Atlas of Plasmodium falciparum intraerythrocytic development using expansion microscopy

  1. Benjamin Liffner
  2. Ana Karla Cepeda Diaz
  3. James Blauwkamp
  4. David Anaguano
  5. Sonja Frolich
  6. Vasant Muralidharan
  7. Danny W Wilson
  8. Jeffrey D Dvorin
  9. Sabrina Absalon  Is a corresponding author
  1. Department of Pharmacology and Toxicology, Indiana University School of Medicine, United States
  2. Biological and Biomedical Sciences, Harvard Medical School, United States
  3. Division of Infectious Diseases, Boston Children’s Hospital, United States
  4. Center for Tropical and Emerging Global Diseases, University of Georgia, United States
  5. Department of Cellular Biology, Franklin College of Arts and Sciences, University of Georgia, United States
  6. Research Centre for Infectious Diseases, School of Biological Sciences, University of Adelaide, Australia
  7. Institute for Photonics and Advanced Sensing, University of Adelaide, Australia
  8. Burnet Institute, 85 Commercial Road, Australia
  9. Department of Pediatrics, Harvard Medical School, United States

Abstract

Apicomplexan parasites exhibit tremendous diversity in much of their fundamental cell biology, but study of these organisms using light microscopy is often hindered by their small size. Ultrastructural expansion microscopy (U-ExM) is a microscopy preparation method that physically expands the sample by ~4.5×. Here, we apply U-ExM to the human malaria parasite Plasmodium falciparum during the asexual blood stage of its lifecycle to understand how this parasite is organized in three dimensions. Using a combination of dye-conjugated reagents and immunostaining, we have cataloged 13 different P. falciparum structures or organelles across the intraerythrocytic development of this parasite and made multiple observations about fundamental parasite cell biology. We describe that the outer centriolar plaque and its associated proteins anchor the nucleus to the parasite plasma membrane during mitosis. Furthermore, the rhoptries, Golgi, basal complex, and inner membrane complex, which form around this anchoring site while nuclei are still dividing, are concurrently segregated and maintain an association to the outer centriolar plaque until the start of segmentation. We also show that the mitochondrion and apicoplast undergo sequential fission events while maintaining an association with the outer centriolar plaque during cytokinesis. Collectively, this study represents the most detailed ultrastructural analysis of P. falciparum during its intraerythrocytic development to date and sheds light on multiple poorly understood aspects of its organelle biogenesis and fundamental cell biology.

eLife assessment

This important study provides an unprecedented overview of the subcellular organization of proliferative blood-stage malaria parasites using expansion microscopy. The localization of multiple parasite organelles is comprehensively probed using three-dimensional super-resolution microscopy throughout the entire intraerythrocytic development cycle. This work provides a compelling framework to investigate in future more deeply the unconventional cell biology of malaria-causing parasites.

https://doi.org/10.7554/eLife.88088.3.sa0

Introduction

The human malaria parasite Plasmodium falciparum has a complex lifecycle that involves both human and mosquito hosts. Of its many lifecycle stages, the asexual replication of P. falciparum inside human red blood cells (RBCs) is responsible for the clinical symptoms of malaria. This asexual blood stage starts when a merozoite invades a host RBC and transitions through several morphologies before forming approximately 30 new daughter merozoites (Rudlaff et al., 2020), which egress from their host cell and invade new RBCs (Figure 1a). Host RBCs are approximately 7–8 µm in diameter (Kinnunen et al., 2011) and contain dozens of parasites, each with their own sets of organelles and structures. The small size of P. falciparum and its organelles still poses a challenge to the study of many facets of P. falciparum cell biology, especially when immunostaining is required.

Figure 1 with 2 supplements see all
Ultrastructural expansion microscopy (U-ExM) workflow and summary of parasite structures imaged in this study.

(a) Diagram of asexual blood-stage lifecycle of P. falciparum. (b) U-ExM workflow used in this study. PFA = paraformaldehyde, FA = formaldehyde, AA = acrylamide, PG = propyl gallate. Snowflake indicates steps where gels were cryopreserved. (c) Comparison of brightfield and DAPI staining of unexpanded P. falciparum parasites (inset) with P. falciparum prepared by U-ExM, stained with N-hydroxysuccinimide (NHS) ester (protein density; grayscale) and SYTOX Deep Red (DNA; cyan), and imaged using Airyscan microscopy. Images are maximum-intensity projections, number on image = Z-axis thickness of projection in µm. Scale bars = 2 µm. (d) Summary of all organelles, and their corresponding antibodies, imaged by U-ExM in this study.

Expansion microscopy is a set of sample preparation techniques that isotropically increase the physical size of a microscopy sample (Wassie et al., 2019). While many expansion microscopy methods have been developed, ultrastructure expansion microscopy (U-ExM) (Gambarotto et al., 2019) was the first used in Plasmodium and since has been used in P. falciparum and across multiple apicomplexan parasites (Bertiaux et al., 2021; Dave et al., 2022; Liffner and Absalon, 2021; Oliveira Souza et al., 2022; Rashpa and Brochet, 2022; Severo et al., 2022). U-ExM results in the ~4.5-fold isotropic expansion of the sample and largely preserves its proteome, making it compatible with antibody staining and many fluorescent dyes (Gambarotto et al., 2019). The increase in physical sample size results in a dramatic increase in the ability to identify and distinguish different parasite structures. Thus, some structures that could previously only be investigated using electron microscopy can now be studied with the flexibility, scalability, and inexpensive nature of conventional light microscopy.

Application of U-ExM to Plasmodium, and other Apicomplexa, has already enhanced our understanding of parasite cell biology tremendously, resulting in the identification of new parasite structures and better characterization of the size and shape of others (Bertiaux et al., 2021; Liffner and Absalon, 2021; Qian et al., 2022; Rashpa and Brochet, 2022; Simon et al., 2021; Tomasina et al., 2022; Tosetti et al., 2020). Its significant impact on the field in such a short amount of time indicates U-ExM will be a technique heavily used in both Plasmodium and Apicomplexa more broadly for the foreseeable future. Considering this, we set out to image P. falciparum structures and organelles across the asexual blood stage of the lifecycle to serve as a reference for the expanding number of U-ExM users who study Apicomplexa and uncover previously invisible aspects of the cell biology of P. falciparum.

Results

U-ExM reveals multiple parasite structures without the use of antibodies

Dyes that are not antigen-specific are commonplace in light microscopy. N-hydroxysuccinimide (NHS) esters conjugated to dyes are amino-reactive and can be used for fluorescent labeling of protein density (Nanda and Lorsch, 2014). Similarly, BODIPY TR ceramide (BODIPY TRc) and other dye-conjugated fatty acids are commonly used for labeling lipids (Marks et al., 2008). Coupling U-ExM with these dyes has already revealed parasite structures for which specific antibodies did not exist (Bertiaux et al., 2021; Liffner and Absalon, 2021; Simon et al., 2021). While antibody-based labeling provides high specificity, this labeling lacks complexity and is limited to the specific protein or antigen that is being targeted. The more general stains increase the number of parasite features or organelles we can observe in the same sample without additional antibody markers. Therefore, these general stains allow for low-specificity but high-complexity imaging. An example of this principle is the use of uranyl acetate in electron microscopy to increase contrast by increasing the electron density of phosphate-rich structures in the cell (Rudlaff et al., 2020). While NHS esters and uranyl acetate can bring out similar features in the cell, they do not have the same reactivity and are therefore not equivalent stains. To better profile the subcellular organization of P. falciparum during the asexual blood stages, we set out to determine what parasite structures could, and could not, be visualized by U-ExM when using some of these dyes.

We, and others, have previously shown that combining BODIPY TRc and dye-conjugated NHS ester with U-ExM allows visualization of the parasite plasma membrane (PPM), parasitophorous vacuolar membrane (PVM), nuclear envelope, rhoptries, endoplasmic reticulum (ER), centriolar plaque (CP), basal complex, and apical polar rings (APRs) (Bertiaux et al., 2021; Liffner and Absalon, 2021; Simon et al., 2021). However, many important organelles and parasite structures are either not identifiable using these stains or have yet to be validated, including the mitochondrion, apicoplast, and cytostomes.

To identify and validate the location of as many parasite organelles and structures as possible, we utilized U-ExM coupled with BODIPY TRc, Alexa Fluor 405-conjugated NHS ester (which we will refer to as ‘NHS ester’), the nucleic acid (i.e., DNA) stain SYTOX Deep Red, and antibodies directed against 13 different subcellular targets (microtubules, CP, basal complex, inner membrane complex [IMC], mitochondrion, apicoplast, cytostome, rhoptry bulb, rhoptry neck, micronemes, cytoplasm, ER, and Golgi). In this study, all parasites were fixed in 4% paraformaldehyde (PFA), unless otherwise stated, and anchored overnight at 37°C before gelation, denaturation at 95°C, and expansion. Expanded gels were measured (Figure 1—figure supplement 1), before shrinking in PBS antibody staining, washing, re-expansion, and imaging (Figure 1b). Parasites were harvested at multiple time points during the intraerythrocytic asexual stage and imaged using Airyscan 2 super-resolution microscopy, providing high-resolution three-dimensional imaging data (Figure 1c). A full summary of all target-specific stains used in this study can be found in Figure 1d.

The first protein we imaged was aldolase, a marker of the parasite cytoplasm (Figure 1—figure supplement 2). Aldolase staining was present in all asexual replication stages. During the ring stage, the ‘ameboid’ shape of the parasite is readily visualized, consistent with previous studies of this stage in time-lapse microscopy of live parasites (Grüring et al., 2011; Figure 1—figure supplement 2). Regions within the parasite where both aldolase and NHS ester staining were absent are consistent with the expected area of the food vacuole. Typically, the food vacuole would be filled with hemozoin; however, this crystal likely cannot expand and therefore leaves a large space inside the parasite that does not contain significant protein density (Figure 1—figure supplement 2).

The CP, nuclear MTOC, and microtubules

The first major transition during the blood stage of the lifecycle occurs when the parasites turn from rings into trophozoites. Soon after this transition, the parasite will begin to replicate its DNA and undergo mitosis followed by nuclear fission (Gerald et al., 2011). Mitosis is coordinated by microtubules, which are in turn nucleated by structures called microtubule organizing centers (MTOCs) (Sanchez and Feldman, 2017). P. falciparum has a structure known as the CP that spans the nuclear envelope, with intranuclear and cytoplasmic portions (Simon et al., 2021). In this study, we will refer to the intranuclear portion of the CP as the inner CP and the cytoplasmic portion of the CP as the outer CP. The inner CP acts as the nuclear MTOC that coordinates P. falciparum mitosis, while the function of the outer CP is unknown in asexual blood-stage parasites. The most commonly used MTOC markers are the centrins, which in Plasmodium comprise four proteins that localize to the outer CP and appear after intranuclear microtubules have already formed (Simon et al., 2021). Given that an MTOC is required for microtubule formation, this implies that an MTOC forms before centrin is visible. To investigate these processes in more detail, we visualized the biogenesis and dynamics of the CP and microtubules during the trophozoite and schizont stages by pairing NHS ester, which we have recently shown can stain both the inner and outer CP (Liffner and Absalon, 2021), with an anti-centrin antibody (Clone 20H5, raised against centrin from Chlamydomonas) (Figure 2a). This antibody likely recognizes centrin 3 in P. falciparum (Mahajan et al., 2008), but a recent study suggests that all four P. falciparum centrins share an outer CP localization (Voß et al., 2022).

Figure 2 with 2 supplements see all
Centriolar plaque (CP) biogenesis and dynamics.

3D7 parasites were prepared by ultrastructural expansion microscopy (U-ExM), stained with N-hydroxysuccinimide (NHS) ester (grayscale), BODIPY TRc (white), SYTOX (cyan), and anti-centrin (outer CP; magenta) antibodies and imaged using Airyscan microscopy. (a) Images of whole parasites throughout asexual blood-stage development. (b) Whole parasite panel (left) followed by individual CP or CP pair zooms following our proposed timeline of events in CP biogenesis, dynamics, and disassembly. Yellow line = cytoplasmic extensions, blue line = nuclear envelope, green line = parasite plasma membrane. Images are maximum-intensity projections, number on image = Z-axis thickness of projection in µm. White scale bars = 2 µm, yellow scale bars = 500 nm.

CP biogenesis and disassembly

Neither a recognizable CP nor above-background centrin staining were observed in ring-stage parasites (Figure 2b; Simon et al., 2021). The inner CP first appeared in mononucleated trophozoites but changed morphology as these parasites got closer to their first nuclear division. In the 23 mononucleated trophozoites we imaged, 52% of CPs lacked cytoplasmic extensions (Figure 2—figure supplement 1a and b). These CPs contained only the inner CP and lacked the outer CP as observed by NHS ester (Figure 2a and b). All 12 of the trophozoites with an inner CP but no outer CP also lacked centrin staining (Figure 2—figure supplement 1c). This matches previous reports that centrin is specifically associated with the outer CP (Simon et al., 2021). As expected, this early CP lacking the outer CP was capable of nucleating microtubules (Figure 2—figure supplement 1a). The centrin focus and outer CP became visible in mononucleated trophozoites after nucleation of the intranuclear microtubules but prior to the first CP duplication event of the first mitosis (Figure 2—figure supplement 1a), consistent with previous reports (Simon et al., 2021). The cytoplasmic extensions that form the outer CP began at the nuclear membrane and ended at an NHS ester-dense focus located at the PPM (Figure 2b). This association between the outer CP and the PPM has previously been observed in gametocytes and asexual blood stages (Li et al., 2022; Rashpa et al., 2023), but the temporal nature of this association during the asexual blood stages remained uncharacterized. Our observation of the outer CP using NHS ester, discussed further below, combined with the temporal pattern of outer CP–PPM association, suggests that nuclei are physically anchored to the PPM while parasites are undergoing mitosis.

For as long as parasites continue to undergo mitosis throughout schizogony, the outer CP remains in contact with the PPM (Figure 2a). The outer CP appears as one or two elongated bundles, referenced throughout this paper as ‘branches,’ that stain densely with NHS ester. These branches showed one or two centrin foci that largely matched branch number. That is, of the 139 single-branched outer CPs (outer CPs with a single cytoplasmic extension branch) observed, all had a single centrin focus, and of the 30 double-branched outer CPs observed, 92% had two centrin foci (Figure 2b, Figure 2—figure supplement 1c). The overwhelming coordination between number of cytoplasmic extension branches and number of centrin foci suggests centrin duplication and duplication of the branches of the outer CP happens quickly and simultaneously. The few cases when there is a mismatch between the number of cytoplasmic extensions and centrin foci all occur in double-branched outer CPs and can be attributed to limitations in our ability to resolve two centrin foci (Figure 2—figure supplement 1c and d).

The relative abundance of nuclei with a single CP, defined as a CP not forming a mitotic spindle, versus mitotic CPs, CPs anchored to a mitotic spindle, varied throughout schizogony as did their branch numbers (Figure 2—figure supplement 1). As we previously observed that the inner CP forms first, we hypothesized that CPs start out without an outer CP and then develop a single cytoplasmic extension and centrin focus, which duplicate ahead of CP duplication and then segregate with each CP during miotic spindle formation and karyokinesis (Figure 2b). This means that during the rapid mitotic events of schizogony that take place at the 6–12 nuclei stage, single outer CPs with a single cytoplasmic extension are very rare. We observed them in just 6% of 221 imaged CPs (Figure 2—figure supplement 1b). Single outer CPs with a single cytoplasmic extension were only abundant when the pace of nuclear replication was slow at the early schizont stage (24% of 76 CPs imaged in cells with 2–5 nuclei) and the end of segmentation (44% of 290 CPs imaged in segmenting parasites).

This observed pattern of duplication and segregation also suggests that double-branched outer CPs with two centrin foci have committed to undergoing a new round of mitosis and that the duplication of cytoplasmic extensions represents the first identifiable step in CP duplication. In line with this hypothesis, the most common CP states in schizonts prior to segmentation are single CPs with two cytoplasmic extensions (26% of CPs in cells with 2–5 nuclei, 52% in cells with 6–12 nuclei) that have committed to the next round of mitosis and mitotic CPs with one extension each (42% of CPs in cells with 2–5 nuclei, 36% in cells with 6–12 nuclei) that are finishing a round of mitosis. Interestingly, 8% of CPs observed in the 2–5 nuclei stage and 6% in the 6–12 nuclei stage were mitotic and double-branched, suggesting the duplication of cytoplasmic extensions, and commitment to the next round of mitosis, can happen before karyokinesis is completed (Figure 1—figure supplements 1e and 2).

Outer CP branch number reaches semi-synchrony at the beginning of segmentation as defined by the first appearance of a basal complex by NHS ester. At this point, rather than seeing a variety of CP states and branch numbers, virtually all CPs in the same cell share the same mitotic state and branch number. Most CPs are mitotic with a single cytoplasmic extension during early segmentation and then appear as single CP with a single extension during mid-segmentation (Figure 2—figure supplement 1b). By the time segmentation is completed, the CP is no longer visible by NHS ester, suggesting that it may disassemble after all mitotic events are finished (Figure 2b, Figure 2—figure supplement 1b). To ensure imaged parasites were fully segmented, we arrested parasite development by adding the reversible protein kinase G inhibitor compound 1 (C1) (Collins et al., 2013; Gurnett et al., 2002; Taylor et al., 2010). This inhibitor arrests parasite maturation after the completion of segmentation but before egress. When C1 is washed out, parasites egress and invade normally, ensuring that observations made in C1-arrested parasites are physiologically relevant and not a developmental artifact due to arrest. Of 159 nuclei imaged in 6 C1-arrested schizonts, none showed the presence of a CP.

