PBS, and allowed to air dry completely before use. Freshly egressed T. gondii parasites were harvested, centrifuged at 6000 rpm for 1 min, and resuspended in calcium saline solution at a concentration of 1 × 106 parasites per sample. The calcium saline solution consisted of 137 mM NaCl, 5mM KCl, 1 mM Na₂HPO₄, 5.5 mM glucose, and 21 mM HEPES, 5 mM CaCl₂ (pH ~ 7). Parasites were added to the poly-L-lysine-coated dishes and incubated at 37°C for 30 min to allow gliding. Samples were then gently washed once with PBS and fixed with 4% paraformaldehyde and 0.06% glutaraldehyde in PBS for 15 min at room temperature. Following fixation, samples were washed with PBS and blocked with 1.5% BSA in PBS for 30 min, then incubated with a rabbit anti-P30 antibody (a kind gift from Dr. Lloyd Kasper, Darthmouth College, MA) at 1:1000 for 30 min. After three gentle washes with PBS, samples were incubated with an Alexa Fluor 488-conjugated goat anti-rabbit secondary antibody (Invitrogen, A11034) for 30 min, washed three additional times with PBS, and imaged using a 40X objective lens.
Electron microscopy
Suspensions of extracellular parasites were treated with A23187 and processed for negative staining as described in [19]. Samples were imaged on a Talos transmission electron microscope (Thermo Fisher) operated at 120 keV.
Microneme immunofluorescence and secretion assays
Immunofluorescence labeling of micronemes in intracellular parasites was performed as previously described [19]. For assessing microneme labeling in extracellular parasites using expansion microscopy, parasites treated with 5 μM A23187 and processed as described above for expansion were labeled with a mouse-anti-tubulin (1:250, T6793-6-11B-1, Sigma-Aldrich) and a rat-anti-MIC2 antibodies (1:200, a kind gift from Dr. Vern Carruthers at the University of Michigan), and corresponding secondary antibodies [goat anti-mouse Alexa568 (1:400, A11031, Invitrogen) and goat anti-rat Alexa488 (1:200, A11006, Invitrogen)] for immunofluorescence.
For microneme secretion assays, freshly egressed parasites were collected and treated with 5 μM A23187 as described in [68] in DMEM growth medium or L15 imaging medium. Secreted and pellet fractions were analyzed by Western blot as described in [68].
Supporting information
S1 Video. Time-lapse microscopy of RHΔku80 parental (WT), ΔkinesinAΔapr9 (kinAapr9DKO), and ΔkinesinAΔapr9:mE-APR9 complement (DKO:mE-APR9 comp) parasites.
The cell impermeant DNA binding dye DAPI was added to the culture medium prior to the start of the experiment. The parasite-dependent host cell-permeabilization was detected by the DNA labeling by DAPI entering the host cell nucleus. Time is shown in min:s, Video speed: 12 frames/s. Scale bar: 5 µm.
https://doi.org/10.1371/journal.ppat.1014378.s001
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S2 Video. Time-lapse microscopy of RHΔku80 parental (WT) parasites expressing actin-Cb-mE.
Time is shown in min:s, Video speed: 6 frames/s.
https://doi.org/10.1371/journal.ppat.1014378.s002
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S3 Video. Time-lapse microscopy of Δapr2 (apr2 KO) parasites expressing actin-Cb-mE.
Time is shown in min:s, Video speed: 6 frames/s.
https://doi.org/10.1371/journal.ppat.1014378.s003
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S4 Video. Time-lapse microscopy of ΔkinesinAΔapr9 (kinAapr9DKO) parasites expressing actin-Cb-mE.
Time is shown in min:s, Video speed: 6 frames/s.
https://doi.org/10.1371/journal.ppat.1014378.s004
(AVI)
S1 Fig. Magenta-green version of the images in Fig 2C-2E.
https://doi.org/10.1371/journal.ppat.1014378.s005
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S2 Fig. Predicted structures of TgAPR9, TgAPR4, and orthologs from Plasmodium berghei and Chromera velia by AlphaFold3.
