Microtubule number and length determine cellular shape and function in Plasmodium
2019; Springer Nature; Volume: 38; Issue: 15 Linguagem: Inglês
10.15252/embj.2018100984
ISSN1460-2075
AutoresBenjamin Spreng, Hannah Fleckenstein, Patrick Kübler, Claudia Di Biagio, Madlen Benz, Pintu Patra, Ulrich S. Schwarz, Marek Cyrklaff, Friedrich Frischknecht,
Tópico(s)Malaria Research and Control
ResumoArticle24 May 2019Open Access Source DataTransparent process Microtubule number and length determine cellular shape and function in Plasmodium Benjamin Spreng Benjamin Spreng Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Hannah Fleckenstein Hannah Fleckenstein Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Patrick Kübler Patrick Kübler orcid.org/0000-0002-6079-5103 Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Claudia Di Biagio Claudia Di Biagio orcid.org/0000-0001-9096-3166 Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Madlen Benz Madlen Benz Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Pintu Patra Pintu Patra orcid.org/0000-0002-2275-3738 Institute for Theoretical Physics and Bioquant, Heidelberg University, Heidelberg, Germany Search for more papers by this author Ulrich S Schwarz Ulrich S Schwarz orcid.org/0000-0003-1483-640X Institute for Theoretical Physics and Bioquant, Heidelberg University, Heidelberg, Germany Search for more papers by this author Marek Cyrklaff Marek Cyrklaff Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Friedrich Frischknecht Corresponding Author Friedrich Frischknecht [email protected] orcid.org/0000-0002-8332-6668 Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Benjamin Spreng Benjamin Spreng Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Hannah Fleckenstein Hannah Fleckenstein Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Patrick Kübler Patrick Kübler orcid.org/0000-0002-6079-5103 Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Claudia Di Biagio Claudia Di Biagio orcid.org/0000-0001-9096-3166 Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Madlen Benz Madlen Benz Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Pintu Patra Pintu Patra orcid.org/0000-0002-2275-3738 Institute for Theoretical Physics and Bioquant, Heidelberg University, Heidelberg, Germany Search for more papers by this author Ulrich S Schwarz Ulrich S Schwarz orcid.org/0000-0003-1483-640X Institute for Theoretical Physics and Bioquant, Heidelberg University, Heidelberg, Germany Search for more papers by this author Marek Cyrklaff Marek Cyrklaff Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Friedrich Frischknecht Corresponding Author Friedrich Frischknecht [email protected] orcid.org/0000-0002-8332-6668 Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany Search for more papers by this author Author Information Benjamin Spreng1, Hannah Fleckenstein1,‡, Patrick Kübler1,‡, Claudia Di Biagio1,‡, Madlen Benz1,‡, Pintu Patra2,‡, Ulrich S Schwarz2, Marek Cyrklaff1 and Friedrich Frischknecht *,1 1Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany 2Institute for Theoretical Physics and Bioquant, Heidelberg University, Heidelberg, Germany ‡These authors contributed equally to this work *Corresponding author. Tel: +49 6221 566537; Fax: +49 6221 564643; E-mail: [email protected] The EMBO Journal (2019)38:e100984https://doi.org/10.15252/embj.2018100984 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Microtubules are cytoskeletal filaments essential for many cellular processes, including establishment and maintenance of polarity, intracellular transport, division and migration. In most metazoan cells, the number and length of microtubules are highly variable, while they can be precisely defined in some protozoan organisms. However, in either case the significance of these two key parameters for cells is not known. Here, we quantitatively studied the impact of modulating microtubule number and length in Plasmodium, the protozoan parasite causing malaria. Using a gene deletion and replacement strategy targeting one out of two α-tubulin genes, we show that chromosome segregation proceeds in the oocysts even in the absence of microtubules. However, fewer and shorter microtubules severely impaired the formation, motility and infectivity of Plasmodium sporozoites, the forms transmitted by the mosquito, which usually contain 16 microtubules. We found that α-tubulin expression levels directly determined the number of microtubules, suggesting a high nucleation barrier as supported by a mathematical model. Infectious sporozoites were only formed in parasite lines featuring at least 10 microtubules, while parasites with 9 or fewer microtubules failed to transmit. Synopsis The number and length of microtubules are