The apical polar rings, Golgi, and rhoptries are all segregated with the CP

Given the cytoplasmic coordination of mitotic events and the physical tethering of the nucleus to the PPM throughout schizogony, we investigated whether we could observe any coordination extending to the apical organelles and other structures known to be present near the CP at these stages. We observed close association between the outer CP and the rhoptries, Golgi, basal complex, and an apical density reminiscent of the APR.

The number of basal complex structures closely matched the number of outer CP extensions throughout schizogony (156 observed cytoplasmic extensions, 153 observed basal complexes). The basal complex, discussed in detail below, appears as a ring-like structure from early schizogony to the completion of segmentation. No basal complex structures were observed in CPs without cytoplasmic extensions. Of 82 single-branched outer CPs imaged, 81 (99%) showed a single basal complex structure. Of 37 double-branched outer CPs imaged, 33 (89%) showed two basal complex structures. As we observed when imaging centrin foci, the few cases when there is a mismatch between the number of cytoplasmic extensions and basal complex structures mostly occur in double-branched outer CPs. However, in this case, the mismatch cannot be attributed to resolution and most likely reflects a transition state in basal complex replication. CINCH stains the basal complex in a punctate pattern until early segmentation, allowing us to visualize basal complex division events as gaps in the punctate staining (Figure 4a and b). The occasional presence of two cytoplasmic extensions in the absence of these gaps suggests that basal complex division, when visualized by breaks in CINCH staining, is less simultaneous with outer CP branch duplication than centrin focus duplication.

Rhoptry biogenesis, discussed further below, was also closely tied to the number and position of the outer CP. This association of one rhoptry per branch, however, is broken at the start of segmentation, when 205 of 211 imaged CPs (97%) had a rhoptry pair per outer CP branch. Further suggestive of CP–rhoptry interaction is the fact that the rhoptries were positioned immediately next to the outer CP for as long as these were visible by NHS ester. While we had no APR protein marker, the cytoplasmic extensions always ended in an NHS ester-dense focus at the plasma membrane. At the beginning of segmentation, this focus obtained a morphology suggestive of the APR based on its ring shape, position, and size. As described above, a small percentage of parasites commit to the next round of mitosis before they finish segregating their genetic material. In these parasites, CP-associated organelles continued to match the number of cytoplasmic extensions. This gave rise to nuclei that were anchored to four basal complexes, four single rhoptries, and four proto-APR densities in close proximity (Figure 4—figure supplement 1). This matches previous observations by three-dimensional electron microscopy that a single nucleus can have four sets of apical buds (Rudlaff et al., 2020).

We also characterized the distribution of the Golgi and its location near the CP and apical organelles, this spatial correlation had been previously described by electron microscopy but has not been thoroughly explored (Bannister et al., 2000). The Golgi was visualized using an antibody to ER lumen protein retaining receptor 2 (ERD2), a cis-Golgi marker expressed throughout intraerythrocytic development (Elmendorf and Haldar, 1993; Figure 2—figure supplement 2). Small regions of Golgi were visible at all development stages. In ring-stage parasites and mononucleated trophozoites, one or two Golgi foci were observed near the nucleus but had no clear proximity to the CP. In all five imaged parasites with less than two nuclei and an outer CP, no Golgi was observed near the inner CP. In parasites that had undergone the first round of mitosis and had an outer CP, the Golgi was proximal to the extensions of the outer CP (Figure 2—figure supplements 1b and 2) and closely matched their number and presence. Specifically, 21 of 22 imaged parasites with visible outer CPs had Golgi staining near each branch of their outer CP (Figure 2—figure supplement 1b). While increased Golgi–CP proximity coincides with the appearance of the cytoplasmic extensions, the Golgi is able to remain toward the apical end of the parasite after these tethers and the CP are no longer visible by NHS ester. However, at this point the Golgi loses its close proximity to the apical organelles and is situated closer to the nucleus. In C1-arrested schizonts, each merozoite has a single Golgi that remains at the apical end of the parasite, typically between the rhoptry bulb and the nucleus (Figure 2—figure supplement 2).

In contrast, we did not observe a CP association in the distribution of the ER (Figure 4—figure supplement 2) within the parasite. The ER was visualized using an antibody to binding immunoglobulin protein (BIP), a constitutively expressed ER lumen protein (Kumar et al., 1991). As expected, ER was detected at all stages of intraerythrocytic development (Figure 4—figure supplement 2). In ring-stage and mononucleated trophozoite-stage parasites, the ER could be seen wrapping around the nucleus and forming recognizable cisternae. In multinucleated parasites, the ER was too dense to observe cisternae, but large regions of the cell were occupied by the ER. Following segmentation in C1-arrested schizonts, the ER was only observed contiguous with the nuclear envelope.

Combining the observations that the CP is physically tethered to the PPM through the outer CP and that this anchoring is closely associated with organelles that will define the apical end of the parasite (Golgi, rhoptries, basal complex, and APRs), we suggest that this tethering by the outer CP establishes apical–basal polarity in the parasite early in schizogony. Considering that rhoptries are formed from Golgi-derived cargo (Ben Chaabene et al., 2021; Counihan et al., 2013), it is unsurprising to find the Golgi forms part of this apical cluster of organelles throughout schizogony. The confined space between nuclear envelope and PPM that these organelles are packed into, for example, may allow each nucleus to provide rhoptry cargo locally to their own rhoptries rather than to all rhoptries in the cell. The same principle could apply to other apical Golgi-derived organelles. However, it remains unclear what role, if any, the outer CP plays in this association, whether any organelles besides the nucleus are physically tethered by these extensions, and how these clusters of organelles remain together during the rapid mitotic events constantly separating sister CPs.

Characterization of intranuclear microtubules

P. falciparum asexual blood stages are known to have two classes of microtubules; intranuclear microtubules, which partake in mitosis (Liffner and Absalon, 2021; Simon et al., 2021), and subpellicular microtubules (SPMTs), which are cytosolic and extend in a single spine from the apical end of merozoites (Liffner and Absalon, 2021; Simon et al., 2021). Investigating microtubules with an anti-α-tubulin antibody, we failed to detect microtubules in ring-stage parasites, consistent with previous observations (Figure 3—figure supplement 1; Simon et al., 2021). Intranuclear microtubules were first visible in mononucleated trophozoite-stage parasites and were present until early segmentation stages, with no intranuclear microtubules visible by the end of segmentation (Figure 3a). Intranuclear microtubules arrange into three distinct spindle structures: hemispindles, mitotic spindles, and interpolar spindles. Hemispindles are microtubule structures coming from a single CP that retract prior to CP duplication. Mitotic spindles appear following CP duplication and separate sister chromatids during mitosis. When the two CPs migrate away from each other, they remain connected by an elongated microtubule structure called the interpolar spindle (or elongated spindle), which retracts prior to nuclear fission (Liffner and Absalon, 2022; Machado et al., 2022; Simon et al., 2021). It has recently been shown that the interpolar spindle is short-lived relative to the hemispindle and mitotic spindle (Machado et al., 2022; Simon et al., 2021). In this study, we observed 24 interpolar spindles, which allowed us to perform the first detailed characterization of this spindle type (Figure 3b).

Figure 3 with 1 supplement see all
Characterization of intranuclear and subpellicular microtubules (SPMTs).

3D7 parasites were prepared by ultrastructural expansion microscopy (U-ExM), stained with N-hydroxysuccinimide (NHS) ester (grayscale), BODIPY TRc (white), SYTOX (cyan), and anti-tubulin (microtubules; magenta) antibodies, and imaged using Airyscan microscopy. (a) Images of whole parasites throughout asexual blood-stage development. (b) Nuclei in the process of dividing, with their CPs connected by an interpolar spindle. (c) The number and type of microtubule branches in interpolar spindles and (d) length of interpolar microtubules. (e) SPMTs stained with an anti-poly-glutamylation (PolyE; yellow) antibody. (f) Quantification of the number of SPMTs per merozoite from C1-treated schizonts. (g) SPMT biogenesis throughout segmentation. (h) Model for SPMT biogenesis. PPM = parasite plasma membrane, APRs = apical polar rings, BC = basal complex, CP = centriolar plaque. Images are maximum-intensity projections, number on image = Z-axis thickness of projection in µm. Scale bars = 2 µm.

Interpolar spindles have microtubule branches that connect the two distant CPs (interpolar microtubules), and microtubule branches that do not connect the CPs (non-interpolar microtubules). Each interpolar spindle contained an average of 12.5 (±2.6 SD) total microtubules, of which 1.3 (±0.6 SD) were interpolar microtubules and 11.2 (±2.8 SD) were non-interpolar microtubules (Figure 3c). The average number of non-interpolar branches per inner CP was 5.6, which is similar to the previously reported average number of branches in a hemispindle of 5–6 (Liffner and Absalon, 2021; Simon et al., 2021). This suggests that only the interpolar microtubules retract during the interpolar spindle to hemispindle transition. We measured interpolar microtubules in 3D, adjusting for expansion factor by dividing the measured distance by 4.25, the median expansion factor observed in this study (Figure 1—figure supplement 1; ‘Materials and methods’). All of the following measurements in this study are reported in this expansion-corrected format. Interpolar microtubules ranged from ~1 to 5 µm, with a mean length of 2.9 µm (±1.0 µm SD) or 12.47 µm before expansion factor correction (Figure 3d). In all cases, the CPs connected by interpolar spindles were anchored to the plasma membrane by their cytoplasmic extensions. The large variability in interpolar microtubule size and the continued tethering of the outer CPs to the PPM suggest that interpolar microtubules push PPM-anchored CPs to opposite sides of the cell without causing detachment from the PPM. It is unclear how parasites achieve this sliding effect or how CP-associated organelles are able to retain this association while CPs are moved large distances.

SPMT length and biogenesis

SPMTs are nucleated in the cytoplasm and have long been observed in merozoites (Aikawa, 1967). SPMTs have been shown to be stabilized by polyglutamylation (Bertiaux et al., 2021) and can be identified specifically using a combination of anti-tubulin and anti-PolyE antibodies (Figure 3e). Using this approach, we characterized 86 SPMTs in 50 merozoites from C1-arrested schizonts. These nascent merozoites had between 1 and 3 SPMTs, with an average of 1.7 (±0.6 SD) (Figure 3f). Of 50 imaged merozoites, 48 had at least one SPMT that extended >50% of cell length from the APR to the basal complex. This longest microtubule in a merozoite had an average length of 1.01 µm (±0.24 µm SD). In merozoites with more than one SPMT, the second and third microtubules were shorter than the first, having an average length of 0.8 µm (±0.21 µm SD). Given the large variation in SPMT size and observation that, in segmenting schizonts, the basal end of the SPMTs was in contact with the basal complex throughout segmentation, we hypothesize that most SPMTs measured in our C1-treated schizonts had partially depolymerized. P. falciparum microtubules are known to rapidly depolymerize during fixation (Liffner and Absalon, 2022; Simon et al., 2021). It is unclear, however, why this depolymerization was observed most often in C1-arrested parasites. Thus, we cannot determine whether these shorter microtubules are a by-product of drug-induced arrest or a biologically relevant native state that occurs at the end of segmentation.

Little is known about SPMT biogenesis during the asexual blood stage of P. falciparum, but it is currently hypothesized that they are nucleated by the APRs (Hanssen et al., 2013; Morrissette and Sibley, 2002), as is the case in Toxoplasma (Morrissette and Sibley, 2002; Tran et al., 2010). Curiously, TgCentrin 2 localizes to the APR of Toxoplasma tachyzoites (Hu, 2008), but no Centrin 2 has been observed to localize to the APRs of P. falciparum. Furthermore, it was recently shown that the SPMTs of P. falciparum gametocytes, which lack an APR, are formed at the outer CP, in the space between the nuclear envelope and PPM (Li et al., 2022). Leveraging our ability to specifically detect SPMTs using PolyE, we investigated the possibility that merozoite SPMTs are also formed at the outer CP and subsequently transferred onto the APR during segmentation. In schizonts where nuclei are approaching or have completed their final mitosis (~15n), we observed small cytoplasmic microtubules that stained strongly with PolyE appear in the area between the outer CP and PPM (Figure 3g and h). However, we did not achieve a resolution that allowed us to distinguish individual APRs or to confidently pinpoint whether the microtubules were nucleated at the APRs or the cytoplasmic extensions. Likely, higher resolution imaging techniques are needed to resolve the site of SPMT nucleation in merozoites.

Segmentation machinery (IMC and basal complex)

Following replication of their genetic material during the trophozoite and early schizont stages, parasites partition their nuclei and organelles into ~30 daughter merozoites from the common cytoplasm of a schizont (Francia and Striepen, 2014). This form of cytokinesis, called segmentation, takes place in the final hours of schizogony and culminates with the physical separation of each daughter cell and their egress from the host RBC. The IMC is a double lipid bilayer formed from flattened vesicles that scaffolds the process of segmentation as well as anchors many proteins important for parasite shape and motility (Harding and Meissner, 2014).

The IMC cannot be distinguished from the plasma membrane by U-ExM

The IMC forms de novo during segmentation starting at the apical end of the parasite, where the outer CP is anchored to the plasma membrane (Figure 4—figure supplement 3a; Harding and Meissner, 2014). This can be observed using the IMC-anchored protein glideosome-associated protein 45 (GAP45), which bridges the IMC and plasma membrane, as well as using BODIPY TRc, which shows increased membrane staining in the area overlapping GAP45 (Jones et al., 2006; Figure 4—figure supplement 3a). As segmentation progresses, the IMC expands around the nucleus and associated organelles until it envelops the daughter cell, leaving an opening at the apical end, where the APR is located, and the basal end, where the basal complex resides (Figure 4—figure supplement 3a). While the pellicle was easily visualized as a whole, we were unable to distinguish the IMC membranes from the PPM (Figure 4—figure supplement 3b and c). We stained parasites using the plasma membrane marker MSP1 and two different IMC markers: GAP45, which lies between the IMC and PPM, and IMC1g, which is attached to the cytoplasmic face of the IMC (Blackman et al., 1994; Cepeda Diaz et al., 2023; Kono et al., 2012). In both cases, we were unable to resolve the IMC marker from MSP1 (Figure 4—figure supplement 3b and c).

Basal complex dynamics throughout segmentation

The basal complex is an essential ring structure located at the basal end of the IMC (Morano and Dvorin, 2021). It is hypothesized to act as a contractile ring that guides IMC biogenesis and mediates abscission of newly formed merozoites by separating the IMC and plasma membrane from the residual body. We used parasites where PfCINCH, a basal complex marker, was tagged with a spaghetti monster V5 (smV5) tag to follow basal complex development throughout schizogony (Figure 4a; Rudlaff et al., 2019). CINCH is first visible at early schizogony (3–5 nuclei stage) as a small ring-like structure surrounding an NHS ester-dense focus on the plasma membrane that is tethered to the CP (Figure 4b). Of 55 early schizont CPs imaged, 44 (80%) had matching numbers of outer CP branches and basal complex structures. This suggests that as CPs divide, they each inherit a CINCH ring that has been split by the duplication of the cytoplasmic tethers (Figure 4b). Early IMC proteins have been described to form cramp-like structures like these prior to attaining their characteristic ring structure later in schizogony (Hu et al., 2010; Kono et al., 2012). During the rapid nuclear divisions of schizogony, 77% of the CINCH structures of mitotic CPs show a break in the ring (Figure 4—figure supplement 1b). This break faces a sister basal complex with its own cytoplasmic extension (Figure 4b, Figure 4—figure supplement 1b). These observations suggest that, upon duplication of the outer CP branches, the basal complex ring likely ‘breaks’ into two semicircles, which re-seal to form their own ring prior to the next branch duplication (Figure 4b, Figure 4—figure supplement 1b).

Figure 4 with 3 supplements see all
Basal complex biogenesis and development throughout segmentation.

Parasites expressing an smV5-tagged copy of the basal complex marker CINCH were prepared by ultrastructural expansion microscopy (U-ExM), stained with N-hydroxysuccinimide (NHS) ester (grayscale), BODIPY TRc (white), SYTOX (cyan), and anti-V5 (basal complex; magenta) antibodies and imaged using Airyscan microscopy across segmentation. (a) Images of whole parasites throughout asexual blood-stage development. (b) Basal complex development during schizogony. The basal complex is formed around the parasite plasma membrane (PPM) anchor of the outer centriolar plaque (CP). In nuclei whose outer CP has two branches, and will therefore undergo mitosis, the basal complex rings are duplicated. From early segmentation, the basal complex acquires a stable, expanding ring form. Cytostomes that will form part of merozoites are marked with a white asterisk, while those outside merozoites are marked with a yellow asterisk. Images are maximum-intensity projections, number on image = Z-axis thickness of projection in µm. Scale bars = 2 µm.

