Cell Motility of Early Emerging Apicomplexans

FACULTY OF SCIENCE

Cell motility of early emerging apicomplexans

Ph.D. Thesis

MAGDALÉNA KOVÁČIKOVÁ

Supervisor: RNDr. Andrea Bardůnek Valigurová, Ph.D. Department of Botany and Zoology

Brno 2019

Bibliographic Entry

Author: Mgr. Magdaléna Kováčiková Faculty of Science, Masaryk University Department of Botany and Zoology

Title of Thesis: Cell motility of early emerging apicomplexans

Degree programme: Biology

Specialization: Parasitology

Supervisor: RNDr. Andrea Bardůnek Valigurová, Ph.D.

Academic Year: 2018/2019

Number of Pages: 278

Keywords: actin; Apicomplexa; archigregarines; α-tubulin; blastogregarines; confocal laser scanning microscopy; electron microscopy; eugregarines; gregarines; motility; myosin; protococcidians; ultrastructure; Western blot

Bibliografický záznam

Autorka: Mgr. Magdaléna Kováčiková Přírodovědecká fakulta, Masarykova univerzita Ústav botaniky a zoologie

Název práce: Buněčná motilita raných linií výtrusovců

Studijní program: Biologie

Specializace: Parazitologie

Vedoucí práce: RNDr. Andrea Bardůnek Valigurová, Ph.D.

Akademický rok: 2018/2019

Počet stran: 278

Klíčova slova: aktin; Apicomplexa; archigregariny; α-tubulin; blastogregariny; elektronová mikroskopie; eugregariny; gregariny; konfokální laserová skenovací mikroskopie; motilita; myozin; protokokcídie; ultrastruktura; Western blot

Abstract

Apicomplexans represent an intensively studied group of unicellular organisms, comprising exclusively of the parasitic genera. Besides significant human and animal pathogens belonging to higher coccidians, Apicomplexa includes several early branching lineages such as protococcidians, blastogregarines and gregarines that are widely dispersed in marine, freshwater and terrestrial invertebrate hosts. Despite apicomplexans enormous diversity, the knowledge of their basal lineages still remains insufficient. Using a combination of experimental, microscopic, biochemical and molecular approaches, the presented thesis describes and compares the motility, subcellular organisation and phylogeny of the poorly investigated lineages of basal apicomplexans. The main aim of thesis was to investigate the cytoskeletal elements (their structure, organisation, nature, and function) responsible for highly diversified motility modes observed in studied marine and terrestrial apicomplexan representatives. The obtained results indicate that these diverse motility modes depend on the modifications of the parasite cell cortex (which is closely interconnected with its parasitism strategies and environment) and differ from substrate-dependent gliding generally described for apicomplexan zoites. In blastogregarines, Siedleckia nematoides performs active pendular/twisting motility similar to the bending/nematode like movement observed in Selenidium archigregarines (S. pygospionis, S. pherusae), while Chattonaria mesnili shows only weak motility with slow and intermittent bending movement. Though blastogregarines and archigregarines are phylogenetically separated groups, they show considerable similarities in their motility patterns and the organisation of their cell cortex with subpellicular microtubules functioning as the main motility motor. Trophozoites and gamonts of both these groups, despite bearing a striking resemblance to overgrown apicomplexan zoites, show no signs of gliding motility. In eugregarines, only the sporozoite stage exhibits the subcellular organisation characteristic of apicomplexan zoites. Ultrastructural study of sporozoites of urosporid eugregarines confirmed the presence of an apical set of organelles and subpellicular microtubules, the structures that are usually not observed at later stages in a majority of eugregarine species. Gliding motility, often accompanied with changes in the cell shape, was observed in eugregarines from terrestrial (Gregarina garnhami, Blabericola cubensis and Protomagalhaensia granulosae from insects) and marine (Polyrhabdina sp. from polychaetes, Cephaloidophora cf. communis from barnacles)

hosts. The main leading motor structure in the eugregarine gliding motility appears to be the polymerised form of actin that forms bundles of actin filaments (mostly rib-like myonemes and ectoplasmic network), supported by additional cytoskeletal structures (e.g. specific architecture of epicytic folds and cortical microtubules in some monocystid species) along with secretion of mucopolysaccharides coating the eugregarines surface. In contrast to the above mentioned basal apicomplexans, in protococcidian Eleutheroschizon duboscqi (developing in epicellular position within host-derived parasitophorous sac and sharing the features of gregarines and cryptosporidians) motility was not proven, despite the presence of actin rich subpellicular filaments presumably functioning as the parasite cytoskeleton.