The numbers of aligned residues were generated by TM-align (Version 20190822, Zhang, Y. and J. Skolnick, 2005. TM-align: a protein structure alignment algorithm based on the TM-score. Nucleic Acids Res 33, 2302–2309.).
https://doi.org/10.1371/journal.ppat.1014378.s006
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S3 Fig. Magenta-green version of the images in Fig 3C-3H.
https://doi.org/10.1371/journal.ppat.1014378.s007
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S4 Fig. Southern blot analysis of the apr4 locus in the WT, mEmeraldFP-APR4 (mE-APR4 KI) and Δapr4parasites.
A. Schematic for the predicted APR4 locus in WT, mE- tagged APR4 knock-in, and Δapr4 lines. Restriction sites, hybridization targets of the Southern blot probes for the apr4 coding region (CDS probe, orange bar), regions upstream (“5’ UTR probe”, blue bar) and downstream (“3’ UTR probe”, purple bar) of the CDS, and the corresponding DNA fragment sizes expected are indicated. B. Southern blots confirmed the homologous integration of the mE-APR4 fusion in the knock-in line, and the deletion of the apr4 locus in the Δapr4 lines.
https://doi.org/10.1371/journal.ppat.1014378.s008
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S5 Fig. Magenta-green version of the images in Fig 4C-4E.
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S6 Fig. Two additional sets of images of 2-D gliding motility assay of RHΔku80parental (WT), Δapr2, and ΔkinesinAΔapr9 parasites.
https://doi.org/10.1371/journal.ppat.1014378.s010
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S7 Fig. Magenta-green version of the images in Fig 8B.
https://doi.org/10.1371/journal.ppat.1014378.s011
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S1 Table. List of proteins identified in MudPIT analysis of an immunoprecipitation using GFP-Trap and lysate from the mEmeraldFP-APR2 knock-in line.
The RHΔku80 parental parasite with untagged APR2 was used as the negative control.
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S2 Table. Quantification of A23187-induced MIC2 secretion of WT and ΔkinesinAΔapr9 parasites in DMEM growth medium and L15 imaging medium.
Values represent background-subtracted intensities of the secreted MIC2 band in the A23187-induced samples relative to WT parasites in DMEM growth medium, normalized to the corresponding tubulin loading control in the pellet fraction.
https://doi.org/10.1371/journal.ppat.1014378.s013
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S3 Table. List of primers used in this study.
https://doi.org/10.1371/journal.ppat.1014378.s014
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Acknowledgments
We thank Matea Susac for tissue culture support and helpful discussions and the EM facility at Arizona State University for instrumentation support.
References
- 1. Seeber F, Steinfelder S. Recent advances in understanding apicomplexan parasites. F1000Res. 2016;5. pmid:27347391
- 2. Checkley W, White AC Jr, Jaganath D, Arrowood MJ, Chalmers RM, Chen X-M, et al. A review of the global burden, novel diagnostics, therapeutics, and vaccine targets for cryptosporidium. Lancet Infect Dis. 2015;15(1):85–94. pmid:25278220
- 3. Torgerson PR, Mastroiacovo P. The global burden of congenital toxoplasmosis: a systematic review. Bull World Health Organ. 2013;91(7):501–8. pmid:23825877
- 4.
World Health Organization. World Malaria Report 2022. 2022.