highly variable within metazoan cells. This study reveals that tuning the number and the length of cytoplasmic microtubules affects the formation, infectivity, and length of sporozoites of the protozoan parasite Plasmodium. Depletion of microtubules in Plasmodium does not inhibit chromosome segregation. Reducing the number and length of microtubules reduces cell size and decreases infectivity of sporozoites. Mathematical modeling predicts that tubulin expression levels dictate microtubule number rather than length, because nucleation rate is higher than elongation rate. Introduction Microtubules are cytoskeletal filaments formed as hollow cylinders from dimers of α-tubulin and β-tubulin (Olmsted & Borisy, 1973; Fojo, 2008). Microtubules can nucleate spontaneously or from a template such as the γ-tubulin ring complex (Roostalu & Surrey, 2017; Wu & Akhmanova, 2017) and can undergo phases of rapid growth and shrinkage (Desai & Mitchison, 1997; Goodson & Jonasson, 2018). Eukaryotic cells can arrange microtubules into different assemblies, including those forming axonemes of flagella and cilia, spindles for genome segregation during cell division, or cytoplasmic asters originating from the microtubule organizing centre and mediating intracellular transport and force distribution (Fojo, 2008). Microtubules provide an attractive target for drugs that interfere with these dynamic assemblies and which are used to treat cancer and infections by pathogenic worms (Jordan & Wilson, 2004; Fennell et al, 2008). In some organisms, different isoforms of tubulin are expressed in different cells or tissues, suggesting that the different physical and biological properties of microtubules can be encoded in the subtle variations of the protein sequence of these isoforms (Panda et al, 1994; Hutchens et al, 1997; Ludueña & Banerjee, 2008; Sirajuddin et al, 2014). Cells can contain from just a few to many hundreds of microtubules (Aikawa, 1971; Osborn & Weber, 1976). While the number of microtubules in axonemes is fixed and their length is stable (Linck et al, 2014), these two key parameters are rarely investigated in cytoplasmic or spindle microtubules. Work in neurons showed that the number of cytoplasmic microtubule is different in neurite-forming axons or dendrites (Yu & Baas, 1994), with the axon containing about 10-fold more microtubules. However, individual microtubule length varied over three orders of magnitude. Owing to those large variations, the precise number and length of cytoplasmic microtubules are rarely taken into account or even investigated in most mammalian or model cells under study. Also, the high rate of growth and shrinkage of microtubules (Mitchison & Kirschner, 1984; Goodson & Jonasson, 2018) make the determination of these parameters difficult if not meaningless as they constantly change. Work on spindle microtubules shows that their number depends on the numbers of kinetochores (Nannas et al, 2014). Both, number and length of spindle microtubules, can be fixed and are likely important for spindle function, as shown in fission yeast (Ward et al, 2014). Similar to metazoans, protozoans can also contain unique assemblies of microtubules, such as those making up the suction disc of the intestinal parasite Giardia lamblia (Nosala et al, 2018) or the axopodia of the free-living heliozoan Echinosphaerium nucleofilum (Tilney & Porter, 1965). Key protozoan organisms that are intensely investigated for their diverse biology and medical relevance include the parasites Trypanosoma brucei causing sleeping sickness and various species of Plasmodium, the causative agents of malaria. Both contain arrays of so-called subpellicular (cytoplasmic) microtubules located in close contact to the surface (pellicle) of the parasites (Garnham et al, 1960, 1961, 1962, 1963; Vanderberg et al, 1967; Sinden & Garnham, 1973; Gull, 1999; Kappes & Rohrbach, 2007; Lacomble et al, 2009), although these parasites are found on different branches on the eukaryotic tree of life. Here, we studied the impact of the subpellicular microtubules on the formation and function of the form of Plasmodium transmitted by mosquitoes using the rodent model species Plasmodium berghei. These so-called Plasmodium sporozoites develop in oocysts at the midgut wall of mosquitoes and enter the salivary glands of the insect from where they are transmitted to a vertebrate host (Ménard et al, 2013; Frischknecht & Matuschewski, 2017; Fig 1A). Sporozoites are highly flexible and motile cells that need to pass several tissue barriers including the salivary glands, the skin and blood vessel endothelium before they infect liver cells to differentiate into thousands of merozoites, which then infect red blood cells (Frischknecht & Matuschewski, 2017; Vaughan & Kappe, 2017). Sporozoites are the target of the first approved malaria vaccine, which contains part of the major sporozoite surface protein as an antigen (Clemens & Moorthy, 2016; Olotu et al, 2016). They are also used in ongoing clinical trials as live vaccines in combination with blood-stage killing drugs or as attenuated parasites (Matuschewski, 2017; Singer & Frischknecht, 2017). Sporozoites of P. berghei contain a fixed number of 16 subpellicular microtubules that extend from an apical polar ring at the front of the sporozoite to the nucleus at the centre of the cell (Vanderberg et al, 1967; Kudryashev et al, 2010, 2012; Fig 1B). These microtubules are arranged in a typical 1 + 15 pattern (Fig 1B) and have been suggested to be important for vesicular trafficking, morphogenesis, cellular mechanics, polarity and motility of sporozoites (Vanderberg et al, 1967; Cyrklaff et al, 2007; Schrevel et al, 2007). The subpellicular microtubules are considered extremely stable and cannot be depolymerized by classic microtubule depolymerizing agents, while addition of taxol derivatives during replication of parasites showed the importance of microtubules in the Plasmodium blood stage (Pouvelle et al, 1994; Schrével et al, 1994; Sinou et al, 1996; Kappes & Rohrbach, 2007). Figure 1. α1-tubulin is essential for sporozoite formation Cartoon of a simplified Plasmodium life cycle indicating in red the stages important for our work. Note that transfection is performed in blood-stage parasites. Schematic cartoon of a sporozoite with an indicated apical cross section. Subpellicular (cytoplasmic) microtubules (green) can be found below the three membrane layers (plasma membrane and inner membrane complex) of the pellicle (bottom). Isolated sporozoites from the hemolymph (HL) and salivary glands (SG) of infected mosquitoes stained with Hoechst and SiR-tubulin. Note the similar staining of WT and α1-tubulin(-) complemented lines. Scale bar: 5 μm. SiR-tubulin staining (green, I and II) and TEM (III) show that microtubules shrink after wild-type sporozoite formation. During budding, microtubules can be up to 15 μm long (I), while after formation they measure on average only 6 μm (graph). Inset in II shows sporozoite cross section at the sporoblast membrane, note the SiR-tubulin fluorescence (green, arrowheads) next to the nuclear Hoechst stain (blue). (III) TEM longitudinal section of budding sporozoites with the length of a subpellicular microtubule indicated by a white line. Green arrowheads point to microtubules close to the sporoblast. Rh: rhoptry; N: nucleus; ER: endoplasmic reticulum; Mi: mitochondrion; Sb: sporoblast. **** indicates P < 0.0001; Kruskal–Wallis test. Scale bars: I and II: 5 μm, magnification box in II: 1 μm, III: 1 μm, magnification boxes in III: 0.1 μm. Comparison of sporozoite development in oocysts of WT, α1-tubulin(-) and complemented lines. Hoechst (blue) and SiR-tubulin (green) staining reveal that the complemented line generates sporozoites indistinguishable from WT controls, while SiR-tubulin staining is absent from α1-tubulin(-) oocysts. mCherry stains the cytoplasm of the parasite. Scale bars: 5 μm. Transmission electron micrographs of developing sporozoites in WT and α1-tubulin(-) parasite lines. Note the slender sporozoites in the WT oocysts, which are absent in the mutant. Sb: sporoblast; scale bars: 5 μm. See also Appendix Figs S2 and S3. TEM cross sections of WT and α1-tubulin(-) parasites showing the aberrant structure of the latter. Subpellicular and centriolar plaque (spindle pole body)-associated (hemi-spindle) microtubules are readily recognizable in the WT sporozoites (green arrowheads in insert) but absent in the mutant, while the inner membrane complex is formed below the plasma membrane of both parasites (white arrowheads). Note the different scale chosen to highlight the malformed shapes in the mutant. N: nucleus; scale bars: 0.2 μm, magnification boxes: 0.1 μm. Source data are available online for this figure. Source Data for Figure 1D [embj2018100984-sup-0005-SDataFig1D.xlsx] Download figure Download PowerPoint Transgenic parasites can be generated in the blood stages of Plasmodium and then transmitted to mosquitoes if the genetic modification is not essential for parasite growth and development in the rodent. In order to assess microtubule function in sporozoites, we now used a gene deletion and replacement approach for α1-tubulin to assess the function of microtubules for sporozoite formation and the impact of modulating their numbers and length on sporozoite biology. Strikingly, these perturbations revealed that Plasmodium genomes can be separated without microtubules and that microtubule