Once segmentation begins and the outer CP stops duplicating, CINCH forms a bona fide ring with smooth borders (Figure 4). This matches previously reported behavior of early IMC proteins and supports our hypothesis that the cramp-like structures arise from IMC and basal complex division. At this point, nuclei reach a point of semi-synchronicity. All 64 imaged CPs in early segmentation parasites had duplicated and were forming a mitotic spindle. This event marks the last nuclear division the parasite will undergo. Each of these CPs had a single uninterrupted basal complex ring (Figure 4b). Thus, we cease to observe events where a single nucleus is attached to four basal complexes as CPs have ceased committing to future rounds of mitosis. As the parasite undergoes segmentation, the basal complex expands and starts moving in the basal direction. By the time the basal complex reaches its maximum diameter, all nuclear divisions have been completed, each nucleus has a single CP and basal complex, and no mitotic spindles are visible (Figure 4). After this point, the basal complex contracts and continues to move away from the apical end. By the time segmentation is completed, the basal complex is an NHS ester-dense ring that is smaller than the APR.

NHS ester as a basal complex marker

While the basal complex stains brightly with NHS ester at the end of segmentation (Figure 4), this staining is not consistent throughout schizogony. NHS ester staining of the basal complex is not visible or is very faint during early schizogony. Once the basal complex attains its bona fide ring form during early segmentation, it stains reliably, though faintly, with NHS ester. This staining intensifies after the basal complex begins to contract. The denser staining observed during basal complex contraction could be due to recruitment of more basal complex proteins at the midpoint of schizogony, an increase in protein density as the ring area decreases during contraction, or both. Once the parasites finish segmentation, the basal complex is at its brightest (Figure 4b). While NHS ester staining correlates with CINCH, it does not perfectly overlap with it. CINCH consistently appears as a larger ring with a slight basal shift relative to NHS ester after the basal complex reaches maximum diameter, an effect most visible at the end of segmentation (Figure 4b). Since this shift is consistent with parasite anatomy regardless of parasite orientation, it suggests it is not an imaging artifact. There is no primary antibody against CINCH at this point, and so it is not possible to determine whether the lack of overlap with NHS ester is due to distance between the smV5 tag and the main protein density of CINCH (CINCH is 230 kDa). It is also possible that this difference in localization reflects basal complex architecture similar to that previously observed in Toxoplasma gondii, where the basal complex consists of multiple concentric rings (Anderson-White et al., 2012; Engelberg et al., 2022; Hu, 2008; Roumégous et al., 2022).

Mitochondrion and apicoplast

The apicoplast and mitochondrion undergo pronounced morphological changes during the P. falciparum blood-stage lifecycle (van Dooren et al., 2005; van Dooren et al., 2006). Both are long, and often branching, organelles whose complex three-dimensional morphologies have only been robustly studied using electron microscopy-based techniques (Rudlaff et al., 2020).

Looped regions of the mitochondrion display low membrane potential

To visualize the mitochondria, we stained live parasites using MitoTracker Orange CMTMRos prior to fixation and expansion (Figure 5—figure supplement 1a). MitoTracker Orange CMTMRos accumulates in live mitochondria, driven electrophoretically by membrane potential, and is retained after fixation (Elmore et al., 2004; Poot et al., 1996). When imaged at high resolution, MitoTracker can be used to observe individual cristae in the mitochondria of mammalian cells (Wolf et al., 2019). Plasmodium cristae morphology is different from that found in mammalian mitochondria; cristae are thought to be bulbous or tubular rather than lamellar and are present in gametocytes but absent from asexual blood-stage parasites (Evers et al., 2021; Evers et al., 2023). To our surprise, rather than showing continuous staining of the mitochondria, MitoTracker staining of our expanded parasites revealed alternating regions of bright and dim staining that formed MitoTracker-enriched pockets (Figure 5—figure supplement 1b). These clustered areas of MitoTracker staining were highly heterogeneous in size and pattern. Small staining discontinuities like these are commonly observed in mammalian cells when using MitoTracker dyes due to the heterogeneity of membrane potential from cristae to cristae as well as due to fixation artifacts. At this point, we cannot determine whether the staining we observed represents a true biological phenomenon or an artifact of this sample preparation approach. Our observed MitoTracker-enriched pockets could be an artifact of PFA fixation, a product of local membrane depolarization, a consequence of heterogeneous dye retention, or a product of irregular compartments of high membrane potential within the mitochondrion, to mention a few possibilities. Further research is needed to conclusively pinpoint an explanation.

In addition to these small staining discontinuities, we observed large gaps in MitoTracker staining within parasites at all stages of development. This included pre-segmentation parasites, where we would expect a single continuous mitochondrion to be present (Figure 5—figure supplement 1a). To our knowledge, no membrane potential discontinuities or fixation artifacts of this size have been reported in mammalian cells. So, as a secondary way to visualize the mitochondria and better characterize these staining discontinuities, we generated a transgenic cell line with the putative ATP synthase F0 subunit-d (ATPd, Pf3D7_0311800) tagged with a spaghetti monster HA tag (Viswanathan et al., 2015; Figure 5—figure supplement 2). ATPd is a membrane-embedded proton channel that had not previously been localized to the mitochondria in P. falciparum but was identified as a mitochondrial protein in a recent proteomics study (van Esveld et al., 2021; Evers et al., 2021). Furthermore, its Toxoplasma homolog has been shown to localize to the mitochondria (Barylyuk et al., 2020; Sheiner et al., 2011). We confirmed that ATP synthase subunit F0 localizes to P. falciparum mitochondria, as it largely co-localized with MitoTracker staining, forming a border around it due to its membrane association (Figure 5a, Figure 5—figure supplement 1a). ATPd, like MitoTracker, had a heterogeneous distribution throughout the mitochondria, but it did not show the same large gaps in staining. ATPd allowed us to better visualize regions of the mitochondria that appeared to fold onto themselves and fuse with each other, as has been previously described (van Dooren et al., 2005). Thus, MitoTracker and ATPd are both useful but imperfect markers for the mitochondria, with neither of them showing a continuous, even distribution throughout the organelle.

Figure 5 with 2 supplements see all
Growth and fission of the mitochondrion.

Parasites with an smHA-tagged copy of the ATP Synthase F0 Subunit D (ATPd, Pf3D7_0311800) as a mitochondrial marker were prepared by ultrastructural expansion microscopy (U-ExM), stained with N-hydroxysuccinimide (NHS) ester (grayscale), BODIPY TRc (white), SYTOX (cyan), and anti-HA (mitochondrion; magenta) antibodies and imaged using Airyscan microscopy. (a) Images of whole parasites throughout asexual blood-stage development. Maximum-intensity projections of both a subsection of the cell (partial mito) and the full cell (full mito) are shown. (b) ATPd staining was compared against MitoTracker Orange CMTMRos (yellow), which showed discontinuous staining in looped regions. (c) Area of the mitochondrion was quantified for parasites of varying nucleus number. Seventy-three cells were counted across four biological replicates. ****p<0.001, ns = p >0.05 by one-way ANOVA, error bars = SD. (d) Schizont with mitochondria that have undergone fission (yellow zoom), mitochondria that are shared between two nascent merozoites (black zoom), and mitochondria left outside merozoites in the forming residual body (gray). (e) During fission, mitochondria associate with the outer centriolar plaque (oCP). Images are maximum-intensity projections, number on image = Z-axis thickness of projection in µm. White scale bars = 2 µm, yellow scale bars = 500 nm.

Curiously, 25 of 26 imaged parasites showed MitoTracker discontinuities specifically in regions where the ATPd signal formed looped structures (Figure 5b). These structures were defined as areas where the mitochondria showed a turn or fold of ~180°. Of the 41 looped regions identified, 75% lacked MitoTracker staining. This suggests that mitochondria looped regions in P. falciparum have some degree of depolarization that prevents MitoTracker accumulation or that MitoTracker initially accumulates in these regions but is not bound and retained. The biological significance of these areas, if any, is currently unclear.

Growth of the apicoplast and mitochondrion

To visualize the apicoplast, we utilized a previously established cell line that expresses GFP fused to the apicoplast transit peptide of acyl carrier protein (ACP) (Florentin et al., 2020), which we will refer to as apicoplast-GFP (Figure 6a). This marker allowed for a relatively even and continuous staining of the organelle. We quantified mitochondrion and apicoplast signal area using ATPd-smHA and apicoplast-GFP respectively as a proxy measurement of size (Figures 5c and 6b). Tracking this in parallel to parasite nucleus number allowed us to determine whether the growth of these organelles occurred progressively with simultaneous rounds of mitosis and nuclear division. In mononucleated ring and trophozoite parasites, both the mitochondria and the apicoplast are relatively small, having an average area of 13.39 µm2 (±15.1 µm2 SD) and 4.81 µm2 (±2.62 µm2 SD), respectively, in expanded parasites (Figures 5c and 6b). As expected from live cell observations (van Dooren et al., 2005), both organelles show significant growth and spread throughout the cell in multinucleated parasites, adopting an elongated and branching morphology (Figures 5 and 6). Mitochondria grow almost exclusively during the first two rounds of nuclear replication, achieving an average size of 87.9 µm2 (±15.9 µm2 SD) at the 2–5 nuclei stage. This size remains relatively constant until segmentation, with the average mitochondria size right before the start of fission being 99.6 µm2 (±37.4 µm2 SD) (Figure 5c). In contrast, the apicoplast continues to grow past the 2–5 nuclei stage, having an average size of 25.5 µm2 (±7.21 µm2 SD) at the 2–5 nuclei stage and 34.97 µm2 (±11.23 µm2 SD) in cells with >15 nuclei (Figure 6b). These data suggest that the mitochondrion and apicoplast do not grow simultaneously with or as a response to nuclear replication during schizogony. Rather, both organelles show the largest increase in size during the 1–2 nuclei transition and either plateau in size, in the case of the mitochondria, or enter a second phase of slower growth that ends shortly before segmentation, in the case of the apicoplast.

Growth and fission of the apicoplast.

Parasites expressing GFP-conjugated to the apicoplast transit signal of ACP (ACPTs-GFP) were prepared by ultrastructural expansion microscopy (U-ExM), stained with N-hydroxysuccinimide (NHS) ester (grayscale), BODIPY TRc (white), SYTOX (cyan), and anti-GFP (apicoplast) (magenta) antibodies and using Airyscan microscopy. (a) Images of whole parasites throughout asexual blood-stage development. Maximum-intensity projections of both a subsection of the cell (partial mito) and the full cell (full mito) are shown. (b) Area of the apicoplast was quantified for parasites of varying nucleus number. Seventy cells were counted across three biological replicates. ****p<0.001, *p<0.05 by one-way ANOVA, error bars = SD. (c) Representative images of the different stages of apicoplast fission. Images are maximum-intensity projections, number on image = Z-axis thickness of projection in µm. Asterisks represent centriolar plaques. Scale bars = 2 µm.

Fission of the mitochondrion and apicoplast

P. falciparum has a single, large, branching, mitochondrion and apicoplast throughout most of the asexual blood stage (van Dooren et al., 2005; van Dooren et al., 2006; Verhoef et al., 2021). During segmentation, however, these organelles undergo fission such that each merozoite inherits an individual apicoplast and mitochondrion (Rudlaff et al., 2020; van Dooren et al., 2005). While it has been shown that apicoplast fission occurs before mitochondrial fission, it is unclear how fission occurs (Rudlaff et al., 2020). A recent review (Verhoef et al., 2021) posed three possible mechanisms: synchronous fission where the organelle simultaneously divides into all daughter parasites at once, outside-in fission where fission occurs at the ends of the organelle, or branching point fission where a first fission event divides the organelle into larger segments and a subsequent fission event leaves each merozoite with an individual organelle (Verhoef et al., 2021). It also remains unclear how accurate segregation into daughter cells is monitored. In T. gondii, the apicoplast associates with the centrosomes prior to undergoing fission. A similar association between apicoplasts and CPs has been proposed in Plasmodium but still lacks evidence due to the difficulty of observing the Plasmodium CP in live cells (van Dooren et al., 2005). The mitochondrion is not thought to associate with the CP in Toxoplasma or Plasmodium, and its mechanism for ensuring accurate segregation remains unknown.

In the process of imaging the mitochondria and apicoplasts of segmenting parasites, we observed a transient outer CP association prior to and during fission in both organelles. Just before the start of segmentation, there is little association between the outer CP and apicoplast. Of five parasites imaged that had >10 nuclei but had not yet started segmentation, three had no contact points between the outer CP and apicoplast branches and the other two had <4 contact points. Once segmentation starts, all apicoplast branches contact an outer CP each and remain in contact with the outer CP until the end of apicoplast fission and CP degradation (Figure 6c). In 11 imaged segmenting schizonts, all outer CPs showed contact with an apicoplast branch each. This association starts in early segmentation, when a single branched apicoplast connects all outer CPs. By the time the basal complex reaches maximum diameter and begins contraction, we observe seven parasites where all branches have completed fission and two parasites that had at least one or more apicoplast segments still connecting multiple merozoites (Figure 6c). Mitochondria fission follows a very similar pattern, but later in parasite development. Prior to the basal complex reaching maximum diameter, we observe no significant connection between the outer CP and mitochondria. When the basal complex entered its contraction phase, we observed nine parasites where all outer CPs were in contact with one branch of the mitochondria each (Figure 5d and e). Only one of these nine parasites had an intact pre-fission mitochondrion, while the other eight had undergone at least one fission event. Matching previous descriptions of mitochondria and apicoplast segmentation, C1-arrested schizonts having completed segmentation show elongated mitochondria (Figure 5a) and small, rounded apicoplasts (Figure 6a).

Both mitochondria and apicoplast fission showed neighboring nascent merozoites that shared a single branch of mitochondria or apicoplast passing through both of their basal complexes while others had an individual mitochondrion or apicoplast that had already separated from the rest (Figures 5d and 6c). This suggests that fission does not occur synchronously (Figure 5d) and supports the model of branching point fission. In other words, parasites seem to undergo a primary fission event that leaves only some merozoites sharing stretches of the organelles and then a subsequent fission event leaves each merozoite with an individual apicoplast and mitochondrion (Figure 5e). Unfortunately, BODIPY TRc does not distinctly stain the membranes of the mitochondria and apicoplast. So, it is not possible for us to determine whether the observed breaks in staining of our chosen organelle markers truly indicate a complete fission of the mitochondria or apicoplast membranes. Thus, while suggestive of branching point fission, our data is not sufficient to conclusively determine the sequence of fission events in these organelles. More research with additional mitochondrial and apicoplast markers is needed to confirm the observations made in this study and conclusively map out the growth and fission of these organelles.

Characterization of residual body mitochondria

At the completion of segmentation, the parasite forms a structure known as the residual body, which contains parasite material, such as the hemozoin crystal, that was not incorporated into merozoites during segmentation (Rudlaff et al., 2020). The residual body is poorly understood in Plasmodium, but in Toxoplasma it has been shown that a significant amount of the mitochondria, and not the apicoplast, is left behind in the residual body following segmentation (Nishi et al., 2008).

There is no well-characterized marker of the residual body in Plasmodium. So, for this study, we defined the residual body as any area within the parasitophorous vacuole membrane but visibly external to any merozoite in a C1-arrested schizont as determined by BODIPY TRc staining. We imaged 35 C1-arrested schizonts and observed that 54% had mitochondrial staining inside the residual body (Figure 5d, Figure 5—figure supplement 1c). To determine the proportion of total mitochondria that gets included in the residual body, we quantified the fluorescence of both mitochondria in the residual body and mitochondria in merozoites. Of the 19 parasites that showed mitochondria staining inside the residual body, the amount of material ranged from 1 to 12% of the total mitochondrial staining in the parasite (Figure 5—figure supplement 1d). On average, the residual body had approximately 1.5× more mitochondrial staining than the average merozoite (Figure 5—figure supplement 1e). No significant apicoplast staining was ever observed in the residual body, similarly to what has been reported for Toxoplasma (Nishi et al., 2008).

Cytostomes

During its intraerythrocytic development, P. falciparum engulfs host cell cytoplasm from which it catabolizes hemoglobin as a source of amino acids (Francis et al., 1997). The parasite is separated from its host cell by the parasitophorous vacuole, and therefore the uptake of host-cell cytosol requires invagination of both the PPM and PVM. The cytostome coordinates this endocytic process and is comprised of two key regions: a protein-dense collar region, which forms the pore through which membrane invagination will occur, and the membranous bulb region, which contains the RBC-derived cargo (Milani et al., 2015; Xie et al., 2020).

NHS ester staining reveals pore-like structures at the PPM

Prior to this study, cytostomes were not immediately obvious by NHS ester staining given the large number of features that were visible using this stain but pending validation. While observing the basal complex of segmenting schizonts (Figure 4a), we noticed that merozoites contained a second NHS ester-dense ring (Figure 7a). The size and position of this NHS ester ring matched that of an endocytic micropore recently identified in Toxoplasma tachyzoites (Koreny et al., 2022). In that study, the micropore was identified using Kelch13 (K13) as a marker (Koreny et al., 2022). The Plasmodium equivalent to this K13 micropore is the cytostome, so to determine whether this NHS ester-dense ring was indeed a cytostome, we evaluated a parasite strain where the endogenous K13 was fused to GFP (Birnbaum et al., 2017). Investigation using this parasite line revealed that the NHS ester-dense ring also stained with K13, suggesting that this structure is a cytostome (Figure 7b).

Figure 7 with 1 supplement see all
Cytostomes are observable by ultrastructural expansion microscopy (U-ExM) throughout the asexual blood stage of the lifecycle.