- 5. Morrissette NS, Sibley LD. Cytoskeleton of apicomplexan parasites. Microbiol Mol Biol Rev. 2002;66(1):21–38; table of contents. pmid:11875126
- 6. Nichols BA, Chiappino ML. Cytoskeleton of Toxoplasma gondii. J Protozool. 1987;34(2):217–26. pmid:3585817
- 7. Morrissette NS, Murray JM, Roos DS. Subpellicular microtubules associate with an intramembranous particle lattice in the protozoan parasite Toxoplasma gondii. J Cell Sci. 1997;110 (Pt 1):35–42. pmid:9010782
- 8. Hu K, Roos DS, Murray JM. A novel polymer of tubulin forms the conoid of Toxoplasma gondii. J Cell Biol. 2002;156(6):1039–50. pmid:11901169
- 9. Russell DG, Burns RG. The polar ring of coccidian sporozoites: a unique microtubule-organizing centre. J Cell Sci. 1984;65:193–207. pmid:6715423
- 10. Wang X, Fu Y, Beatty WL, Ma M, Brown A, Sibley LD, et al. Cryo-EM structure of cortical microtubules from human parasite Toxoplasma gondii identifies their microtubule inner proteins. Nat Commun. 2021;12(1):3065. pmid:34031406
- 11. Sun SY, Segev-Zarko L-A, Chen M, Pintilie GD, Schmid MF, Ludtke SJ, et al. Cryo-ET of Toxoplasma parasites gives subnanometer insight into tubulin-based structures. Proc Natl Acad Sci U S A. 2022;119(6):e2111661119. pmid:35121661
- 12. Li Z, Du W, Yang J, Lai D-H, Lun Z-R, Guo Q. Cryo-Electron Tomography of Toxoplasma gondii indicates that the conoid fiber may be derived from microtubules. Adv Sci (Weinh). 2023;10(14):e2206595. pmid:36840635
- 13. Hu K, Johnson J, Florens L, Fraunholz M, Suravajjala S, DiLullo C, et al. Cytoskeletal components of an invasion machine--the apical complex of Toxoplasma gondii. PLoS Pathog. 2006;2(2):e13. pmid:16518471
- 14. Heaslip AT, Nishi M, Stein B, Hu K. The motility of a human parasite, Toxoplasma gondii, is regulated by a novel lysine methyltransferase. PLoS Pathog. 2011;7(9):e1002201. pmid:21909263
- 15. Graindorge A, Frénal K, Jacot D, Salamun J, Marq JB, Soldati-Favre D. The conoid associated motor MyoH is indispensable for Toxoplasma gondii entry and exit from host cells. PLoS Pathog. 2016;12(1):e1005388. pmid:26760042
- 16. Katris NJ, van Dooren GG, McMillan PJ, Hanssen E, Tilley L, Waller RF. The apical complex provides a regulated gateway for secretion of invasion factors in Toxoplasma. PLoS Pathog. 2014;10(4):e1004074. pmid:24743791
- 17. Leung JM, He Y, Zhang F, Hwang Y-C, Nagayasu E, Liu J, et al. Stability and function of a putative microtubule-organizing center in the human parasite Toxoplasma gondii. Mol Biol Cell. 2017;28(10):1361–78. pmid:28331073
- 18. Tosetti N, Dos Santos Pacheco N, Soldati-Favre D, Jacot D. Three F-actin assembly centers regulate organelle inheritance, cell-cell communication and motility in Toxoplasma gondii. Elife. 2019;8:e42669. pmid:30753127
- 19. Munera Lopez J, Tengganu IF, Liu J, Murray JM, Arias Padilla LF, Zhang Y, et al. An apical protein, Pcr2, is required for persistent movement by the human parasite Toxoplasma gondii. PLoS Pathog. 2022;18(8):e1010776. pmid:35994509
- 20. Dos Santos Pacheco N, Brusini L, Haase R, Tosetti N, Maco B, Brochet M, et al. Conoid extrusion regulates glideosome assembly to control motility and invasion in Apicomplexa. Nat Microbiol. 2022;7(11):1777–90. pmid:36109645
- 21. Stadler RV, White LA, Hu K, Helmke BP, Guilford WH. Direct measurement of cortical force generation and polarization in a living parasite. Mol Biol Cell. 2017;28(14):1912–23. pmid:28209732
- 22. Nagayasu E, Hwang Y-C, Liu J, Murray JM, Hu K. Loss of a doublecortin (DCX)-domain protein causes structural defects in a tubulin-based organelle of Toxoplasma gondii and impairs host-cell invasion. Mol Biol Cell. 2017;28(3):411–28. pmid:27932494