Abstrakt

Výtrusovci jsou intenzivně zkoumanou skupinou jednobuněčných organismů, zahrnující výlučně parazitické rody. Kromě významných patogenů lidí a zvířat, náležejících do skupiny vyšších kokcidií, Apicomplexa zahrnují několik raných linií, jako jsou protokokcidie, blastogregariny a gregariny, které jsou široce rozšířeny u mořských, sladkovodních a suchozemských bezobratlých. Navzdory obrovské rozmanitosti výtrusovců, jsou znalosti o zástupcích bazálních liniích stále nedostatečné. S využitím kombinovaných experimentálních, mikroskopických, biochemických a molekulárních přístupů předkládaná práce popisuje a porovnává pohyb, subcelulární uspořádaní a fylogenezi u málo prozkoumaných skupin bazálních výtrusovců. Hlavním cílem práce byl výzkum cytoskeletálních struktur (jejich struktury, organizace, charakteru a funkce), zodpovědných za značně rozmanité způsoby pohybu pozorované u studovaných mořských i suchozemských zástupců výtrusovců. Výsledky studie naznačují, že tyto rozmanité způsoby pohybu závisí od modifikací buněčného kortexu parazita (který je úzce spjat se strategiemi parazitismu a životním prostředím parazita) a liší se od klouzavého, na substrátu závislého pohybu, obecně popsaného u zoitů výtrusovců. V případě blastogregarin, Siedleckia nematoides vykazuje aktivní kyvadlový/kroutivý pohyb podobný ohýbavému pohybu hlístic, pozorovanému u archigregarin rodu Selenidium (S. pygospionis, S. pherusae), zatímco Chattonaria mesnili vykazuje jen nevýrazný pohyb ve formě pomalého a přerušovaného ohýbání. I když jsou blastogregariny a archigregariny řazeny do různých fylogenetických skupin, vykazují značnou podobnost ve způsobu pohybu a v morfologii buněčného kortexu, kde subpelikulární mikrotubuly představují hlavní motor pohybu. Trofozoiti a gamonti obou skupin, navzdory jejich výrazné podobnosti s hypertrofovanými zoitmi výtrusovců, nevykazují žádné známky klouzavého pohybu. U eugregarin byla subcelulární organizace shodná se zoitmi výtrusovců pozorována pouze ve stadiu sporozoita. Ultrastrukturální výzkum sporozoitů urosporidních eugregarin potvrdil přítomnost apikálních organel a mikrotubulů, struktur, které obvykle nejsou pozorovány u pozdních stadií většiny eugregarin. Klouzavý pohyb, ve většině případů spojen se změnou buněčného tvaru, byl pozorován u eugregarin ze suchozemských (Gregarina garnhami, Blabericola cubensis a Protomagalhaensia granulosae z hmyzu) jako i mořských (Polyrhabdina sp. z mnohoštětinatců, Cephaloidophora cf. communis ze svijonožců) hostitelů. Hlavní strukturou zabezpečující

pohyb u klouzajících druhů eugregarin se jeví být aktin ve své polymerizované formě, tvořící provazce aktínových vláken (především žebrovité myonémy a ektoplasmatickú síť vláken), spolu s přídavnými cytoskeletálními strukturami (např. specifická stavba záhybů epicytu či kortikální mikrotubuly u některých monocystidních druhů) a sekrecí mukopolysacharidů pokrývajících povrch gregarin. Na rozdíl od výše zmiňovaných bazálních skupin výtrusovců, u protokokcídie druhu Eleutheroschizon duboscqi (vyvíjející se epicelulárně v parazitiformním vaku odvozeným z hostitelské buňky a sdílející charakteristiky gregarin i kryptosporidií) pohyb nebyl prokázán, i přes přítomnost subpelikulárních filamentů bohatých na aktin, pravděpodobně majících funkci jako cytoskelet parazita.