length and number are essential for efficient transmission from the mosquito to the mammalian host. Our work demonstrates the essential role of microtubule regulation in a small protozoan of high medical relevance and might pave the way to similar insights in other model organisms. Results Deletion of α1-tubulin affects Plasmodium sporozoite formation during budding All sequenced Plasmodium genomes contain one gene encoding β-tubulin and two genes encoding α-tubulins (Fig EV1A and B). The two α-tubulins likely arose by gene duplication from an ancestral α-tubulin. Such duplications can allow for a change in either the level or the timing of gene expression and/or to generate different functionalities arising from differences within the two coding sequences. Both mechanisms were shown to be at work for α-tubulins in different organisms (Ludueña & Banerjee, 2008). Expression analysis suggested that the α2-tubulin gene is expressed between one and two orders of magnitude higher than the α1-tubulin gene in blood stages (Otto et al, 2014). Indeed, the α2-tubulin gene appears essential and cannot be deleted (Kooij et al, 2005), while we could readily delete the α1-tubulin gene in a wild-type (WT) background and in a parasite expressing GFP and mCherry as cytoplasmic markers at different life cycle stages (Fig EV1C and D). α1-tubulin gene deletion was attempted before without success (Kooij et al, 2005); however, the improved transfection methods since then might have allowed us to generate a transgenic line without problems. Both α1-tubulin(-) parasite lines showed no defect in blood-stage growth or initial mosquito infection (Fig EV1E and F). However, no sporozoites were found in the midgut or salivary glands of the mosquito (Tables 1 and 2); thus, a main defect seems to occur at the oocyst stage. This defect could be completely rescued by complementing the gene in the α1-tubulin(-) parasite line (Fig EV2). Click here to expand this figure. Figure EV1. Generation of α1-tubulin(-) parasite lines Plasmodium genomes, like those of yeast, contain two α1-tubulins and one β-tubulin, while other protozoan such as Toxoplasma gondii or metazoans can contain many more. Data collected from (Ludueña & Banerjee, 2008) and PlasmoDB (version 36; Aurrecoechea et al, 2009). Genome localization and gene structure of P. berghei and P. falciparum tubulin genes. Note that α1-tubulin has a longer C-terminus than α2-tubulin. Strategy to delete α1-tubulin from the genome of P. berghei, strain ANKA (wild type) and from the double fluorescent P. berghei strain ANKA line "RG", which expresses two fluorescent proteins. Primers used for analysis in (D) and resultant amplicons (with length) are indicated. Pyrimethamine is used for positive selection, and 5-fluorocytosine is used for negative selection. Note the Pbdhfr 3′UTRs used for negative selection (green). WL: whole locus. PCR analyses showing the successful deletion of α1-tubulin. Numbers below lanes show expected amplicon sizes as indicated in (C). WL: whole locus. Blood-stage growth rate after injection of 100 WT control and α1-tubulin(-) parasites in 4 and 8 C57BL/6 mice, respectively. Note that a similar growth rate was seen in the PlasmoGEM screen for α1-tubulin(-) parasites (Bushell et al, 2017). One-way ANOVA test was used for statistical analysis. Numbers of oocysts per midgut from infected Anopheles stephensi mosquitoes from WT-like control (see Fig EV2) and α1-tubulin(-) parasite lines. n indicates numbers of investigated midguts. Source data are available online for this figure. Download figure Download PowerPoint Table 1. Early sporozoite formation of the generated parasite lines Table 2. Infective capacity of sporozoites from the generated parasite lines Click here to expand this figure. Figure EV2. Complementation of α1-tubulin(-) parasites with α1-tubulin Cartoon showing the complementation strategy. Primers used for PCR analysis in (B) and amplicon sizes are indicated. PCR analysis of the complemented parasite line. Numbers indicate expected amplicon sizes. Comparative analysis of the complemented and α1-tubulin(-) parasite lines with two wild-type parasite lines (P. berghei strain ANKA and a P. berghei strain ANKA derived line expressing two fluorescent proteins and named RG). Sporozoite numbers from midgut and salivary glands of the different lines and the infectivity of salivary gland sporozoites for NIMR mice were analysed. Note that the 16 microtubules of the WT line are taken from the literature (lit.). Red letters indicate difference from the WT. Download figure Download PowerPoint To investigate whether microtubules were affected by the deletion of α1-tubulin, we established the use of SiR-tubulin, a