(a) Parasites expressing an smV5-tagged copy of the basal complex marker CINCH, prepared by U-ExM, and stained with anti-V5, show N-hydroxysuccinimide (NHS) ester-dense rings that are negative for this basal complex marker. (b) Parasites prepared by U-ExM where the cytostome marker Kelch13 was conjugated to GFP (K13-GFP) and stained with anti-GFP. Image shows co-localization between K13-GFP and the putative cytostome. NHS ester-dense ring. (c) Comparison between paraformaldehyde (PFA)-only and PFA-glutaraldehyde fixed U-ExM parasites, showing lysed (PFA only, orange) and intact (PFA-glutaraldehyde, magenta) red blood cell (RBC) membranes. (d) In PFA-only fixed parasites, only the cytostomal collar is preserved, while both the collar and bulb are preserved upon PFA-glutaraldehyde fixation. (e) K13-GFP parasites were either fixed in PFA only or PFA-glutaraldehyde, prepared by U-ExM, stained with NHS ester (grayscale), SYTOX (cyan), and anti-GFP (cytostome) (magenta) antibodies and imaged using Airyscan microscopy across the asexual blood stage. Zoomed regions show cytostomes. Images are maximum-intensity projections, number on image = Z-axis thickness of projection in µm. White scale bars = 2 µm, yellow scale bars = 500 nm.

PFA-glutaraldehyde fixation allows visualization of cytostome bulb

The cytostome can be divided into two main components: the collar, a protein-dense ring at the PPM where K13 is located, and the bulb, a membrane invagination containing RBC cytoplasm (Milani et al., 2015; Xie et al., 2020). While we could identify the cytostomal collar by K13 staining, these cytostomal collars were not attached to a membranous invagination. Fixation using 4% v/v PFA is known to result in the permeabilization of the RBC membrane and loss of its cytoplasmic contents (Tonkin et al., 2004). Topologically, the cytostome is contiguous with the RBC cytoplasm, and so we hypothesized that PFA fixation was resulting in the loss of cytostomal contents and obscuring of the bulb. PFA-glutaraldehyde fixation has been shown to better preserve the RBC cytoplasm (Tonkin et al., 2004). Comparing PFA only with PFA-glutaraldehyde fixed parasites, we could clearly observe that the addition of glutaraldehyde preserves both the RBC membrane and RBC cytoplasmic contents (Figure 7c). Further, while only cytostomal collars could be observed with PFA-only fixation, large membrane invaginations (cytostomal bulbs) were observed with PFA-glutaraldehyde fixation (Figure 7d). Cytostomal bulbs were often much longer and more elaborate spreading through much of the parasite (Video 1), but these images are visually complex and difficult to project, so images displayed in Figure 7 show relatively smaller cytostomal bulbs. Collectively, this data supports the hypothesis that these NHS ester-dense rings are indeed cytostomes and that endocytosis can be studied using U-ExM, but PFA-glutaraldehyde fixation is required to maintain cytostome bulb integrity.

Video 1
Slice-by-slice view of paraformaldehyde/glutaraldehyde fixed parasite.

Schizont stage parasite fixed with paraformaldehyde and glutaraldehyde to preserve erythrocyte membrane, prepared using ultrastructural expansion microscopy (U-ExM) and stained with N-hydroxysuccinimide (NHS) ester (protein density, grayscale) and SYTOX (DNA, cyan). This slice-by slice video through the parasite shows the elongated, protein-dense, cytostome invaginations in the parasite.

We subsequently harvested K13-GFP parasites across the parasite lifecycle and imaged them following either PFA only or PFA-glutaraldehyde fixation. K13-stained cytostomes were detected at all stages of the parasite lifecycle (Figure 7e). Ring-stage parasites typically contained one or two cytostomes, which increased in number during the trophozoite stage and schizogony (Figure 7e).

Single cytostomes appear in the area containing the IMC near the apical organelles at the same time as the basal complex forms a complete ring. Cytostomes remain within the IMC area but change positions within the nascent merozoite as segmentation progresses (Figure 4b, white asterisks). The majority of merozoites in C1-arrested schizonts contained a single cytostome. This suggests that cytostomes are incorporated into the IMC of merozoites and inherited early in segmentation. Clusters of cytostomes that had not been incorporated into merozoites during segmentation were observed either adjacent to nascent merozoites or as part of the residual body (Figure 7—figure supplement 1 and Figure 4b, yellow asterisk). It is currently unclear whether there are any functional differences between the cytostomes that are incorporated into merozoites and those that are left behind.

Non-canonical cytostome collar morphologies

We noticed a number of different cytostome morphologies and organizational patterns (Figure 7—figure supplement 1). Cytostomes frequently clustered together and did not appear randomly distributed across the PPM (Figure 7e). Some cytostomes would form what appeared to be higher order structures where two or three distinct cytostomal collars appeared to be stacked end-on-end (Figure 7—figure supplement 1). Cytostomes have a relatively well-defined and consistent size (Aikawa, 1971; Yang et al., 2019), but occasionally we observed very large cytostomal collars that were approximately twice the diameter of other cytostome collars in the same cell (Figure 7—figure supplement 1). It is unclear what the function of these higher order structures or large cytostomes is, if they represent biogenesis transition states, or indeed if they are performing some specialized endocytosis.

The rhoptries

To invade host RBCs, merozoites secrete proteins from specialized secretory organelles known as the rhoptries and micronemes. While both the rhoptries and micronemes are well studied in the context of Plasmodium biology, neither have been investigated in detail using expansion microscopy in Plasmodium. We previously showed that fully formed rhoptries can be observed by NHS ester staining alone (Liffner and Absalon, 2021), but did not investigate their biogenesis.

Rhoptries can be observed from early in their biogenesis using NHS ester staining

Rhoptries consist of a neck and bulb region, with the tip of the neck being loaded into the APRs of merozoites. Despite both being formed from Golgi-derived cargo, the neck and bulb regions have distinct proteomes (Counihan et al., 2013). We first tracked rhoptry bulb biogenesis across schizogony using antibodies directed against the rhoptry bulb marker rhoptry-associated protein 1 (RAP1).

Nascent rhoptries were detected early in schizogony, with RAP1 foci appearing adjacent to all branches of the outer CP from parasites with 6–10 nuclei in a one-to-one ratio, as described above (Figure 8a, Figure 8—figure supplement 1b). These foci co-localized with NHS ester densities of the same size and round shape, no elongated neck-like structures were visible by NHS ester (neck biogenesis described in more detail below). This matches reports that rhoptry bulb biogenesis occurs first (Bannister et al., 2000; Counihan et al., 2013), with neck biogenesis not occurring until segmentation. As early as the last mitotic event during early segmentation, rhoptry bulbs were observed as pairs, with 88 of 93 (95%) CPs observed forming a mitotic spindle being associated with two RAP1-positive NHS ester densities per outer CP branch. Finally, in newly invaded ring-stage parasites, strong RAP1 staining was observed at the PPM/PVM (Figure 8—figure supplement 1a), supporting previously reported observations that secreted RAP1 coats the merozoite during invasion (Riglar et al., 2011).

Figure 8 with 1 supplement see all
Rhoptries undergo biogenesis near the centriolar plaque and are segregated during nuclear division.

3D7 parasites were prepared by ultrastructural expansion microscopy (U-ExM), stained with N-hydroxysuccinimide (NHS) ester (grayscale), BODIPY TRc (white), SYTOX (cyan), and an anti-rhoptry antibodies and imaged using Airyscan microscopy. (a) Images of whole parasites throughout schizogony stained using an anti-RAP1 (rhoptry bulb; magenta) antibody. (b) Zoom into rhoptry pairs of 3D7 parasites that were prepared for U-ExM and stained with NHS ester (grayscale) along with antibodies against RAMA (rhoptry bulb; magenta) and RON4 (rhoptry neck; yellow) to assess rhoptry neck biogenesis. We observed that the rhoptry neck begins as a single focus inside each rhoptry. Rhoptries then get duplicated and segregated alongside the centriolar plaque. During the final mitosis, the rhoptry neck begins to elongate and the rhoptries separate from centriolar plaque. Images are maximum-intensity projections, number on image = Z-axis thickness of projection in µm. Scale bars = 2 µm.

Our data not only suggests that rhoptry biogenesis occurs well before segmentation, when nuclei still have several rounds of mitosis to complete, but also that rhoptries remain CP-associated during these mitotic events. Instances of this association with the CP have been observed before (Bannister et al., 2000; Rudlaff et al., 2020) but its mechanism remains unknown. To our knowledge, this is the first in-depth documentation of a rhoptry–CP association throughout schizogony in Plasmodium.

Rhoptry heterogeneity during early schizogony and segmentation

Rhoptries associated with the same CP during early schizogony sometimes differ in size (Figure 8—figure supplement 1b). This is not surprising given the speed of mitotic events requires near-constant biogenesis of new rhoptry bulbs. By the time segmentation is underway, instead of inheriting one sister rhoptry in the final mitotic event of schizogony, each CP will inherit a pair of rhoptries each. At this point, the speed of these mitotic events slows and parasites reach a point of semi-synchrony. To our surprise, this synchrony does not extend to rhoptry pairs; the two rhoptries inherited by segmenting daughter cells remain different from each other. This heterogeneity in rhoptry pairs during early segmentation has been documented before by electron microscopy (Bannister et al., 2000; Rudlaff et al., 2020). Of 109 rhoptry pairs imaged in early segmentation schizonts undergoing their last miotic event (where CPs were observed forming mitotic spindles), only 4% had two rhoptries of similar size and density (Figure 8—figure supplement 1d). We observed that 40% of these 109 rhoptry pairs had different size but equal NHS ester density, 21% had the same size but different NHS ester density, and 35% differed in both size and NHS ester density (Figure 8—figure supplement 1e). As expected from previous reports, this heterogeneity was lost after the completion of this last miotic event and rhoptry neck elongation. Of 98 rhoptry pairs imaged in non-mitotic segmenting parasites, 76% had two rhoptries of similar size and density (Figure 8—figure supplement 1d). It is still unclear how the one-to-two rhoptry transition occurs and whether rhoptry heterogeneity has a biological role in biogenesis and maturation of the organelles.

Overall, we present three main observations suggesting that rhoptry pairs undergo sequential de novo biogenesis rather than dividing from a single precursor rhoptry. First, the tight correlation between rhoptry and outer CP branch number suggests that either rhoptry division happens so fast that transition states are not observable with these methods or that each rhoptry forms de novo and such transition states do not exist. Second, the heterogeneity in rhoptry size throughout schizogony favors a model of de novo biogenesis given that it would be unusual for a single rhoptry to divide into two rhoptries of different sizes. Lastly, well-documented heterogeneity in rhoptry density suggests that, at least during early segmentation, rhoptries have different compositions. Heterogeneity in rhoptry contents would be difficult to achieve so quickly after biogenesis if they formed through fission of a precursor rhoptry. While constant de novo biogenesis could explain why one rhoptry can appear smaller or less mature than the other (Bannister et al., 2000; Rudlaff et al., 2020), it is currently unclear why heterogeneity in rhoptry density only appears during early segmentation and not earlier. Thus, this model is not enough to explain all the variation in rhoptry size and density observed throughout schizogony. Furthermore, a lot of unknowns remain about what exactly governs rhoptry number during the rapid rounds of asynchronous nuclear division (Klaus et al., 2022), how the transition to a rhoptry pair is signaled, and how many rounds of de novo rhoptry formation parasites undergo.

Rhoptry neck biogenesis and elongation

In order to observe rhoptry neck biogenesis in more detail, we stained parasites against the rhoptry apical membrane antigen (RAMA, a rhoptry bulb marker) and rhoptry neck protein 4 (RON4, a rhoptry neck marker) (Richard et al., 2010; Topolska et al., 2004). RAMA is anchored to the rhoptry bulb membrane and only stains the periphery of the rhoptry bulb as marked by NHS ester (Figure 8b, Figure 8—figure supplement 1c). RON4 is absent from the earliest rhoptry bulbs, appearing as a focus within the rhoptry bulb shortly before early segmentation and before the rhoptry neck could be distinguished from the bulb by NHS ester staining alone (Figure 8b). During early segmentation, when rhoptry pairs first become visible, we observe an uneven distribution of RON4 within each pair. RON4 preferentially associates with one of the rhoptries, with the staining on the second rhoptry being fainter, more diffuse, or even absent in some cases. Of 84 rhoptry pairs observed at this stage, 72 (86%) showed an uneven distribution of RON4. To our surprise, when these rhoptry pairs were of different NHS ester densities, the larger share of RON4 associated with the less dense rhoptry (Figure 8b). Previous observations of rhoptry density differences by electron microscopy have been ascribed to differences in rhoptry age or maturity, with the denser rhoptry being more mature. So, finding RON4 to be more abundant in the less dense rhoptry suggests that either heterogeneous RON4 accumulation cannot be explained by rhoptry age or that the less dense rhoptry is instead the older rhoptry. The RON4-positive rhoptry neck elongates during segmentation, attaining its characteristic shape by mid to late segmentation and becoming observable by both RON4 staining and NHS ester (Figure 8). At this point, nearly all rhoptry necks had an equal distribution of RON4 (of 76 rhoptry pairs observed at these stages, 72 [95%] had an equal distribution of RON4).

The micronemes

Previous studies have suggested that micronemes may be heterogeneous and that apical membrane antigen 1 (AMA1) and other micronemal markers such as erythrocyte binding antigen-175 (EBA175) reside in different subsets of micronemes (Absalon et al., 2018; Ebrahimzadeh et al., 2019; Healer et al., 2002). We reasoned that individual micronemes may be visible using U-ExM and imaged parasites stained with AMA1 and EBA175 to observe their biogenesis and relative distribution. The first microneme marker to appear during schizogony was AMA1. Large puncta of AMA1 appear near the rhoptries when the basal complex is at its maximum diameter (Figure 9a). At this point, EBA175 is not yet detectable above background fluorescence. At the end of segmentation, we observe AMA1 has arranged itself into small, densely arranged puncta below the APR and around the rhoptry neck. We also observe EBA175 staining in puncta that are less densely arranged and have little co-localization with AMA1. EBA175 puncta are basal to the AMA1 puncta, being closer to the rhoptry bulb. They also form a cloud of larger diameter than the one formed by AMA1 such that, when viewed from above the APR, two concentric clouds are observed with the core being AMA1 positive and the periphery being EBA175 positive (Figure 9b and c). We could observe a punctate NHS ester staining pattern at the apical end of merozoites (Figure 9b), which we reasoned could be micronemes. The AMA1 and EBA175 staining we observed in late-stage schizonts partially overlaps with this punctate NHS ester pattern, suggesting that NHS ester punctae are micronemes (Figure 9c). However, many NHS ester-positive foci did not stain with either AMA1 or EBA175 despite being morphologically indistinguishable from those which did. This suggests that NHS ester stains more than just the micronemes and that some of these foci may be exonemes, dense granules, or other apical vesicles. Alternatively, it is also possible that these additional NHS ester-positive foci represent micronemes that lack both AMA1 and EBA175.

Micronemal proteins AMA1 and EBA175 reside in separate micronemes.

3D7 parasites were prepared by ultrastructural expansion microscopy (U-ExM), stained with N-hydroxysuccinimide (NHS) ester (grayscale), SYTOX (cyan), and antibodies against the micronemal markers AMA1 (magenta) and EBA-175 (yellow), and imaged using Airyscan microscopy in segmenting schizonts. (a) Images of whole parasites throughout schizogony. In schizonts still undergoing segmentation, AMA1 localized to the apical end while EBA175 was not detected. In late schizonts, both EBA175 and AMA1 were present in the micronemes. In E64-arrested schizonts, AMA1 was translocated to the merozoite surface while EBA175 remained micronemal. 3D rendering (b) and zooms (c) of merozoites with either micronemal or translocated AMA1. Foci of AMA1 and EBA175 to not routinely co-localize with each other. Images are maximum-intensity projections, number on image = Z-axis thickness of projection in µm. White scale bars = 2 µm, RGB scale bars for 3D rendering = 1 µm.

Using the protease inhibitor E64, we arrested parasites ‘post-egress’ such that AMA1 was translocated. E64 allows for normal daughter cell maturation but prevents RBC plasma membrane rupture once segmentation is complete (Hale et al., 2017). We observe that once AMA1 is translocated, the density of apical AMA1 decreases. EBA175, which does not translocate, increases in density, and moves apically, taking the space AMA1 occupied prior to translocation (Figure 9c and d). This is consistent with existing models of sequential microneme translocation and microneme fusion near the APR (Dubois and Soldati-Favre, 2019).

Discussion

In this work, we apply U-ExM to the asexual blood stage of P. falciparum to provide new insights into the role of the CP in establishing apical–basal polarity and in coordinating organelle segregation during schizogony (Figure 10). In the process, we demonstrate U-ExM can be used to study the biogenesis and protein distribution of a variety of organelles and structures within the parasite. Globally, our observations suggest that the CP is involved in establishing apical–basal polarity within the parasite and that this polarity is established early in schizogony.

Summary of organelle organization and fission during schizogony.