Acknowledgements

I am very grateful to my supervisor Dr. Andrea Bardůnek Valigurová for her valuable advice and consultations, as well as for managing many of the practical aspects of this study and helping me with samples evaluations and interpretations. I also would like to thank Assoc. Prof. Milan Gelnar, who gave me an opportunity to work on parasitic protist and financially supported this research. Many thanks to my colleagues and co-authors from different scientific institutions, namely Dr. Andrei Dakin, Dr. Naděžda Vaškovicová, Assoc. Prof. Gita G. Paskerova, Assoc. Prof. Timur G. Simdyanov and Prof. Isabelle Florent, for sharing their valuable knowledge and for the opportunity to work in their laboratories. I am very grateful for the support from the Department of Botany and Zoology, Faculty of Science, Masaryk University, towards my study. I would also like to thank the staff of the White Sea Biological Station of Lomonosov Moscow State University for help with material obtaining and to the members of Laboratory of Electron Microscopy, Biology Centre Czech Academy of Science for assistance in obtaining the electron microscopic data. Finally, I would like to express my gratitude to my family and friends for their ongoing support and encouragement in helping me to complete this work.

Financial support for this study was provided by the Czech Science Foundation, project No. GBP505/12/G112 (ECIP - Centre of excellence). Travel expenses were partially covered by the Mobility project 7AMB14FR013 from the Ministry of Education, Youth and Sports of the Czech Republic.

Contents 1 Introduction ...... 3 2 Aims of study ...... 4 3 Literature overview ...... 5 3.1 Introduction to the Apicomplexa ...... 5 3.1.1 General morphology and life cycle ...... 5 3.1.2 Motility and cytoskeleton ...... 8 3.2 Introduction to the studied groups of early emerging apicomplexans ...... 12 3.2.1 Protococcidians ...... 12 3.2.2 Blastogregarines ...... 13 3.2.3 Gregarines ...... 14 4 Material and methods ...... 25 4.1 Material collection and identification ...... 25 4.1.1 Apicomplexan parasites from marine polychaete, Scoloplos armiger ...... 25 4.1.2 Apicomplexan parasites from marine polychaete, Pygospio elegans ...... 25 4.1.3 Apicomplexan parasites from marine polychaete, Travisia forbesii ...... 26 4.1.4 Apicomplexan parasites from marine barnacle, Balanus balanus ...... 26 4.1.5 Apicomplexan parasites from desert locust, Schistocerca gregaria ...... 27 4.1.6 Apicomplexan parasites from tropical cockroach, Blaberus dicoidalis ..... 27 4.2 Experimental assays and light microscopy ...... 30 4.2.1 Experiments with cytoskeletal drugs ...... 30 4.2.2 Mucus shedding in drug-treated eugregarines ...... 31 4.2.3 Experiments with artificial seawater with changed ionic composition ...... 31 4.3 Electron microscopy ...... 33 4.3.1 Transmission electron microscopy (TEM) ...... 33 4.3.2 Scanning electron microscopy (SEM) ...... 33 4.3.3 Freeze-etching (FE TEM)...... 34 4.4 Confocal laser scanning microscopy (CLSM) ...... 35 4.5 Western blot ...... 36 4.6 Molecular analysis ...... 39 5 Results ...... 41 5.1 Published data ...... 41 5.2 Unpublished data...... 51 5.2.1 Manuscript in preparation...... 51

5.2.2 The effect of seawater ionic composition on motility of marine gregarines ...... 60 5.2.3 Western blot ...... 61 5.2.4 Morphological observations on terrestrial eugregarines ...... 63 5.2.5 Morphological observations on marine eugregarine Polyrhabdina sp...... 68 5.2.6 Molecular analysis ...... 70 6 Discussion ...... 72 7 Conclusions and perspectives ...... 79 8 References ...... 81 9 Original publications and author contribution ...... 94