derivative of taxol that only fluoresces when associated with microtubules (Lukinavičius et al, 2014) for sporozoite (Fig 1C) and oocyst labelling (Fig 1D and E). This showed that microtubules could readily be detected in oocysts and isolated sporozoites with spinning disc confocal microscopy and that their length could be determined with great precision (Fig 1D and Appendix Fig S1). This showed that during late budding in the oocyst microtubules were longest and that they subsequently shrank as the sporozoites matured in the oocysts to again slightly grow as the parasites reside in the salivary gland (Fig 1D). This suggests that, contrary to previous suggestions (Cyrklaff et al, 2007), cytoplasmic microtubules in Plasmodium undergo some level of shrinkage and growth, although much more slowly than in mammalian cells, on the time scale of hours to days rather than seconds. Labelling of nuclei with the DNA-dye Hoechst confirmed the close association of microtubules and nuclei (Fig 1C and D) as expected during the formation of wild-type sporozoites (Vanderberg et al, 1967; Schrevel et al, 2007; Appendix Fig S2). However, in the oocysts of α1-tubulin(-) parasites we could not detect any microtubules (Fig 1E), while they were readily visible in the complemented line (Fig 1C–E). Electron microscopy analysis further showed that no sporozoites were found in oocysts from α1-tubulin(-) parasites (Fig 1F and Appendix Figs S2 and S3). Detailed views of the micrographs revealed that, as in wild-type parasites, the plasma membrane was still subtended by the inner membrane complex (IMC), an organelle defining the alveolates (Harding & Meissner, 2014). However, no normally shaped parasites could be detected in α1-tubulin(-) oocysts. Instead, the parasites presented as undulating shapes with many vesicular structures of unclear provenance. In some sections, we could detect a nucleus enclosed in those deformed "parasites" (Fig 1G). Close inspection of the cytoplasm next to the IMC revealed that in contrast to wild-type, α1-tubulin(-) parasites lacked microtubules (Fig 1G). Sporozoites form by budding from the plasma membrane surrounding the sporoblast, which contains the cytoplasm, nuclei and essential organelles (Sinden & Garnham, 1973; Schrével et al, 1977; Sinden & Strong, 1978; Thathy et al, 2002; Schrevel et al, 2007; Appendix Fig S2). The undulating shapes of the α1-tubulin(-) parasites suggested that initial budding might still proceed. To investigate this in more detail, we studied different steps of sporozoite formation in wild-type and α1-tubulin(-) parasites by transmission electron microscopy (TEM) and by tomographic reconstructions from serial TEM sections (Fig 2A and B). TEM revealed that sporozoite budding appears to initiate normally in α1-tubulin(-) parasites (Fig 2A and Appendix Figs S2 and S3). The nuclei appeared aligned underneath the plasma membrane as in wild type with their spindle pole body (centriolar plaque) oriented towards the growing apical end of the sporozoite and possibly still connected to the growing tip by the rootlet fibre, a dynamic tether that links the centriolar plaque to the apical end of the emerging parasites (Sinden & Strong, 1978; Schrevel et al, 2007; Francia et al, 2012). However, the growing parasites in the mutants showed irregular shapes that contrasted the more regular and elongated WT sporozoite form (Fig 2A and B). While the inner membrane complex subtended the plasma membrane of both budding WT and mutant sporozoites (Fig 2A), the growing α1-tubulin(-) parasites were malformed and contained numerous vesicular structures (Fig 2A and B). The nuclei lacked intra-nuclear microtubules and were not elongated like those within the growing WT sporozoites, where intra-nuclear microtubules extended along the entire longitudinal axis of the ovoid nuclei (Fig 2A and B; Movie EV1). Later during development, α1-tubulin(-) parasites showed large extensions from the sporoblast membrane with multiple bends and diverse diameters and rarely contained nuclei (Fig 2C and D, Movie EV2). These data suggest that microtubules play an essential structural role during sporozoite formation in the oocyst and that specifically α1-tubulin is required for microtubule formation at this stage. Figure 2. Sporozoite budding is impaired in α1-tubulin(-) parasites Electron micrographs showing different stages of development in WT and α1-tubulin(-) parasites. Key structural elements are highlighted in the following colours: blue: plasma membrane; yellow: inner membrane complex; green: microtubules; light blue: micronemes; lilac: rhoptries; red: rootlet fibre; and cyan: nucleus. Note the undulating shapes of the elongated mutants and that micronemes are hard to distinguish from other small vesicles in the α1-tubulin(-) parasites. The rootlet fibre in panel II starts at the apical tip and in panel V is associated with the centriolar plaque (dark structure in nuclear envelope). Panel III corresponds to Fig 1D. Scale bars: 200 nm. 3D reconstruction from serial sectioning reveals that growing α1-tubulin(-) "sporozoites" lack apical polarity and fail to pull in nuclei. Microtubuli: green; rootlet fibre (connecting apical pole to centriolar plaque): red; rhoptries: purple; endomembraneous vesicles (Golgi, premicronemes): brown; nucleus: cyan; sporozoite plasma membrane and inner membrane complex: blue; dotted lines show location of sporoblast membrane. 