- 23. Gui L, O’Shaughnessy WJ, Cai K, Reetz E, Reese ML, Nicastro D. Cryo-tomography reveals rigid-body motion and organization of apicomplexan invasion machinery. Nat Commun. 2023;14(1):1775. pmid:36997532
- 24. Zeng J, Fu Y, Qian P, Huang W, Niu Q, Beatty WL, et al. Atomic models of the Toxoplasma cell invasion machinery. Nat Struct Mol Biol. 2026;33(1):157–70. pmid:41366525
- 25. Tran JQ, de Leon JC, Li C, Huynh M-H, Beatty W, Morrissette NS. RNG1 is a late marker of the apical polar ring in Toxoplasma gondii. Cytoskeleton (Hoboken). 2010;67(9):586–98. pmid:20658557
- 26. Barylyuk K, Koreny L, Ke H, Butterworth S, Crook OM, Lassadi I, et al. A comprehensive subcellular atlas of the Toxoplasma proteome via hyperLOPIT provides spatial context for protein functions. Cell Host Microbe. 2020;28(5):752–766.e9. pmid:33053376
- 27. Koreny L, Zeeshan M, Barylyuk K, Tromer EC, van Hooff JJE, Brady D, et al. Molecular characterization of the conoid complex in Toxoplasma reveals its conservation in all apicomplexans, including Plasmodium species. PLoS Biol. 2021;19(3):e3001081. pmid:33705380
- 28. Arias Padilla LF, Munera Lopez J, Shibata A, Murray JM, Hu K. The initiation and early development of apical-basal polarity in Toxoplasma gondii. J Cell Sci. 2024;137(19):jcs263436. pmid:39239869
- 29. Ren B, Haase R, Patray S, Nguyen Q, Maco B, Dos Santos Pacheco N, et al. Architecture of the Toxoplasma gondii apical polar ring and its role in gliding motility and invasion. Proc Natl Acad Sci U S A. 2024;121(46):e2416602121. pmid:39514309
- 30. Burette M, Pezinsky M, Nguyen Q, Benharoual A, Patray S, Maynadier M, et al. Two anchoring proteins control daughter apical complex assembly in Toxoplasma gondii. bioRxiv. 2026.
- 31. Li W, Koczy O, Qin P, Forné I, Gras S, Grünert J, et al. An apical ring protein essential for conoid complex assembly and daughter cell formation in Toxoplasma gondii. Nat Commun. 2025;16(1):10149. pmid:41315181
- 32. Padilla LFA, Murray JM, Hu K. The initiation and early development of the tubulin-containing cytoskeleton in the human parasite Toxoplasma gondii. Mol Biol Cell. 2024;35(3):ar37. pmid:38170577
- 33. Haase R, Puthenpurackal A, Maco B, Guérin A, Soldati-Favre D. γ-tubulin complex controls the nucleation of tubulin-based structures in Apicomplexa. Mol Biol Cell. 2024;35(9):ar121. pmid:39046777
- 34. Engelberg K, Bauwens C, Ferguson DJP, Gubbels M-J. Co-dependent formation of the Toxoplasma gondii subpellicular microtubules and inner membrane skeleton. mBio. 2025;16(9):e0138925. pmid:40801525
- 35. Florens L, Washburn MP, Raine JD, Anthony RM, Grainger M, Haynes JD, et al. A proteomic view of the Plasmodium falciparum life cycle. Nature. 2002;419(6906):520–6. pmid:12368866
- 36. Florens L, Korfali N, Schirmer EC. Subcellular fractionation and proteomics of nuclear envelopes. Methods Mol Biol. 2008;432:117–37. pmid:18370014
- 37. Florens L, Liu X, Wang Y, Yang S, Schwartz O, Peglar M, et al. Proteomics approach reveals novel proteins on the surface of malaria-infected erythrocytes. Mol Biochem Parasitol. 2004;135(1):1–11. pmid:15287581
- 38. Roumégous C, Abou Hammoud A, Fuster D, Dupuy J-W, Blancard C, Salin B, et al. Identification of new components of the basal pole of Toxoplasma gondii provides novel insights into its molecular organization and functions. Front Cell Infect Microbiol. 2022;12:1010038. pmid:36310866
- 39. Anderson-White BR, Ivey FD, Cheng K, Szatanek T, Lorestani A, Beckers CJ, et al. A family of intermediate filament-like proteins is sequentially assembled into the cytoskeleton of Toxoplasma gondii. Cell Microbiol. 2011;13(1):18–31. pmid:20698859
- 40. Dubey R, Harrison B, Dangoudoubiyam S, Bandini G, Cheng K, Kosber A, et al. Differential roles for inner membrane complex proteins across Toxoplasma gondii and Sarcocystis neurona development. mSphere. 2017;2:e00409–17.