(a) Apical organelle biogenesis: Biogenesis of the rhoptries, Golgi, basal complex, and apical polar rings occur at outer centriolar plaque (CP), between the nuclear envelope and parasite plasma membrane. Duplication and segregation of these organelles appears to be tied to CP duplication and segregation following nuclear division. Mid-segmentation merozoite: the rhoptry neck is distinguishable from the bulb, and AMA1-positive micronemes are present at the apical end of the forming merozoite. Each merozoite has inherited a cytostome. Subpellicular microtubules stretch the entire distance from the apical polar rings and the basal complex. The apicoplast has attached to the outer CP and begun fission. Mature merozoite: the parasite has completed segmentation, and each merozoite contains a full suite of organelles. The CP is no longer visible, and EBA175-positive micronemes are both visible and separate from AMA1-positive micronemes. (b) Model for fission of the mitochondrion and apicoplast. Prior to fission, both the apicoplast and mitochondrion branch throughout the parasite cytoplasm, before associating with the outer CP of each forming merozoite. For the apicoplast, this occurs during the final mitosis, but not until late in segmentation for the mitochondrion. Following outer CP association, the apicoplast and mitochondrion undergo a first fission event, which leaves an apicoplast and mitochondrion shared between forming merozoite pairs. Subsequently, both organelles undergo a second fission event, leaving each forming merozoite with a single apicoplast and mitochondrion.

We show that the branches of the outer CP appear to act as physical ‘tethers,’ connecting the nucleus to PPM. This creates a space between the PPM and nuclear envelope where we observe multiple parasite structures and organelles including the Golgi, basal complex, rhoptries, and APRs. These structures and organelles remain CP-associated despite the constant movement and segregation of CPs during mitotic events. In other systems, MTOC or centrosome positioning has been described as a source of cellular polarity, polarizing a cell by directing cargo of secretory organelles to a defined area (Cowan and Hyman, 2004; de Anda et al., 2005; Huse, 2012). While this has not been previously described in Plasmodium, this model fits well with our observation of the Golgi being adjacent to the CP throughout schizogony. This Golgi positioning could grant local control over the biogenesis of rhoptries, micronemes, dense granules, and IMC, which are all at least partially formed by Golgi-derived cargo (Counihan et al., 2013; Dubois and Soldati-Favre, 2019; Griffith et al., 2022; Sloves et al., 2012). While the biogenesis of the APRs seems to also be coupled to the CP, we do not yet know how they are nucleated or whether they are also dependent on Golgi-derived cargo.

This study also shows CP-coupled organelle segregation and biogenesis to an extent never observed before. Specifically, we observe that the Golgi, rhoptries, and basal complex match the branches of the outer CP in number throughout early schizogony and segregate with CPs during the mitotic events of schizogony. As this study used fixed cells, we lack the temporal power to precisely define the order of these events. Using known markers of parasite age, however, we describe putative transition states that nascent rhoptries and basal complexes adopt when being segregated with CPs. We put forward a model where outer CP-associated events occur before inner CP-associated events. That is, the duplication of the outer CP and organelles precedes inner CP duplication and the formation of a mitotic spindle such that this cytoplasmic duplication represents the first identifiable step in the commitment of a nucleus to the next round of mitosis. Interestingly, nuclei can sometimes have CPs forming a mitotic spindle that have already committed to the next round of mitosis by duplicating their cytoplasmic cargo. That is, a mitotic nucleus can show four outer CP branches and sets of apical organelles at the same time. While this supports our hypothesis that duplication of apical organelles and the outer CP components happens upstream from intranuclear mitotic events, it does not tell us how this relates to the genome copy number inside the nucleus. So, we cannot say whether DNA replication and associated checkpoints occur upstream or downstream from the cytoplasmic events that seem to commit a CP to mitosis.

Lastly, we contribute important evidence toward hypotheses across several open questions regarding organelle biogenesis and segregation in Plasmodium. We observe contacts between the outer CP and both the mitochondria and apicoplast during fission that suggest a role for this structure in monitoring copy number of these organelles. We also document sequential fission events in both the apicoplast and mitochondria that are suggestive of branching point fission. Lastly, we see temporal and spatial localization patterns of AMA1, EBA175, and RON4 that support theories of heterogeneity within the apical organelles.

U-ExM represents an affordable and adaptable sample preparation method that can be applied to any microscope to produce images with far greater visible detail than conventional light microscopy platforms. Applying this technique to the visualization of organelle biogenesis and segregation throughout schizogony allowed us to observe these processes in the context of structures that could previously only be investigated in detail using electron microscopy. The flexibility and scalability of this technique allowed us to image more than 600 individual parasites at a variety of developmental stages, increasing the confidence of our observations and giving them some temporal resolution. To our knowledge, this article represents the most comprehensive study of a single organism using U-ExM, with a total of 13 different subcellular structures investigated.

Our inability to pinpoint the nucleation site of the SPMTs or resolve the plasma membrane from the IMC highlights some of the limitations of U-ExM as applied in this work. The subpellicular network that holds IMC1g and lines the cytoplasmic face of the IMC sits approximately 20 nm below the parasite surface, where we would find MSP1 (Kudryashev et al., 2012). Thus, the distance between IMC1g and MSP1 post-expansion is around 90 nm. This is below our imaging resolution with Airyscan 2 and close to the maximum resolution we could achieve through other super-resolution methods compatible with our current setup when antibody effects are considered. Thus, some parasite structures remain beyond the resolution achieved in this study. In order to resolve the IMC from the plasma membrane or resolve multiple APRs using light microscopy, we would need to employ single-molecule localization microscopy or iterative expansion microscopy (Louvel et al., 2022), which increases expansion factor to ~20×.

We also noticed some drawbacks and artifacts introduced by U-ExM. Most visually striking was that the hemozoin crystal of the food vacuole does not expand (Coronado et al., 2014), which leaves a large space that lacks NHS ester staining. For nearly all antibodies used in this study, significant off-target fluorescence was observed inside the food vacuole. Thus, U-ExM may not be as useful for studying food vacuole biology. Occasionally, significant SYTOX (DNA stain) fluorescence was observed at either the nuclear envelope or PPM. It is unclear if this represents an expansion-induced artifact or a PFA-fixation artifact that is only now observable. Lastly, for cells stained with MitoTracker, some non-specific background was observed that seemed to correlate with protein density as observed by NHS ester.

Malaria parasites have been extensively studied using electron microscopy to determine their ultrastructure and live-cell microscopy to observe their most dynamic processes in real time. Much of what we uncovered in this study involved dynamic processes that are too small to be resolved using conventional live-cell microscopy. Specifically, we made important observations about P. falciparum organelle biogenesis and the organization of Plasmodium cell division around the CP. Rather than a replacement for any existing microscopy techniques, we see U-ExM as a complement to the suite of techniques available to study the cell biology of malaria parasites, which bridges some of the limitations of electron microscopy and live-cell microscopy. As such, there are many parasite processes that are logical candidates for investigation by U-ExM. Some of these have been highlighted in this article, but others remain completely unexplored. Merozoite invasion, for example, has been well-studied using a variety of microscopy techniques (Geoghegan et al., 2021; Hanssen et al., 2013; Liffner et al., 2022; Riglar et al., 2011; Weiss et al., 2015), but U-ExM would allow us to visualize how all the apical organelles associate with each other and rearrange in three dimensions across the established time course (Weiss et al., 2015) of this process. Another logical candidate for investigation by U-ExM is the rapid disassembly of the IMC and other merozoite organelles immediately following invasion (Ferreira et al., 2020), a process where the parasite undergoes rapid and dramatic morphological rearrangements that lie beyond the resolution of live-cell microscopy.

Materials and methods

Plasmodium falciparum culture

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Unless otherwise stated, all parasites in this study were 3D7-Cas9 (Rudlaff et al., 2019). For imaging of the apicoplast, the previously generated ACP-transit-peptide-GFP cell line was used (Florentin et al., 2020). For imaging of Kelch13, the previously generated 2xFKBP-GFP-K13 parasites were used (Birnbaum et al., 2017). For imaging of the basal complex, the previously generated CINCH-smV5 cell line was used (Rudlaff et al., 2019).

Identity of 3D7-Cas9 parasites was confirmed by whole-genome sequencing. Identity of transgenic parasites was confirmed by resistance to drug selectable markers, expression of fluorescently tagged proteins, and PCR. All parasite lines were tested periodically for Mycoplasma contamination.

All parasites were cultured in O+ human RBCs at 4% hematocrit in RPMI-1640 containing 25 mM HEPES, 50 mg/L hypoxanthine, 0.21% sodium bicarbonate, and 0.5% w/v Albumax II (Trager and Jensen, 1976). All parasite cultures were incubated on a shaker at 37°C in a gas mixture of 1% O2, 5% CO2, and 94% N2 as previously described. The smHA-tagged Pf3D7_0311800 (ATP Synthase F0 Subunit D) cell line was maintained under selection of 5 nm WR99210. Apicoplast targeting signal-GFP line was maintained under selection of 2.5 ug/ml of Blasticidin-hcl. 2xFKBP-GFP-K13 parasites were maintained under selection of 0.9 µm DSM1.

Parasites were routinely synchronized using sorbitol lysis. Briefly, parasite cultures were resuspended in 5% w/v D-sorbitol, resulting in the selective lysis of schizont-stage parasites (Lambros and Vanderberg, 1979).

For samples where parasites were arrested as schizonts using either trans-epoxysuccinyl-L-leucylamido(4guanidino)butane (E64) (Salmon et al., 2001) or compound 1 (C1) (Taylor et al., 2010), late schizont-stage cultures were treated with either 10 µm E64 for ~3 hr or 5 µm C1 for ~5 hr.

Plasmid generation and transfection

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For imaging of the mitochondria, a cell line where ATP-Synthase F0 Subunit D (Pf3D7_0311800) had a C-terminal spaghetti-monster HA tag was generated (Figure 5—figure supplement 2). To create the Pf3D7_0311800 smHA HDR plasmid, the 3D7_0311800 5′ and 3′ homology regions were PCR amplified from 3D7 genomic DNA with oligonucleotides oJDD4893/oJDD4894 and oJDD4891/oJDD4892, respectively. The two pieces were fused together using Sequence Overlap Extension PCR (SOE PCR) using oJDD4891/oJDD4894 and the piece was digested with NotI/XhoI and ligated with T4 ligase to generate pSAB55. To create the PF3D7_0311800 targeting guide RNA plasmid, oJDD4889/oJDD4890 were annealed, phosphorylated, and ligated into BpiI-digested pRR216 to generate pSAB81. All oligonucleotide sequences are shown in Table 1.

Table 1
Oligonucleotides for cloning and integration PCR.
Oligo/gBlock nameSequence (5′→3′)
JDD44TGGGGTGATGATAAAATGAAAG
JDD56ACACTTTATGCTTCCGGCTCGTATGTTGTG
JDD4889TATTGTCAAATCGTTACCTCTATG
JDD4890AAACCATAGAGGTAACGATTTGAC
JDD4891TAGgcggccgcGGTCCTACACCAATAAATATCA
JDD4892GTCTGATTCTTCCCATCaggccttccggaccgcggGGTCCCTTCATTGTAGACTTTTTATTATTGAAC
JDD4893GTCTACAATGAAGGGACCccgcggtccggaaggcctGATGGGAAGAATCAGACAAATGGT
JDD4894GATctcgagcAGcGGcAAtGAcTTcACgAATTCTCTTATTTCTTGTTTTTGCATTTCCT
SAB257CGGACCGAGAATTTATGTCCATTAACGTC
SAB471TGGTATTAATGGATGAAGACACACA
SAB472GTAATGGAATAGCTTTATATATGTACCTTCAT
SAB473TATGTGATCCATACATACCTGTTCAGAC

For transfection, 100 µg of pSAB55 plasmid was linearized with StuI and transfected into 3D7-Cas9, along with 100 µg of pSAB81. A day following transfection, parasites were treated with 5 nm WR99210 until 13 days, when resistant parasites were detected.

Ultrastructure expansion microscopy

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U-ExM was performed as previously described with minor modification (Bertiaux et al., 2021; Gambarotto et al., 2019; Liffner and Absalon, 2021). Then, 12 mm round Coverslips (Fisher, Cat# NC1129240) were treated with poly-d-lysine for 1 hr at 37°C, washed twice with MilliQ water, and placed in the wells of a 12-well plate. Parasite cultures were set to 0.5% hematocrit, and 1 mL of parasite culture was added to the well containing the coverslip for 15 min at 37°C. Culture supernatants were removed, and cultures were fixed with 1 mL of 4% v/v PFA in 1× PBS for 15 min at 37°C. For some experiments visualizing cytostomes, cultures were instead fixed in 4% v/v PFA + 0.01% v/v glutaraldehyde in 1× PBS. Following fixation, coverslips were washed three times with 37°C PBS before being treated with 1 mL of 1.4 % v/v formaldehyde/2% v/v acrylamide (FA/AA) in PBS. Samples were then incubated at 37°C overnight.

Monomer solution (19% w/w sodium acrylate [Sigma, Cat# 408220], 10% v/v acrylamide [Sigma, Cat# A4058, St. Louis, MO], 0.1% v/v N,N’-methylenebisacrylamide [Sigma, Cat# M1533] in PBS) was typically made the night before gelation and stored at −20°C overnight. Prior to gelation, FA/AA solution was removed from coverslips, and they were washed once in PBS. For gelation, 5 µL of 10% v/v tetraethylenediamine (TEMED; Thermo Fisher, Cat# 17919) and 5 µL of 10% w/v ammonium persulfate (APS; Thermo Fisher, Cat# 17874) were added to 90 µL of monomer solution and briefly vortexed. Subsequently, 35 µL was pipetted onto parafilm and coverslips were placed (cell side down) on top. Gels were incubated at 37°C for 30 min before being transferred to wells of a 6-well plate containing denaturation buffer (200 mM sodium dodecyl sulfate [SDS], 200 mM NaCl, 50 mM Tris, pH 9). Gels were incubated in denaturation buffer with shaking for 15 min, before separated gels were transferred to 1.5 mL tubes containing denaturation buffer. 1.5 mL tubes were incubated at 95°C for 90 min. Following denaturation, gels were transferred to 10 cm Petri dishes containing 25 mL of MilliQ water for the first round of expansion and placed onto a shaker for 30 min three times, changing water in between. Gels were subsequently shrunk with two 15 min washes in 25 mL of 1× PBS, before being transferred to 6-well plates for 30 min of blocking in 3% BSA-PBS at room temperature. After blocking, gels were incubated with primary antibodies, diluted in 3% BSA-PBS, overnight. After primary antibody incubation, gels were washed three times in 0.5% v/v PBS-Tween 20 for 10 min before incubation with secondary antibodies diluted in 1× PBS for 2.5 hr. Following secondary antibody incubation, gels were again washed three times in PBS-Tween 20, before being transferred back to 10 cm Petri dishes for re-expansion with three 30 min MilliQ water incubations.

Gels were either imaged immediately following re-expansion or stored in 0.2% w/v propyl gallate in MilliQ water until imaging. For gels stained with BODIPY TRc, the fully expanded gel was incubated overnight at room temperature in 0.2% w/v propyl gallate (Acos Organics, Cat# 131581000) solution containing BODIPY TRc (2 µM final concentration).

For parasites stained with MitoTracker Orange CMTMRos (Thermo Fisher, M7510), parasite cultures were resuspended in incomplete media (RPMI-1640 containing 25 mM HEPES, 50 mg/L hypoxanthine, and 0.21% sodium bicarbonate) containing 300 nM MitoTracker Orange CMTMRos. Parasite cultures were then stained with MitoTracker for 35 min while settling on poly-d-lysine-coated coverslips. From this point, the expansion protocol was followed as described above, with the exception that all steps when possible were carried out protecting the sample from light.

Cryopreservation and thawing of gels

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A proportion of gels imaged in this study were cryopreserved and subsequently thawed prior to imaging (Louvel et al., 2022). Gels were frozen either unstained, following the first round of expansion, or frozen stained, following the second round of expansion. To freeze, a portion of the expanded gel was placed into a 6-well dish and washed three times with 50% glycerol in MilliQ water for 30 min. Fresh glycerol was then added, and the gels were stored at –20°C for future use. To thaw unstained gels, the glycerol was replaced with MilliQ water and incubated at room temperature for 30 min. Gels were then washed and shrunk with three 20 min washes in 1× PBS at room temperature before proceeding with the antibody staining process normally. Stained gels were thawed with three washes in MilliQ water for 30 min before proceeding with imaging as normal.

Stains and antibodies

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A comprehensive list of all stains and antibodies used in this study, their working concentrations, and source(s) is provided in Table 2.