1 Introduction

Apicomplexans (Apicomplexa Levine 1970, emend. Adl et al. 2012) are one of the most investigated group of protists, comprising exclusively parasites. Besides important pathogens of human and agricultural animals (e.g. Toxoplasma gondii, Plasmodium spp., Cryptosporidium spp., and Eimeria spp.), this group comprises the basal lineages as protococcidians, blastogregarines and gregarines restricted to invertebrate host. While the principles of motility and host cell invasion in apicomplexan parasites of the vertebrates are deeply investigated, the motility mechanisms and cytoskeletal elements driving the movement in early emerging apicomplexans are still poorly understood. In the last decades, numerous studies on Apicomplexa have dealt with the motility of their invasive stages (zoites) as they represent a potential target for chemotherapeutic intervention. Apicomplexan zoites perform a substrate-dependent gliding motility facilitated by a conserved form of actomyosin motor system that is intimately associated with their pellicle (the so called glideosome) and requires a stable network of subpellicular microtubules (Keeley and Soldati, 2004; Opitz and Soldati, 2002; Soldati et al., 2004; Tardieux and Baum, 2016). In contrast to vertebrate pathogens, the motility mechanism in early branching groups of Apicomplexa, such as lower coccidians, blastogregarines and gregarines parasitising invertebrates, still remains unclear. In basal apicomplexans several types of motility were described, such as typical wavy or twisting movement in blastogregarines, pendular/rolling/bending motility in Selenidium archigregarines, progressive linear gliding with or without obvious changes in cell shape in intestinal eugregarines or peristaltic/metabolic movement performed by coelomic and some intestinal eugregarines. Though several published studies focused on cytoskeletal elements involved in their motility, these usually lack the complex data set from experimental assays and combined microscopic approaches. In addition, a comparison of data obtained from various representatives is essential, to identify the correlations between the cell cortex organisation and motility of parasites. Until now, the majority studies focused on motility in gliding eugregarines from insect hosts (Ghazali and Schrével, 1993; Ghazali et al., 1989; Heintzelman, 2004; Heintzelman and Mateer, 2008; Valigurová et al., 2013). Filamentous structures associated with eugregarine cytoskeleton, rich in actin and myosin, and presumably involved in motility were demonstrated in several investigated species. Nevertheless, a very few experimental studies have been performed to verify the role of actin filaments in

eugregarine gliding. On the top of that, archigregarines and eugregarines from marine hosts are considered to have an ancestral position in phylogeny of Apicomplexa as supported by molecular evidence (Leander, 2008), however, only a few researches deal with structures responsible for their motility. Similarly, blastogregarines exhibiting both the coccidian and gregarine features, were for a long time an enigmatic group with uncertain phylogenetic position within the Apicomplexa and their motility mechanism was completely unknown. The publications involved in the presented thesis indicate that locomotion in early emerging apicomplexans differs from the substrate-dependent gliding observed in apicomplexan zoites and aims to clarify and compare the principles of motility within individual groups.

2 Aims of study

The main aim of this study was to investigate the motility in representatives of phylogenetically distant groups of basal Apicomplexa. Partial aims were set to achieve expected results:  Field collection and processing of material.  Observation and documentation of cell motility in selected apicomplexans in vitro.  Experimental motility assays using various cytoskeletal drugs affecting the motility or incubation of parasites in media with different ionic composition.  Comparative morphological analysis of structures responsible for motility.  Biochemical and molecular analysis of target proteins.