- 41. Seidi A, Muellner-Wong LS, Rajendran E, Tjhin ET, Dagley LF, Aw VY, et al. Elucidating the mitochondrial proteome of Toxoplasma gondii reveals the presence of a divergent cytochrome c oxidase. Elife. 2018;7:e38131. pmid:30204084
- 42. Abramson J, Adler J, Dunger J, Evans R, Green T, Pritzel A, et al. Accurate structure prediction of biomolecular interactions with AlphaFold 3. Nature. 2024;630(8016):493–500. pmid:38718835
- 43. Sidik SM, Huet D, Ganesan SM, Huynh M-H, Wang T, Nasamu AS, et al. A genome-wide CRISPR screen in Toxoplasma identifies essential Apicomplexan Genes. Cell. 2016;166(6):1423–1435.e12. pmid:27594426
- 44. Mital J, Schwarz J, Taatjes DJ, Ward GE. Laser scanning cytometer-based assays for measuring host cell attachment and invasion by the human pathogen Toxoplasma gondii. Cytometry A. 2006;69(1):13–9. pmid:16342112
- 45. Carey KL, Westwood NJ, Mitchison TJ, Ward GE. A small-molecule approach to studying invasive mechanisms of Toxoplasma gondii. Proc Natl Acad Sci U S A. 2004;101(19):7433–8. pmid:15123807
- 46. Opitz C, Soldati D. “The glideosome”: a dynamic complex powering gliding motion and host cell invasion by Toxoplasma gondii. Mol Microbiol. 2002;45(3):597–604. pmid:12139608
- 47. Periz J, Whitelaw J, Harding C, Gras S, Del Rosario Minina MI, Latorre-Barragan F, et al. Toxoplasma gondii F-actin forms an extensive filamentous network required for material exchange and parasite maturation. Elife. 2017;6:e24119. pmid:28322189
- 48. Del Rosario M, Periz J, Pavlou G, Lyth O, Latorre-Barragan F, Das S, et al. Apicomplexan F-actin is required for efficient nuclear entry during host cell invasion. EMBO Rep. 2019;20(12):e48896. pmid:31584242
- 49. Periz J, Del Rosario M, McStea A, Gras S, Loney C, Wang L, et al. A highly dynamic F-actin network regulates transport and recycling of micronemes in Toxoplasma gondii vacuoles. Nat Commun. 2019;10(1):4183. pmid:31519913
- 50. Howard BL, Harvey KL, Stewart RJ, Azevedo MF, Crabb BS, Jennings IG, et al. Identification of potent phosphodiesterase inhibitors that demonstrate cyclic nucleotide-dependent functions in apicomplexan parasites. ACS Chem Biol. 2015;10(4):1145–54. pmid:25555060
- 51. Rabenau KE, Sohrabi A, Tripathy A, Reitter C, Ajioka JW, Tomley FM, et al. TgM2AP participates in Toxoplasma gondii invasion of host cells and is tightly associated with the adhesive protein TgMIC2. Mol Microbiol. 2001;41(3):537–47. pmid:11532123
- 52. Jewett TJ, Sibley LD. The toxoplasma proteins MIC2 and M2AP form a hexameric complex necessary for intracellular survival. J Biol Chem. 2004;279(10):9362–9. pmid:14670959
- 53. Huynh M-H, Carruthers VB. Toxoplasma MIC2 is a major determinant of invasion and virulence. PLoS Pathog. 2006;2(8):e84. pmid:16933991
- 54. Gras S, Jackson A, Woods S, Pall G, Whitelaw J, Leung JM, et al. Parasites lacking the micronemal protein MIC2 are deficient in surface attachment and host cell egress, but remain virulent in vivo. Wellcome Open Res. 2017;2:32. pmid:28630943