Table 2
Summary of all antibodies and stains used in this study.
Primary antibodiesAntibody speciesAntibody source (Cat#)Ab concentrationReference
Anti-alpha tubulin (Clone B-5-1-2)Mouse (IgG1)Thermo Fisher (32-2500)1:500
Anti-centrin (Clone 20H5)Mouse (IgG2a)Sigma-Aldrich (04-1624)1:200
Anti-Hscentrin1RabbitThermo Fisher (PA5-29986)1:500
Anti-polyE (IN105)RabbitAdipogen (AG-25B-0030-C050)1:500
Anti-ERD2 (MRA-1)RabbitBEI Resources MR41:2000Elmendorf and Haldar, 1993
Anti-HA (3F10)RatRoche (12158167001)1:50
Anti-GFPRabbitOriGene (TP401)1:2000
Anti-RAP1 (2.29)MouseEuropean Malaria Reagent Repository1:500Hall et al., 1983
Anti-RON4MouseGift from Alan Cowman1:100
Anti-RAMARabbitGift from Ross Coppel1:200Topolska et al., 2004
Anti-AMA1RabbitGift from Carole Long1:500
Anti-EBA175 (3D7)MouseGift from Alan Cowman1:500Sim et al., 2011
Anti-AldolaseRabbitAbcam (ab207494)1:2000
Anti-Histone H3RabbitAbcam (ab1791)1:1000
Anti-BIPRabbitGenerated by Dvorin Lab1:2000
Anti-GAP45RabbitGift from Julian Rayner1:2000Jones et al., 2009
Anti-IMC1gRabbitGenerated by Dvorin Lab1:1000Cepeda Diaz et al., 2023
Anti-MSP1 (1E1)RabbitGift from Anthony Holder1:250Blackman et al., 1994
Anti-hemoglobinRabbitThermo Fisher (PA5-102943)1:1000
Secondary antibodiesAntibody speciesAntibody sourceAntibody concentration
Anti-mouse IgG Alexa Fluor 488GoatThermo Fisher (A28175)1:500
Anti-mouse IgG Alexa Fluor 555GoatThermo Fisher (A21428)1:500
Anti-rabbit IgG Alexa Fluor 488GoatThermo Fisher (A11034)1:500
Anti-rabbit IgG Alexa Fluor 555GoatThermo Fisher (A21428)1:500
Anti-rat IgG Alexa Fluor 488GoatThermo Fisher (A11006)1:500
Anti-mouse IgG2a Alexa Fluor 488GoatThermo Fisher (A21131)1:500
Anti-mouse IgG1 Alexa Fluor 594GoatThermo Fisher (A21125)1:500
StainsStain source (Cat#)Stain concentration
NHS ester Alexa Fluor 405Thermo Fisher (A30000)1:250 (8 µM) in DMSO
BODIPY TR ceramideThermo Fisher (D7549)1:500 (2 µM)
SYTOX Deep RedThermo Fisher (S11381)1:1000 (1 µM) in DMSO

Image acquisition

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Immediately before imaging, a small slice of gel ~10 mm × ~10 mm was cut and mounted on an imaging dish (35 mm Cellvis coverslip bottomed dishes NC0409658, Fisher Scientific) coated with poly-d lysine. The side of the gel containing sample is placed face down on the coverslip and a few drops of ddH20 are added after mounting to prevent gel shrinkage due to dehydration during imaging. All images presented in this study were taken using either a Zeiss LSM800 AxioObserver with an Airyscan detector, or a Zeiss LSM900 AxioObserver with an Airyscan 2 detector. Imaging on both microscopes was conducted using a ×63 Plan-Apochromat objective lens with a numerical aperture of 1.4. All images were acquired as Z-stacks that had an XY pixel size of 0.035 µm and a Z-slice size of 0.13 µm.

Image analysis

Image processing and presentation

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All images were Airyscan processed using 3D processing at moderate filter strength on ZEN Blue Version 3.1 (Zeiss, Oberkochen, Germany).

The majority of images presented in this study are presented as maximum-intensity z-projections, but those that contain BODIPY TRc are presented as average intensity projections for viewing and interpretation purposes. For images containing NHS ester, the gamma value of this channel was set to 0.45 rather than 1 as this allowed discernment of a greater number of parasite structures. It should be noted that because of this the fluorescence intensities shown in the NHS ester channel are not linear.

3D rendering

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3D renderings of micronemal proteins AMA1 and EBA175 were produced using the 3D analysis package on ZEN Blue version 3.5.

Measurement of interpolar spindles and SPMTs

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All length measurements reported in this study were obtained using the ‘Measure 3D distance’ function of ZEN Blue version 3.1. The length of interpolar spindle microtubules and SPMTs was determined as the 3D distance between the start and end points of continuously stained stretches of anti-tubulin staining. Interpolar microtubules were defined as those whose staining appeared to contact both CPs as defined by NHS ester, while non-interpolar microtubules were those that did not meet these criteria. SPMTs were only measured in C1-arrested schizonts that had visibly completed segmentation based on the basal complex as visualized by NHS ester. Any microtubule that did not appear connected to the APR, or extend toward the basal complex was excluded from the analysis. Cell diameter was defined as the greatest XY distance on any z-slice between two points of the parasite as defined by NHS ester staining. Merozoite length was defined as the 3D distance between the center of the APRs and basal complex as defined by NHS ester staining.

Apicoplast and mitochondria area analysis

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All area measurements presented were obtained using the ‘Area’ function on ZEN Blue version 3.1. Images were presented as a maximum-intensity projection before free hand outlining the apicoplast or the mitochondria in each image. The sum of all fragments was then calculated to find the total area of the organelles per cell.

Mitochondria residual body analysis

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In C1-arrested schizonts, the proportion of total mitochondria staining found in the residual body was calculated as follows. Using ZEN Blue version 3.1, a maximum-intensity projection of the entire cell was generated and based on the NHS staining, the entire parasite was defined as the region of interest. Signal intensity of the channel staining the mitochondria was calculated inside the full cell, which was defined as total mitochondria fluorescence. Subsequently, the residual body was defined the area within the parasite vacuole, but external to all merozoite plasma membranes, as based on BODIPY TRc staining. A second maximum-intensity projection of this subsection of the schizont was made, the residual body was defined as the region of interest, and this signal intensity inside this region of interest was defined as residual body mitochondria fluorescence.

To determine residual body mitochondria fluorescence (RB) as % total mitochondria fluorescence (total), the following equation was used:

(RB fluorescenceTotal fluorescence)×100

In each of the parasites included in this analysis, the number of merozoites was defined as the number of distinct nuclei as determined by SYTOX staining. To determine residual body mitochondria fluorescence as % of one merozoite, the following equation was used:

(RB fluorescence ÷ Total fluorescence-RB fluorescenceNumber of merozoites)×100

Cells where no mitochondria fluorescence was visible inside the residual body, while visible inside merozoites, were defined as having no residual body mitochondria. An attempt was made to do a similar analysis on apicoplast stained cells, but no visible apicoplast staining was ever observed in the residual body.

Statistical analysis

Estimation of actual distance from expanded samples

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Expansion factors for 43 gels used in this study were determined as follows. Gels were assumed to have an initial diameter of 12 mm as they are formed on a 12-mm-diameter coverslip. Gels were subsequently measured following expansion to the nearest whole millimeter, and the expansion factor was defined as the expanded gel diameter divided by the initial gel diameter (12 mm). Gels whose edges were damaged or malformed, and therefore their diameters could not be actually measured, were excluded. Gels in this study had a median expanded diameter of 51 mm, which corresponds to a median expansion factor of 4.25 (Figure 1—figure supplement 1).

Generation of graphs and statistical analysis

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All graphs presented in this study were generated using GraphPad PRISM 9. All error bars in this study represent standard deviation. Differences between samples analyzed by ANOVA was determined as difference where the p-value was <0.05. For scatterplots, slopes were considered significantly non-zero when the p-value was <0.05.

Data accessibility

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Results in this study are underpinned by 647 3D Airyscan images of U-ExM parasites at multiple lifecycle stages with multiple combinations of stains. All images are publicly available through the following data repository: https://doi.org/10.5061/dryad.934mw6mp4.

Data availability

All images underlying this publication are publically available through Dryad at the following link https://doi.org/10.5061/dryad.9s4mw6mp4.

The following data sets were generated
    1. Benjamin L
    2. Diaz AKC
    3. Blauwkamp J
    4. Anaguano D
    5. Frölich S
    6. Muralidharan V
    7. Wilson DW
    8. Dvorin J
    9. Absalon S
    (2023) Dryad Digital Repository
    Data from: Atlas of Plasmodium falciparum intraerythrocytic development using expansion microscopy.
    https://doi.org/10.5061/dryad.9s4mw6mp4

References

    1. Lambros C
    2. Vanderberg JP
    (1979)
    Synchronization of Plasmodium falciparum erythrocytic stages in culture
    The Journal of Parasitology 65:418–420.

Peer review

Reviewer #1 (Public Review):

This paper consists of a comprehensive analysis of the malaria parasite Plasmodium falciparum during its development in erythrocytes, using expansion microscopy. The authors used general dyes to stain membranes or proteins and a set of specific markers to label diverse cellular structures of the parasite, with a particular focus on the centriolar plaque.

This is by nature a purely descriptive study, providing remarkable images with great details on subcellular structures such as the centriolar plaque, the basal complex, the cytostome and rhoptries. The work is extremely well performed and the images are beautiful. This study confirms a number of previous observations and illustrates the strength of expansion microscopy, an affordable and adaptable sample preparation method that will undoubtedly become standard in the field.

This study provides a valuable resource that can serve as a reference dataset for the analysis of P. falciparum and other apicomplexan parasites.

https://doi.org/10.7554/eLife.88088.3.sa1

Reviewer #2 (Public Review):

In this work the authors describe the shape and interconnectedness of intracellular structures of malaria blood stage parasites by taking advantage of expansion microscopy. Compared to previous microscopy work with these parasites, the strength of this paper lies in the increased resolution and the fact that the NHE ester highlights protein densities. Together with the BodipyC membrane staining, this results in data that is somewhere in between EM and standard fluorescence microscopy: it has higher resolution than standard fluorescence microscopy and provides some points of reference of different cellular structures due to the NHE ester/BodipyC.

This study makes many interesting and useful observations and although it is somewhat "old school descriptory" in its presentation, researchers working in many different areas will find something of interest here. This ranges from mitosis, to organisation and distribution of major cellular structures, endocytosis and invasion, overall providing a rich and interesting resource. The results section is long but by taking the space to explain everything in detail, it has the advantage that it clearly transpires how things were done and on how many cells a conclusion is based on. Further the authors often also included a brief interpretation of their findings with a very open assessment what it does and what it does not show, highlighting interesting questions left by the data.

Overall this is a very nice and useful paper that will be of interest to many, particularly those working on nuclear division, cytokinesis, endocytosis or invasion in malaria parasites. The spatiotemporal arrangement and interconnection of subcellular structures will also give a framework for specific functional studies.

https://doi.org/10.7554/eLife.88088.3.sa2

Reviewer #3 (Public Review):

In their study the authors analyze the localization of multiple organelles and subcellular structure of blood stage malaria parasites with unprecedented detail. They use a 3D super-resolution imaging technique that has gained popularity in the protozoan field, ultrastructure expansion microscopy. Building on markers and labels established in the field they generate an appealing collection of images for all stages of the intraerythrocytic developmental stages of asexual blood stage parasites with some focus on nuclear division and cell segmentation stages.

The authors have made a very good effort to address all the comments raised by the reviewers providing more clarity to the manuscript and appropriate interpretations of their results. Particularly the sharing of their image data in the Dryad repository adds significant value to their work.

https://doi.org/10.7554/eLife.88088.3.sa3

Author response

The following is the authors’ response to the original reviews.

We greatly appreciate the overwhelmingly positive summaries from all three reviewers and the eLife editorial team. All reviewers provided extremely detailed feedback regarding the initially submitted manuscript, we appreciate their efforts in helping us improve this manuscript. Below, are listed each of the specific comments made by the reviewers, and our responses to them in a point-by-point format.

The only notable change made to the manuscript that was not in response to comments from a reviewer was regarding nomenclature of the structure that we had previously called the nuclear microtubule organising centre (MTOC). We had used the term MTOC to describe the entire structure, which spans the nuclear envelope and comprises an intranuclear portion and cytoplasmic extensions. Given recent evidence, including findings from this study, it is possible that both the intranuclear region and cytoplasmic extensions both have microtubule nucleating capacity, and therefore both meet the definition of an MTOC. To disambiguate this, we now refer to the overall structure as the centriolar plaque (CP), consistent with previous literature. The intranuclear portion of the CP will be referred to as the inner CP, while the cytoplasmic portion will be referred to as the outer CP.

Reviewer #1 (Recommendations For The Authors):

1. In the first part of the result section, a paragraph on sample processing for U-ExM could be added, with reference to Fig 1b.

The following section has been added to the first paragraph of the results “…In this study all parasites were fixed in 4% paraformaldehyde (PFA), unless otherwise stated, and anchored overnight at 37 °C before gelation, denaturation at 95 °C and expansion. Expanded gels were measured, before shrinking in PBS, antibody staining, washing, re-expansion, and imaging (Figure 1b). Parasites were harvested at multiple time points during the intraerythrocytic asexual stage and imaged using Airyscan2 super-resolution microscopy, providing high-resolution three-dimensional imaging data (Figure 1c). A full summary of all target-specific stains used in this study can be found in Figure 1d.”

1. The order of the figures could be changed for more consistency. For example, fig 2b is cited before 2a.

An earlier reference to figure 2a was added to rectify this discrepancy.

1. In Fig 2b it is difficult to distinguish the blue (nuclear) and green (plasma membrane) lines.x

The thickness of these lines has been doubled.

1. It is unclear what the authors want to show in Fig 2a.

The intention of this figure, as with panel a of the majority of the organelle-specific figures in this manuscript, is simply to show what the target protein/structure looks like across intraerythrocytic development.

1. Lines 154-155, the numbers of MTOC observed do not match those in Supplt Fig2c.

This discrepancy has been addressed, the numbers in Supplementary Figure 2c were accurate so the text has been changed to reflect this.

1. Line 188: the authors should explain the principle of C1 treatment.

The following explanation of C1 treatment has been provided:

“To ensure imaged parasites were fully segmented, we arrested parasite development by adding the reversible protein kinase G inhibitor Compound 1 (C1). This inhibitor arrests parasite maturation after the completion of segmentation but before egress. When C1 is washed out, parasites egress and invade normally, ensuring that observations made in C1-arrested parasites are physiologically relevant and not a developmental artefact due to arrest.”

1. Lines 195-204: this part is rather difficult to follow as analysis of the basal complex is detailed later in the manuscript. The authors refer to Fig4 before describing Fig3.

This has been clarified in the text

1. Lines 225 and 227, the authors cite Supplt Fig 2b about the Golgi, but probably meant Supplt Fig 4? In Supplt Fig 4, the authors could provide magnification in insets to better illustrate the Golgi-MTOC association.

This should have been a reference to Supplementary Figure 2e instead of 2b, which has now been changed. In Supplementary Figure 4, zooms into a single region of Golgi have been provided to more clearly show its MTOC association.

1. Supplt Fig8 is wrong (duplication of Supplt Fig6).

We apologise for this mistake, the correct figure is now present in Supplementary Figure 8.

1. Line 346: smV5 should be defined, and generation of the parasites should be described in the methods.

This has now been defined, but we have not described the generation of the parasites, as this was performed in a previous study that we have referenced.

1. Lines 361-362: "By the time the basal complex reaches its maximum diameter..." This sentence is not very clear, the authors could explain more precisely the sequence of events, indicating that the basal complex starts moving in the basal direction, as clearly illustrated in Fig 4a.

This has been prefaced with the following sentence “…As the parasite undergoes segmentation, the basal complex expands and starts moving in the basal direction.”

1. Supplt Fig6 comes after Supplt Fig9 in the narrative, and therefore could be placed after.

Supplementary Figure 6 and 9 follow the order in which they are referred to in the text.

1. Line 538: Supplt Fig9e instead of 9d.

This has been fixed.

1. Line 581: does the PFA-glutaraldehyde fixation allows visualizing other structures in addition to cytostome bulbs?

While PFA-glutaraldehyde fixation allows visualisation of cytostome bulbs, to date we have not observed any other structure that stains/preserves better using NHS Ester or BODIPY Ceramide in PFA-glutaraldehyde fixed parasites. As a general trend, all structures other than cytostomes become somewhat more difficult to identify using NHS Ester or BODIPY Ceramide in PFA-glutaraldehyde fixed samples due to the local contrast with the red blood cell cytoplasm. It seems likely that this is just due to the preservation of RBC cytoplasm, and would be expected from any fixation method that doesn’t result in RBC lysis, rather than anything unique to glutaraldehyde.

1. Line 652-653: It is unclear how the authors can hypothesize that rhoptries form de novo rather than splitting based on their observations.

This not something we can say with certainty, we have however, introduced the following paragraph to qualify our claims: “Overall, we present three main observations suggesting that rhoptry pairs undergo sequential de novo biogenesis rather than dividing from a single precursor rhoptry. First, the tight correlation between rhoptry and MTOC cytoplasmic extension number suggests that either rhoptry division happens so fast that transition states are not observable with these methods or that each rhoptry forms de novo and such transition states do not exist. Second, the heterogeneity in rhoptry size throughout schizogony favors a model of de novo biogenesis given that it would be unusual for a single rhoptry to divide into two rhoptries of different sizes. Lastly, well-documented heterogeneity in rhoptry density suggests that, at least during early segmentation, rhoptries have different compositions. Heterogeneity in rhoptry contents would be difficult to achieve so quickly after biogenesis if they formed through fission of a precursor rhoptry.”

1. Line 769: is expansion microscopy sample preparation compatible with FISH?

Yes, there are publications of expansion being done with both MERFISH and FISH. Though it has not yet been applied to plasmodium. See examples: Wang, Guiping, Jeffrey R. Moffitt, and Xiaowei Zhuang. "Multiplexed imaging of high-density libraries of RNAs with MERFISH and expansion microscopy." Scientific reports 8.1 (2018): 4847. And Chen, Fei, et al. "Nanoscale imaging of RNA with expansion microscopy." Nature methods 13.8 (2016): 679-684.

1. In the methods, the authors could provide details on the gel mounting step for imaging This is particularly important since this paper will likely serve as a reference standard for expansion microscopy in the field. Also, illustration that cryopreservation of gels does not modify the quality of the images would be useful.