17 and 10 sections of 100 and 300 nm thickness were processed for the tomograms in WT and α1-tubulin(-) parasites, respectively. Scale bar: 1 μm. See also Movie EV1. Tomography of serial sections from budding α1-tubulin(-) parasites at day 13 (same as in panel B) and day 17 highlights the increase in surface area (blue) with time. The oocyst wall at day 17 is highlighted by cyan lines (top), and the sporoblast membrane is highlighted by red lines (bottom). The sporoblast membrane at day 13 is indicated by a dashed line (red). 10 and 11 sections of 300 nm thickness were processed for the tomograms at days 13 and 17, respectively. A single continuous 'parasite' was highlighted at day 17. Scale bar: 1 μm. α1-tubulin(-) parasites show multiple attachments (arrowheads) with the sporoblast membrane at day 17 post-infection. Note the highly branched nature of the budding parasite (same parasite as in panel C) shown in two views in I and II. Right: slices through the tomogram in grey scale with the outlines used for the 3D model indicated. Two connections with the sporoblast are indicated by 1 and 2. Scale bars: 1 μm. See also Movie EV2. Download figure Download PowerPoint α1-tubulin deletion does not affect genome replication and nuclear division As the deletion of genes could lead to upregulation of other genes necessary in the affected process, we next investigated mRNA levels of α1-tubulin and α2-tubulin by qRT–PCR. This showed that little α2-tubulin mRNA was present during sporozoite formation (Fig EV3A) and that levels of α2-tubulin mRNA were unaffected by α1-tubulin deletion (Fig EV3B). Similarly, we also noted that the expression of the major surface antigen, circumsporozoite protein (Cohen et al, 2010; Frischknecht & Matuschewski, 2017), was also not affected by the deletion of α1-tubulin (Fig EV3C); albeit, a tendency for lower expression levels could be detected at days 12 and 14 post-mosquito infection. Click here to expand this figure. Figure EV3. Absence of microtubules delays CSP expression A, B. mRNA levels as determined by qRT–PCR analysis of infected midguts at different times post-mosquito infection. Note that α2-tubulin is much less expressed in oocysts than α1-tubulin (A) and that α2-tubulin expression is unchanged in α1-tubulin(-) oocysts (B). Error bars indicate the standard deviation from the mean of technical duplicates. C. CSP expression of α1-tubulin(-) parasites relative to WT on different days post-mosquito blood meal. Relative expression was calculated via the ∆∆ct method with error bars indicating the standard deviation of the mean calculated from technical duplicates. D. Live images of oocysts expressing GFP from the ef1α promoter and mCherry from the csp promoter. Microtubules were labelled with SiR-tubulin, and DNA was labelled with Hoechst in the merged image. Note that CSP expression is not seen in all oocysts (−) and that oocysts with stronger (+) and very strong (++) CSP expression progressed further in development as evidenced by strong SiR-tubulin staining and individualized nuclei. Scale bar: 10 μm. E. Quantification of oocyst maturation as judged from expression of CSP on days 10 and 12 post-blood meal. The number of total investigated oocysts is indicated above each column, and −, +, ++ indicate no, weak and strong CSP expression as indicated in panel D. Source data are available online for this figure. Download figure Download PowerPoint We next analysed the α1-tubulin(-) parasite line that expressed mCherry from the csp promoter. This line also expresses GFP from a constitutively active promoter and thus allows detecting developing oocysts by GFP, the onset of csp promoter activity during parasite formation by mCherry, DNA replication by addition of Hoechst and microtubule formation by SiR-tubulin, which fluoresces in the far red (Lukinavičius et al, 2014). We noted that in cells where few nuclei are present and no alignment of the nuclei to the sporoblast membrane had taken plac
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