- 55. Harding CR, Frischknecht F. The riveting cellular structures of apicomplexan parasites. Trends Parasitol. 2020;36(12):979–91. pmid:33011071
- 56. Bushell E, Gomes AR, Sanderson T, Anar B, Girling G, Herd C, et al. Functional profiling of a Plasmodium genome reveals an abundance of essential genes. Cell. 2017;170(2):260–272.e8. pmid:28708996
- 57. Haase R, Ren B, Tell I Puig A, Bonavoglia A, Marq J-B, Visentin R, et al. RNG2 tethers the conoid to the apical polar ring in Toxoplasma gondii to enable parasite motility and invasion. PLoS Biol. 2025;23(11):e3003506. pmid:41284722
- 58. Paredes-Santos TC, de Souza W, Attias M. Dynamics and 3D organization of secretory organelles of Toxoplasma gondii. J Struct Biol. 2012;177(2):420–30. pmid:22155668
- 59. Aquilini E, Cova MM, Mageswaran SK, Dos Santos Pacheco N, Sparvoli D, Penarete-Vargas DM, et al. An Alveolata secretory machinery adapted to parasite host cell invasion. Nat Microbiol. 2021;6(4):425–34. pmid:33495622
- 60. Sivagurunathan S, Heaslip A, Liu J, Hu K. Identification of functional modules of AKMT, a novel lysine methyltransferase regulating the motility of Toxoplasma gondii. Mol Biochem Parasitol. 2013;189(1–2):43–53. pmid:23685344
- 61. Pivovarova Y, Liu J, Lesigang J, Koldyka O, Rauschmeier R, Hu K, et al. Structure of a novel dimeric SET domain methyltransferase that regulates cell motility. J Mol Biol. 2018;430(21):4209–29. pmid:30148980
- 62. Liu J, He Y, Benmerzouga I, Sullivan WJ Jr, Morrissette NS, Murray JM, et al. An ensemble of specifically targeted proteins stabilizes cortical microtubules in the human parasite Toxoplasma gondii. Mol Biol Cell. 2016;27(3):549–71. pmid:26680740
- 63. Roos DS, Donald RG, Morrissette NS, Moulton AL. Molecular tools for genetic dissection of the protozoan parasite Toxoplasma gondii. Methods Cell Biol. 1994;45:27–78.
- 64. Liu J, Wetzel L, Zhang Y, Nagayasu E, Ems-McClung S, Florens L, et al. Novel thioredoxin-like proteins are components of a protein complex coating the cortical microtubules of Toxoplasma gondii. Eukaryot Cell. 2013;12(12):1588–99. pmid:23873863
- 65. Huynh M-H, Carruthers VB. Tagging of endogenous genes in a Toxoplasma gondii strain lacking Ku80. Eukaryot Cell. 2009;8(4):530–9. pmid:19218426
- 66. Tengganu IF, Hu K. Transfection of the free-living alga Chromera velia enables direct comparisons with its parasitic apicomplexan relative, Toxoplasma gondii. J Cell Sci. 2026;139(1):jcs.264400. pmid:41208444
- 67. Fichera ME, Bhopale MK, Roos DS. In vitro assays elucidate peculiar kinetics of clindamycin action against Toxoplasma gondii. Antimicrob Agents Chemother. 1995;39(7):1530–7. pmid:7492099
- 68. Tengganu IF, Arias Padilla LF, Munera Lopez J, Liu J, Brown PT, Murray JM, et al. The cortical microtubules of Toxoplasma gondii underlie the helicity of parasite movement. J Cell Sci. 2023;136(17):jcs261270. pmid:37675776
- 69. Carruthers VB, Sibley LD. Sequential protein secretion from three distinct organelles of Toxoplasma gondii accompanies invasion of human fibroblasts. Eur J Cell Biol. 1997;73(2):114–23. pmid:9208224