The following section has been added to our “image acquisition” paragraph: “Immediately before imaging, a small slice of gel ~10mm x ~10mm was cut and mounted on an imaging dish (35mm Cellvis coverslip bottomed dishes NC0409658 - FisherScientific) coated with Poly-D lysine. The side of the gel containing sample is placed face down on the coverslip and a few drops of ddH20 are added after mounting to prevent gel shrinkage due to dehydration during imaging.”

We have decided not to illustrate that cryopreservation does not alter gel quality, as this is something that is already covered in the study that first cryopreserved gels, which is referenced in our methods section.

Reviewer #2 (Recommendations For The Authors):

1. Advantages and limitations of the expansion method are generally well discussed. The only matter in that respect that I was wondering is if expansion can always be assumed to be linear for all components of a cell. The hemozoin crystal does not expand (maybe not surprisingly), but could there also be other cellular structures that on a smaller scale separate or expand at a different rate than others? Is there any data on this from other organisms? I am raising this here not as a criticism of this work but if known to occur, it might need mentioning somewhere to alert the reader to it, particularly in regards to the many measurements in the paper (see also point 4). This might be a further factor contributing to the finding that the IMC and PPM could not be resolved.

This is an excellent point and, to our knowledge, one that is currently still under investigation in the field. It is well-documented that expansion protocols need to be customized to each cell type and tissue they are applied to. Each solution used for fixation and anchoring as well as timing and temperature of denaturation can affect the expansion factor achieved as well as how isotropic/anisotropic the expanded structures turn out. However, we do not know of any examples where isotropic expansion was achieved for everything but an organelle or component of the cell. It is our impression that if the cell seems to have attained isotropic expansion, this is assumed to also be the case for the subcellular structures within it. Nonetheless, we think it remains a possibility to be considered specially as more structures are characterized using these methods. In the case of our IMC/PPM findings, when we performed calculations taking into account our experimental expansion factor as well as antibody effects, it was clear that the resolution of our microscope was not enough to resolve the two structures using our current labelling methods. So, we suspect most of the effect is driven by that. However, this still needs to be validated by attempting to resolve the two structures though alternative labelling and imaging methods.

1. I understand that many things described in the results part are interconnected but still the level of hopping around between different figures/supp figures is considerable (see also point 6 on synchronicity of Figure parts). I do not have a simple fix, but maybe the authors could check if they could come up with a way to streamline parts of their results into a somewhat more reader friendly order.

This has been a problem we encountered from the beginning and, after trying multiple presentations of the results and discussion, we realized they all have drawbacks. We eventually settled on this presentation as the “least confusing”. We agree, however, that the figure references and order could be better streamlined and have addressed this to the best of our ability.

1. Are the authors sure the ER expands well and the BIP signal (Fig. S5) gives a signal reflecting the true shape of the ER? The signal in younger parasites seems rather extensive compared to what the ER (in my experience) typically looks like in these stages in live parasites.

While there may be a discrepancy between how the presumably dynamic ER appears in live cells, and how it appears using BiP staining, we think it is unlikely this is a product of expansion. Additionally, if there were to be an artefactual change in the ER, it would be likely under-expansion rather than over-expansion, which to our knowledge has not been reported. In our opinion, the BiP staining we observe is comparable between unexpanded and expanded samples. We have included comparative images in Author response image 1 with DNA in cyan and BiP in yellow, unexpanded (left) and expanded (right) using the same microscope and BiP antibody.

Author response image 1

1. It is nice to have measurements of the apicoplast and mitochondria, but given their size, this could also have been done in unexpanded, ideally live parasites, avoiding expansion and fixing artifacts. While the expansion has many nice features, measuring area of large structures may not be one where it is strictly needed. I am not saying this is not useful information, but maybe a note could be added to the manuscript that the conclusions on mitochondria and apicoplast area and division might be worth confirming in live parasites. A brief mention on similarities and differences to previous work analysing the shape and multiplication of these organelles through blood stage development (van Dooren et al MolMicrobiol2005) might also be useful.

We agree with the reviewer that previous studies such as van Dooren et al. (2005) demonstrate that it is possible to track apicoplast and mitochondrial growth without expansion and share the opinion that live parasites are better for these measurements. Expansion only provides an advantage when more organelle-level resolution is needed. For example, in studying the association between these organelles and the MTOC or visualizing other branch-specific interactions.

1. I could not find the Supp Fig. 8 on the IMC, the current Supp Fig. 8 is a duplication of Supp Fig. 6

This has been addressed, Supplementary Figure 8 now refers to the IMC.

1. Figure order is not very synchronous with the text: Fig. 2a is mentioned after Fig. 2b, Fig. 4b is mentioned first for Fig. 4 (Fig. 4a is not by itself mentioned) and before Fig. 3 is mentioned; Fig. 3b is before Fig. 3a.

We have done our best to fix these discrepancies, but concede that we have not found a way to order these sections that doesn’t lead to some confusion.

1. Fig. S2a, The label "Centrin" on left image is difficult to read

We have increased the font size and changed colour slightly in the hope it is leigible.

1. In Fig. 2a, the centrin foci are very focal and difficult to see in these images, particularly when printed out but also on screen. To a lesser extent this is also the case for CINCH in Fig. 4a (particularly when printed; when zoomed-in on screen, the signal is well visible). This issue of difficulties in seeing the fluorescence signal of some markers, particularly when printed out, applies also to other images of the paper.

In the images of full size parasites, this is an issue that we cannot easily overcome as the fluorescent channels are already at maximum brightness without overexposure. To try and address this, we have provided zooms that we hope will more clearly show the fluorescence in these panels.

1. Expand "C1" in line 188 (first use).

This has been addressed in response to a previous comment.

1. Line 227; does Supp Fig. 2b really show Golgi- cytoplasmic MTOC association?

We have rephrased the wording of this section to clarify that we are observing proximity and not necessarily a physical tethering, however it is worth nothing that this was an accidental reference to Supplementary Figure 2b, and should’ve been Supplementary Figure 2e.

1. Line 230, in segmented schizonts the Golgi was considered to be at the apical end. It might be more precise to call its location to be close to the nucleus on the side facing the apical end of the parasite. It seems to me it often tends to be closer to the nucleus (in line with its proximity to the ER, see also point 13).

We have added more detail to this description clarifying that despite being at the apical end, the Golgi is closer to the nucleus.

1. Supp Fig. S5: Is the top cell indeed a ring? In the second cell there seem to be two nuclei, I assume this is a double infection (please indicate this in the legend or use images of a single infection).

In our opinion, the top cell in Supplementary Figure 5 is a ring. This is based on its size and its lack of an observable food vacuole (an area that lacks NHS ester staining). We typically showed images of ameoboid rings to avoid this ambiguity, but we think this parasite is a ring nonetheless. For the second image, this parasite is not doubly infected, as both DNA masses are actually contained within the same dumbbell shaped nuclear envelope. This parasite is likely undergoing its first anaphase (or the Plasmodium equivalent of anaphase) and will likely soon undergo its first nuclear division to separate these two DNA masses into individual nuclei.

1. Line 244: I would not call the Golgi a part of the apical cluster of organelles. All secretory cargo originates from the ER-Golgi-transGolgi axis in a directional manner and this axis is connected to the nucleus by the perinuclear ER. If seen from a secretory pathway centred view, it is the other way around and you could call the apical organelles part of the nuclear periphery which would be equally non-ideal.

Everything is close together in such a small cell. The secretory pathway likely is arranged in a serial manner starting from the perinuclear region to the transGolgi where cargo is sorted into vesicles for different destinations of which one is for the delivery of material to the apical organelles. The proposition that the Golgi is part of the apical cluster therefore somehow feels wrong, as the Golgi can still be considered to be upstream of the transGolgi before apical cargo branches off from other cargo destined for other destinationsWe agree with the reviewer that claiming a functional association between the Golgi and the apical organelles would be odd and we by no means meant to imply such functional grouping. Our intent was to confirm observations previously made about Golgi positioning by electron microscopy studies such as Bannister et al. (2000) at a larger spatial and temporal scale. These studies make the observation that the Golgi is spatially associated with the rhoptries at the apical end of the parasites. Logically, the Golgi is tied to the apical organelles through the secretory pathway as the reviewer suggests, but we claim no further relationship beyond that of organelle biogenesis. We have made modifications to the text to clarify these points.

1. Lines 300 - 308 (and thereafter): I assume these were also expanded parasites and the microtubule length is given after correction for expansion. I would recommend to indicate in line 274 (when first explaining the expansion factor) that all following measurements in the text represent corrected measures or, if this is not always the case, indicate on each occasion. Is the expansion factor accurate and homogenous enough to draw firm conclusions (see also point 1)? Could it be a reason for the variation seen with SPMTs? Could a cellular reference be used as a surrogate to account for cell specific expansion or would you assume that cellular substructure specific expansion differences exist and prevent this?

This is correct, the reported number is the number corrected for expansion factor, and the corresponding graphs with uncorrected data are present in the Supplementary Figures. We have clarified this in the text.Uneven expansion can be caused when certain organelles/structures do not properly denature. Given that out protocol denatures using highly concentrated SDS at 95 °C for 90 minutes, we do not anticipate that any subcellular compartments would expand significantly differently. In this study our expansion factors varied from ~4.1-4.7 across all gels, and for our corrected values we used the median expansion factor of 4.25. If we are interpreting the length of an interpolar spindle as 20 µm for example, the value would be corrected value would be 4.7 µm when divided by the median expansion factor, 4.9 µm when divided by the lowest, and 4.2 µm when divided by the highest. These values fall well within the measurement error, and so we expect that these small deviations in expansion factor between gels have a fairly minimal influence on variation in microtubule lengths.

1. Line 353: this is non-essential, but a 3D view of the broken basal ring might better illustrate the 2 semicircles

We have added the following panel to Supplementary Figure 3 to illustrate this more clearly:

Author response image 2

1. The way the figure legends are shaped, it often seems only panel (a) is from expansion microscopy while the microscopy images in the other parts of the figures have no information on the method used. I assume all images are from expansion microscopy, maybe this could be clarified by placing this statement in a position of the legend that makes it clear it is for all images in a figure.

This has been clarified in the figure legends.

1. Fig. 8b, is it clear that internal RON4 is not below or above? Consider showing a 3D representation or side view of these max projections.

If in these images, we imagine we are looking at the ‘top’ of the rhoptries, our feeling is that the RON4 signal is on the ‘bottom’, at the part closest to the apical polar ring. We tried projecting this, however, but the images were not particularly due to spherical aberrations. Because of this, we have refrained from commenting on the RON4 location relative to the rhoptry bulb prior to elongation.

1. Line 684 "...distribution or RON4": replace or with of. The information of the next sentence is partly redundant, consider adding it in brackets.

This has been addressed.

1. Fig. 9a the EBA175 signal is not very prominent and a bit noisy, are the authors confident this is indeed showing only EBA175 or is there also some background?-AK

We agree with the reviewer that the EBA175 antibody shows a significant amount of background fluorescence, specially in the food vacuole area. However, we think the puncta corresponding to micronemal EBA175 can be clearly distinguished from background.

1. Fig. 9b, the long appearance of the micronemes in the z-dimension likely is due to axial stretch (due to point spread function in z and refractive index mismatch), in reality they probably are more spherical. It might be worth mentioning somewhere that this likely is not how these organelles are really shaped in that dimension (spherical fluorescent beads could give an estimation of that effect in the microscopy setup used).

After recently acquiring a water-immersion objective lens for comparison, it is clear that the transition from oil to hydrogel causes a degree of spherical aberration in the Z-plane, which in this instance causes the micronemes to be more oblong. As we make no conclusions based on the shape of the micronemes, however, we don’t think this is a significant consideration. This is an assumption that should be made when looking at any image whose resolution is not equal in all 3-dimensions. We also note that the more spherical shape of micronemes can be inferred from the max intensity projections in Figure 9c.

1. Fig. 9b, the authors mention in the text that there is NHS ester signal that overlaps with the fluorescence signal, can occasions of this be indicated in the figure?

Figure 9b was already quite busy, so we instead added the following extra panel to this figure that more clearly shows the NHS punctae we thought may have been micronemes:

Author response image 3

1. Fig. 9, line 695, the authors write that the EBA puncta were the same size as AMA1 puncta. To me it seems the AMA1 areas are larger than the EBA foci, is their size indeed similar? Was this measured?

Since we did not conduct any measurements and doing so robustly would be difficult given the density of the puncta, we have decided to remove our comment on the relative size of the puncta.

1. Materials and methods: Remove "to" in line 871; explain bicarb and incomplete medium in line 885 (non-malaria researchers will not understand what is meant here); line 911 and start of 912 seem somewhat redundant

This has been addressed.

1. is there more information on what the Airyscan processing at moderate filter level does? The background of the images seems to have an intensity of 0 which in standard microscopy images should be avoided (see for instance doi:10.1242/jcs.03433) similar to the general standard of avoiding entirely white backgrounds on Western blots. I understand that some background subtraction processes will legitimately result in this but then it would be nice to know a bit better what happened to the original image.

We have taken the following excerpt from a publication on Airyscan to help clarify:

"Airyscan processing consists of deconvolution and pixel reassignment, which yield an image with higher resolution and reduced noise. This can be a contributor to the low background in some channels. The level of filtering is the processing strength, with higher filtering giving higher resolution but increased chances of artefacts. More information about the principles behind Airyscan processing can be found in the following two publications, though details on the algorithm itself seem to be proprietary: Huff, Joseph. "The Airyscan detector from ZEISS: confocal imaging with improved signal-to-noise ratio and super-resolution." (2015): i-ii. AND Wu, Xufeng, and John A. Hammer. "Zeiss airyscan: Optimizing usage for fast, gentle, super-resolution imaging." Confocal Microscopy: Methods and Protocols. New York, NY: Springer US, 2021. 111-130."

We cannot find any further information about the specifics of Airyscan filtering, however, the moderate filter that we used is the default setting. This information was included just for clarity, rather than something we determined by comparison to other filtering settings.

In regards to the background, the majority of some images having an intensity value of 0 is partially out of our control. For all NHS Ester images, the black point of the images was 0 so areas that lack signal (white in the case of NHS Ester) truly had no signal detected for those pixels. While we appreciate that never altering the black point of images displays 100% of the data in the image, images with any significant background can become impossibly difficult to interpret. We have done our best to try and present images where the black point is modified to remove background for ease of interpretation by the readers only.

Reviewer #3 (Public Review):

1. Most importantly, in order to justify the authors claim to provide an "Atlas", I want to strongly suggest they share their raw 3D-imaging data (at least of the main figures) in a data repository. This would allow the readers to browse their structure of interest in 3D and significantly improve the impact of their study in the malaria cell biology field.

We agree completely that the potential impact of this study is magnified by public sharing of the data. The reason that this was not done at the time of submission is that most public repositories do not allow continued deposition of data, and so new images included in response to reviewers comments would’ve been separated from the initial submission, which we saw as needlessly complicated. All 647 images that underpin the results discussed in this manuscript are now publicly available in Dryad (https://doi.org/10.5061/dryad.9s4mw6mp4)

1. The organization of the manuscript can be improved. Aside some obvious modifications as citing the figures in the correct order (see also further comments and recommendations), I would maybe suggest one subsection and one figure per analyzed cellular structure/organelle (i.e. 13 sections). This would in my opinion improve readability and facilitate "browsing the atlas".

This is actually how we had originally formatted this manuscript, but this structure made discussing inter-connected organelles, such as the IMC and basal complex, impossibly difficult to navigate. We have done our best to make the manuscript flow better, but have not come up with any way to greatly restructure the manuscript so to increase its readability.

1. Considering the importance of reliability of the U-ExM protocol for this study the authors should provide some validation for the isotropic expansion of the sample e.g. by measuring one well defined cellular structure.

The protocol we used comes from the Bertiaux et al., 2021 PLoS Biology study. In this study they show isotropic expansion of blood-stage parasites.

1. In the absence of time-resolved data and more in-depth mechanistic analysis the authors must down tone some of their conclusions specifically around mitochondrial membrane potential, subpellicular microtubule depolymerization, and kinetics of the basal complex.

Our conclusions regarding mitochondrial membrane potential and basal complex kinetics have been dampened. We have not, however, changed our wording around microtubule depolymerisation. Partial depolymerisation of microtubules during fixation is a known phenomenon in Plasmodium, and in our opinion, our explanation of this offers a hypothesis that is balanced with respective to evidence: “we hypothesise that most SPMTs measured in our C1-treated schizonts had partially depolymerised. P. falciparum microtubules are known to rapidly depolymerise during fixation10,29. It is unclear, however, why this depolymerization was observed most often in C1-arrested parasites. Thus, we cannot determine whether these shorter microtubules are a by-product of drug-induced arrest or a biologically relevant native state that occurs at the end of segmentation.”

1. The observation that the centriolar plaque extensions remains consistently tethered to the plasma membrane is of high significance. To more convincingly demonstrate this point, it would be very helpful to show one zoomed-in side view of nucleus with a mitotic spindle were both centriolar plaques are in contact with the plasma membrane.

We of course agree that this is one of our most important observations, but in our opinion this is already demonstrated in Figure 2b. The third panel from the right shows a mitotic spindle and has the location of the cytoplasmic extensions, nuclear envelope and parasite plasma membranes annotated.