3 Literature overview

3.1 Introduction to the Apicomplexa

3.1.1 General morphology and life cycle

Apicomplexa (Levine 1970, emend. Adl et al. 2012) is a group of unicellular eukaryotes living as obligatory parasites widespread in both the vertebrate and the invertebrate hosts. This group includes morphologically and ecologically diverse protists, such as evolutionarily advanced Apicomplexa comprising causative agents of significant diseases of humans and agricultural animals (malaria, toxoplasmosis, cryptosporidiosis, coccidiosis, and piroplasmosis) and plesiomorphic representatives. The basal lineages (e.g. agamococcidians, protococcidians, blastogregarines, archigregarines and eugregarines) inhabiting exclusively the invertebrate hosts are practically unexplored because they are considered of no medical and economic significance, however, are crucial for our understanding of the evolution of parasitism and evolutionary pathways in Apicomplexa as whole (Paskerova et al., 2018, Valigurová et al., 2015). It is assumed that ancestral apicomplexans parasitised marine annelids and afterwards dispersed to other marine invertebrates. Their radiation and adaptation to the parasitic lifestyle likely took place before the era of vertebrates. Evolution of apicomplexans parasitism continued within freshwater and terrestrial invertebrates and finally reached vertebrate hosts (Cox, 1994). At least one stage in the apicomplexans life cycle is characterised by the presence of apical complex consisting of a conoid, polar ring(s), rhoptries, micronemes, and subpellicular microtubules (Adl et al., 2012; Chobotar and Scholtyseck, 1982). Rhoptries and micronemes are specialised secretory organelles that comprise products required for parasites motility, adhesion to and invasion of host cells and establishment of their niche such as parasitophorous vacuole or sac (Aikawa, 1971; Carruthers et al., 1999; Morrissette and Sibley, 2002a; Scholtyseck and Mehlhorn, 1970; Valigurová et al., 2007, 2015). The apical polar ring serves as microtubule-organising center (MTOC) anchoring the subpellicular microtubules by insertion (Russell and Burns, 1984). Apicomplexan parasites are covered by a three-layered pellicle consisting of plasma membrane, underlined by a closely apposed inner membrane complex (IMC) formed by flattened cortical alveoli and associated with cytoskeletal elements such as

microtubules, actin, myosin and a network of intermediate filamentous proteins (Morrissette and Sibley, 2002a) termed alveolins (Frénal et al., 2017) (Fig. 1). This cell envelope, consisting of pellicle with associated cytoskeletal structures, is referred to as a cell cortex. The IMC interconnected with the cytoskeleton plays a fundamental role in maintaining cell shape and contributes to the parasite motility (Kono et al., 2012). The subpellicular microtubules, organised in a single or several layers, are located just beneath the pellicle. These longitudinally oriented microtubules radiate from the polar ring and continue towards the cytosolic face of the pellicle, and in majority of apicomplexan zoites they end freely in the region behind the nucleus (approximately two-thirds of the length of the parasite) (de Souza and Attias, 2010; Morrissette and Sibley, 2002a) or run over the entire length of the parasite in eugregarine sporozoites, blastogregarine and archigregarine trophozoites/gamonts (Diakin et al., 2018; Simdyanov and Kuvardina, 2007; Valigurová et al., 2017). The arrangement of subpellicular microtubules varies among apicomplexan species, but the number, length, and organization are absolutely stereotyped within the life cycle stage of a species (Morrissette and Sibley, 2002a). Majority of apicomplexans examined to date (with the exception of the cryptosporidians and gregarines) possess an apicoplast organelle that is assumed to be an ancient secondary endosymbiosis with an algal cell (Toso and Omoto, 2007; Valigurová and Koudela, 2008; Valigurová et al., 2007; Zhu et al., 2000).

Figure 1. The pellicle of apicomplexans (taken from Frénal et al., 2017). Pellicle is composed of the plasma membrane (PM) and an inner membrane complex (IMC) formed from a single or several flattened vesicular sacs. Network of intermediate filament-like proteins, the alveolins, connects the IMC to the subpellicular microtubules.

A majority of apicomplexan species have a specific and complex parasitic life cycles, with alternating asexual and sexual multiplication (Siński and Behnke, 2004). Their life cycle in general starts with a sporozoite stage (an invasive, motile stage) develops through the vegetative stage of trophozoite into meronts (schizonts) and then divides repeatedly in host by merogony (schizogony) to produce numerous merozoites.