1. Please verify the consistent use of the term trophozoite and schizont. In Fig. 1c a parasite with two nuclei, likely in the process of karyofission is designated as trophozoite, which contrasts with the mononucleated trophozoite shown in Fig. 1a. The reviewer is aware of the more "classical" description of the schizont as parasite with more than 2 nuclei, but based on the authors advanced knowledge of cell cycle progression and mitosis I would encourage them to make a clear distinction between parasites that have entered mitotic stages and pre-mitotic parasites (e.g. by applying the term schizont, and trophozoite, respectively).

For this study, we have interpreted any parasite having three or more nuclei as being a schizont. We are aware this morphological interpretation is not universally held and indeed suboptimal for studying some aspects of parasite development, but all definitions of a schizont have some drawbacks. Whether a parasite has entered mitosis or not is obviously a hugely significant event in the context of cell biology, but in a mononucleated parasite this could only be determined using immunofluorescence microscopy with cell cycle or DNA replication markers.

1. Aldolase does not localize diffusely in the cytoplasm in schizont stages as in contrast to earlier stage. The authors should comment on that.

We are unclear if this is an interpretation of the images in supplementary figure 1, or inferred from other studies. If this is an interpretation of the images in Supplementary Figure 1, we do not agree that the images show a significant change in the localisation of aldolase. It is possible that this difference in interpretation comes from the strong punctate signal observed more readily in the schizont images. This is the strong background signal in or around the food vacuole we mention in the text. These punctae are significantly brighter than the cytosolic aldolase signal, making it difficult to see them on the aldolase only channel, but aldolase signal can clearly be seen in the cytoplasm on the merge images.

1. Line 79. Uranyl acetate is just one of the contrasting agents used in electron microscopy. The authors might reformulate this statement. Possibly this would also be a good opportunity to briefly mention that electron density measured in EM and protein-density labeled by NHS-Ester can be similar but are not equivalent.

We have expanded on this in the text.

1. The authors claim that they investigate the association between the MTOC and the APR (line 194), but strictly speaking only look at subpellicular microtubules and an associated protein density. The argument that there is a "NHS ester-dense focus" (line 210) without actual APR marker is not quite convincing enough to definitively designate this as the APR.

While an APR marker would of course be very useful, there are currently no published examples of APR markers in blood-stage parasites. We therefore think that the timing of appearance, location, and staining density are sufficient for identifying this structure as the APR, as it has previously been designated through EM studies. We have nonetheless softened our language around APR-related observations.

1. Line 226: The authors should also discuss the organization of the Golgi in early schizonts (Fig. S4). (not only 2 nuclei and segmenter stages).

We did not mean to imply that all 22 parasites had only 2 nuclei, but instead that they had 2 or more nuclei. Therefore, early schizonts are included in this analysis, with Golgi closely associated with all their MTOCs.

1. Line 242: To the knowledge of the reviewer the nuclear pore complexes, although clustered in merozoites and ring stages, don't particularly "define the apical end of the parasite".

The MTOC is surrounded by NPCs, which because of the location of the MTOC end up being near the forming apical end of the merozoite, but we have removed this as it was needlessly confusing.

1. Supplementary Figure 8 is missing (it's a repetition of Fig. S6).

This has been addressed.

1. Line 253: asexual blood stage parasites have two classes of MTs. Other stages can have more.

This has been clarified.

1. Fig. 3f: Please comment how much of these observations of "only one" SPMT could result from suboptimal resolution (e.g. in z-direction) or labeling. Otherwise use line profiles to argue that you can always safely distinguish SPMT pairs.

In the small number of electron tomograms of merozoites where the subpellicular microtubules have been rendered, they have been seen to have 2 or 3 SPMTs. Despite this, we don’t think it is likely that the single SPMT merozoites observed in this study are caused by a resolution limitation. SPMTs were measured in 3D, rather than from projections, and any schizont where the SPMTs were pointing towards the objective lens, elongating the parasite in Z, were not imaged. Additionally, our number of merozoites with a single SPMT correspond with the same data collected in the Bertiaux et al., 2021 PLoS Biology study. We cannot rule this out as a possibility, as sometimes SPMTs cross over each other in three-dimensions, and at these intersection points they cannot be individually resolved. We, however, think it is very unlikely that two SPMTs would be so close that they can never be resolved across any part of their length.

1. Lines 302ff: the claim that variability in SPMT size must be a consequence of depolymerzation is unfounded. The dynamics of SPMT are unknown at this point. Similarly unfounded is the definitive claim that it is known that P.f. MTs depolymerize upon fixation. Other possibilities should be considered. SPMT could also simply shorten in C1-arrested parasites.

While we agree with the reviewer that much about SPMT dynamics in schizonts remains unknown, we disagree with the claim that our consideration of SPMT depolymerization as a possible explanation for our observations is unfounded. Microtubule depolymerization is a well-known fixation and sample preparation artefact in both mammalian cells and a well-documented phenomenon in Plasmodium when parasites are washed with PBS prior to fixation. We convey in the text our belief that it is possible that SPMTs shorten in C1-arrested parasites as a result of drug treatment. However, it is our opinion that there simply is not enough evidence at this moment to conclusively pinpoint the cause of our observed depolymerization. As we mention in the text, further experiments are needed in order to determine with confidence whether depolymerization is a consequence of our fixation protocol, a consequence of C1 treatment (or the length of that treatment), or a biological phenomenon resulting from parasite maturation.

1. Line 324: "up to 30 daughter merozoites"

Schizonts can have more than 30 daughter merozoites, so we have not altered this statement.

1. Figure 4b. Line 354 The postulated breaking in two is not well visible and here the authors should attempt a more conservative interpretation of the data (especially with respect to those early basal complex dynamics).

We think that the basal complex dividing or breaking in two is the more conservative interpretation of our data. There is no evidence to suggest that a second basal complex is formed de novo and, while never before described using a basal complex protein, the cramp-like structure and dynamics we observe are consistent with that observed in early IMC proteins. We have updated the text to provide additional context and make the reasoning behind our hypothesis clearer.

1. Line 365: Commenting on their relative size would require a quantification of APR and basal complex size (can be provided in the text).

We are unsure what this is in reference to, as there is no mention of the APR in the basal complex section.

1. Lines 375ff: The claim that NHS Ester is a basal complex marker should be mitigated or more convincing images without the context of anti-CINCH staining being sufficient to identify the ring structure should be presented.

We have provided high quality, zoomed-in images without anti-CINCH staining in Fig. 5D&E, 6C, 7b, and Supplementary Fig. 8 that show that even in the absence of a basal complex antibody, the basal complex still stains densely by NHS ester.

.20. Line 407: The claim that there are differences in membrane potential along the mitochondria needs to be significantly mitigated. There are several alternative explanations of this staining pattern (some of which the authors name themselves). Differences in local compartment volume, differences in membrane surface, diffusibility/leakage of the dye can definitively play a role in addition to fixation and staining artefacts (also brought forward recently for U-ExM by Laporte et al. 2022 Nat Meth). Confirming the hypothesis of the authors would need significantly more experimental evidence that is outside the scope of this study.

We have significantly dampened and qualified the wording in this section. It now reads: “These clustered areas of MitoTracker staining were highly heterogeneous in size and pattern. Small staining discontinuities like these are commonly observed in mammalian cells when using MitoTracker dyes due to the heterogeneity of membrane potential from cristae to cristae as well as due to fixation artifacts. At this point, we cannot determine whether the staining we observed represents a true biological phenomenon or an artefact of this sample preparation approach. Our observed MitoTracker-enriched pockets could be an artifact of PFA fixation, a product of local membrane depolarization, a consequence of heterogeneous dye retention, or a product of irregular compartments of high membrane potential within the mitochondrion, to mention a few possibilities. Further research is needed to conclusively pinpoint an explanation.”

1. Fig. 7e: The differences in morphology using different fixation methods are interesting. Can the authors provide a co-staining of K13-GFP together with the better-preserved structures in the GA-containing fixation protocol to demonstrate that these are indeed cytostome bulbs?

Figure 7 has been changed substantially to show more clearly the preservation of the red blood cell membrane following PFA-GA fixation, followed by direct comparison of K13-GFP stained parasites fixed in either PFA only or PFA-GA. The cytostome section of the results has also changed to reflect this, the changed section now reads:

“PFA-glutaraldehyde fixation allows visualization of cytostome bulbThe cytostome can be divided into two main components: the collar, a protein dense ring at the parasite plasma membrane where K13 is located, and the bulb, a membrane invagination containing red blood cell cytoplasm {Milani, 2015 #63;Xie, 2020 #62}.While we could identify the cytostomal collar by K13 staining, these cytostomal collars were not attached to a membranous invagination. Fixation using 4% v/v paraformaldehyde (PFA) is known to result in the permeabilization of the RBC membrane and loss of its cytoplasmic contents65. Topologically, the cytostome is contiguous with the RBC cytoplasm and so we hypothesised that PFA fixation was resulting in the loss of cytostomal contents and obscuring of the bulb. PFA-glutaraldehyde fixation has been shown to better preserve the RBC cytoplasm65. Comparing PFA only with PFA-glutaraldehyde fixed parasites, we could clearly observe that the addition of glutaraldehyde preserves both the RBC membrane and RBC cytoplasmic contents (Figure 7c). Further, while only cytostomal collars could be observed with PFA only fixation, large membrane invaginations (cytostomal bulbs) were observed with PFA-glutaraldehyde fixation (Figure 7d). Cytostomal bulbs were often much longer and more elaborate spreading through much of the parasite (Supplementary Video 1), but these images are visually complex and difficult to project so images displayed in Figure 7 show relatively smaller cytostomal bulbs. Collectively, this data supports the hypothesis that these NHS-ester-dense rings are indeed cytostomes and that endocytosis can be studied using U-ExM, but PFA-glutaraldehyde fixation is required to maintain cytostome bulb integrity.”

1. It would be helpful to the readers to indicate in the schematic in Fig. 1b at which point NHS-Ester staining is implemented.

Figure 1b is slightly simplified in the sense that it doesn’t differentiate primary and secondary antibody staining, but we have updated it to reflect that antibody and dye staining are concurrent, rather than separate.

1. In Fig. 2B the second panel from the right the nuclear envelope boundary does not seem to be accurately draw as it includes the centrin signal of the centriolar plaque.

Thank you for pointing this out, it has now been redrawn.

1. Line 44-45: should read "up to 30 new daughter merozoites" (include citations).

We have included a citation here, but left it as approximately 30 daughter merozoites as the study found multiple cells with >30 daughter merozoites.

1. Line 49: considering its discovery in 2015 the statement that it has gained popularity in the last decade can probably be omitted.

This has been removed.

1. Fig S1 should probably read "2N" (instead of "2n"). Or alternatively "2C" could be fine.

1. Line 154: To help comprehension please define the term "branch number" in this context when it comes up.

A definition for branch has now been provided.

1. Fig. S5: To my estimation it is not an "early trophozoite", which is depicted.

While this parasite technically fits our definition of trophozoite, as it has not yet undergone nuclear division, we have swapped it for a visibly earlier parasite for clarity. This is the new parasite depicted

Author response image 4

1. Fig. 2a is not referenced before Fig. 2b in the text.

This has been addressed.

1. I could not find the reference to Fig. S2e and its discussion.

It was wrongly labelled as Supplementary Figure 2b in the text, this has now been addressed.

1. The next Figure referenced in the text after Fig. 2b is Fig. 4b. Fig.3 is only referenced and discussed later, which was quite confusing.

The numbering discrepancies have been addressed.

1. Line 196: Figure reference is missing.

This data did not have a figure reference, but the numbers have now been provided in-text.

1. Fig. 3c: Is "Branches per MTOC" not just total branches divided by two? If so it can be omitted. If not so please explain the difference.

Yes it was total branched divided by two, this has been removed from Figure 3c.

1. Figure 5c and 6d: The authors should show examples of the image segmentation used to calculate the surface area.

Surface area calculation was done in an essentially one step process. From maximum intensity projections, free-hand regions of interest were drawn, from which ZEN automatically calculates their area. Example as Author response image 5:

Author response image 5

1. Figure 7b should also show the NHS Ester staining alone for the zoom in.

We have included the NHS ester staining alone on the zoom on, but we have slightly changed the presentation of these two panels to show both the basal complex and cytostomes as follows:

Author response image 6

1. To which degree are Rhoptry necks associated with MTOC extensions?

This cannot easily be determined with the images we have so far. Before elongated necks are visible, the RON4 signal does appear pointed towards the MTOC extensions. Rhoptry necks don’t seem to elongate until segmentation, when the MTOC starts to move away from the apical end of the parasite. So it is possible there is a transient association, but we cannot easily discern this from our data.

https://doi.org/10.7554/eLife.88088.3.sa4

Article and author information

Author details

  1. Benjamin Liffner

    Department of Pharmacology and Toxicology, Indiana University School of Medicine, Indianapolis, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing
    Contributed equally with
    Ana Karla Cepeda Diaz
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1573-6139
  2. Ana Karla Cepeda Diaz

    1. Biological and Biomedical Sciences, Harvard Medical School, Boston, United States
    2. Division of Infectious Diseases, Boston Children’s Hospital, Boston, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing
    Contributed equally with
    Benjamin Liffner
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2033-461X
  3. James Blauwkamp

    Department of Pharmacology and Toxicology, Indiana University School of Medicine, Indianapolis, United States
    Contribution
    Data curation, Formal analysis, Investigation, Writing – original draft
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7639-2800
  4. David Anaguano

    1. Center for Tropical and Emerging Global Diseases, University of Georgia, Athens, United States
    2. Department of Cellular Biology, Franklin College of Arts and Sciences, University of Georgia, Athens, United States
    Contribution
    Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  5. Sonja Frolich

    1. Research Centre for Infectious Diseases, School of Biological Sciences, University of Adelaide, Adelaide, Australia
    2. Institute for Photonics and Advanced Sensing, University of Adelaide, Adelaide, Australia
    Contribution
    Methodology
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8673-3437
  6. Vasant Muralidharan

    1. Center for Tropical and Emerging Global Diseases, University of Georgia, Athens, United States
    2. Department of Cellular Biology, Franklin College of Arts and Sciences, University of Georgia, Athens, United States
    Contribution
    Resources, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6367-6284
  7. Danny W Wilson

    1. Research Centre for Infectious Diseases, School of Biological Sciences, University of Adelaide, Adelaide, Australia
    2. Institute for Photonics and Advanced Sensing, University of Adelaide, Adelaide, Australia
    3. Burnet Institute, 85 Commercial Road, Melbourne, Australia
    Contribution
    Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5073-1405
  8. Jeffrey D Dvorin

    1. Division of Infectious Diseases, Boston Children’s Hospital, Boston, United States
    2. Department of Pediatrics, Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Project administration, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5883-7271
  9. Sabrina Absalon

    Department of Pharmacology and Toxicology, Indiana University School of Medicine, Indianapolis, United States
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Methodology, Project administration, Writing – review and editing
    For correspondence
    sabsalon@iu.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2468-8156

Funding

American Heart Association (10.58275/aha.23post1011626.pc.gr.161347)

  • Benjamin Liffner

National Institutes of Health (R01 AI145941)

  • Jeffrey D Dvorin

National Institutes of Health (F31 AI172110)

  • Ana Karla Cepeda Diaz

NHMRC (GNT2020822)

  • Danny W Wilson

Hospital Research Foundation (C-MCF-52-2019)

  • Danny W Wilson

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank David Roos, Akhil Vaidya, and Rodolpho Ornitz Oliveira Souza for insightful discussions, Vincent Louvel for U-ExM expertise and sharing the protocol for cryopreservation and thawing of gels, and Taco Kooij, and Julie Verhoef for their help with MitoTracker experiments and critical reading of the manuscript. We thank Tobias Spielmann and Isabelle Henshall for generously sharing K13-GFP parasites, and insightful discussion. We thank Julian Rayner, Alan Cowman, Carole Long, BEI Resources (NIAID, NIH), and the European Malaria Reagent Repository for provision of antibodies. This work was supported by NIH R01 AI145941 (JDD) and F31 AI172110 (AKCD), and the American Heart Association 23POST1011626 (BL).

Senior Editor

  1. Dominique Soldati-Favre, University of Geneva, Switzerland

Reviewing Editor

  1. Olivier Silvie, Sorbonne Université, UPMC Univ Paris 06, INSERM, CNRS, France

Version history

  1. Preprint posted: March 24, 2023 (view preprint)
  2. Sent for peer review: April 3, 2023
  3. Preprint posted: May 23, 2023 (view preprint)
  4. Preprint posted: November 29, 2023 (view preprint)
  5. Version of Record published: December 18, 2023 (version 1)

Cite all versions

You can cite all versions using the DOI https://doi.org/10.7554/eLife.88088. This DOI represents all versions, and will always resolve to the latest one.

Copyright

© 2023, Liffner, Cepeda Diaz et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Benjamin Liffner
  2. Ana Karla Cepeda Diaz
  3. James Blauwkamp
  4. David Anaguano
  5. Sonja Frolich
  6. Vasant Muralidharan
  7. Danny W Wilson
  8. Jeffrey D Dvorin
  9. Sabrina Absalon
(2023)
Atlas of Plasmodium falciparum intraerythrocytic development using expansion microscopy
eLife 12:RP88088.
https://doi.org/10.7554/eLife.88088.3

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https://doi.org/10.7554/eLife.88088

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