These enter the sexual phase of life cycle by transforming into gamont stage. Fused gamonts form a gametocyst to generate micro and macrogamets. Gametes pair together and develop into the zygote, giving the rise to new sporozoites via sporogony. In species lacking the vector phase in their life cycle, sporozoites are localise within the oocyst and/or sporocyst (Morrissette and Sibley, 2002a; Siński and Behnke, 2004) The most simple - direct - life cycle was observed in gregarines, where it is composed of gamogony (the sexual phase) and sporogony (the asexual phase) (Schrével and Desportes, 2015). Most apicomplexans are obligate intracellular parasites and are either in direct contact with the host cell cytoplasm or are encircled by a parasitophorous vacuole, formed by components of both the host cells and parasite (Votýpka et al., 2017). The epicellular localisation of parasite (previously referred to as intracellular but extracytoplasmatic) represents a transitional form between the extracellular and intracellular parasitism and was described for protococcidians, eimerid coccidians from poikilotherms, cryptosporidians, blastogregarines and eugregarines (Bartošová-Sojková et al., 2015; Jirků et al., 2002; Simdyanov et al., 2018; Valigurová, 2012; Valigurová et al., 2007, 2008, 2015). This specialised epicellular host-parasite interface reflects analogous modes of adaptation for development in similar environments. Extracellular but attached parasites are usually of a heteropolar nature, while intracellular apicomplexans are generally non-polar. In contrast to intracellular coccidians, evolutionary selection has presumably favoured an epicellular niche for above mentioned groups of apicomplexans, allowing them to more effectively evade the host responses, though these parasites thereby became dependent upon their connection with the host cell for nutrient acquisition (Valigurová et al., 2015). Epicellular cryptosporidians and protococcidians are covered with parasitophorous sac derived from the host cell membrane, while attached blastogregarines and gregarines are not enveloped (Simdyanov et al., 2018; Valigurová et al., 2007, 2015). Extracellular type of parasitism occurs in some eugregarine trophozoites living freely within the host (e.g. in coelomic fluid) and in all eugregarine gamonts detached from host tissue, where nutrient uptake is ensure presumably via micropores-mediated nutrition (Desportes and Schrével, 2013). Comparison of the attachment and feeding strategy observed in different apicomplexan lineages suggests that their parasitic lifestyle evolved from the free-living chromerids via the myzocytic feeding (myzocytosis = sucking out the host cell cytoplasm) of colpodellids, blastogregarines and archigregarines along with development of epicellular parasitism accompanied with nutrient transport presumably via modified attachment apparatus in eugregarines and

cryprosporidians. Alternatively, the nutrient transport via micropores was suggested for gregarines. Intracellular apicomplexans use both the transmembrane and micropore modes of nutrient uptake (Bartošová-Sojková et al., 2015; Desportes and Schrével, 2013; Simdyanov and Kuvardina 2007; Simdyanov et al. 2018; Valigurová et al. 2008, 2015).

3.1.2 Motility and cytoskeleton

Apicomplexa share a number of cytoskeletal elements (microtubules, actin, myosin, and intermediate filament-like proteins) with other eukaryotes, but with peculiarities unique to this group of parasites. For example, the actin filaments (F-actin) of Toxoplasma gondii are thought to be strikingly transient and observed only after treatment with actin polymerising drug (such as jasplakinolide) since majority of actin exists primarily in its globular form (Dobrowolski, Niesman and Sibley, 1997; Shaw and Tilney, 1999). Actin in Plasmodium spp. occurs in the form of dimers or short filaments (average length of 100 nm) (Kumpula et al., 2017; Schmitz et al., 2005). Nevertheless, the apparent lack of visible stable filaments is not fit for all apicomplexans, as in studied blastogregarines, archigregarines, eugregarines and protococcidian Eleutheroschizon duboscqi, the phalloidin labelling revealed the presence of F-actin, even without the application of filament-stabilising drugs (Valigurová, 2012; Valigurová et al., 2009, 2013, 2015, 2017; Kováčiková et al., 2017, 2018, 2019). Apicomplexans myosins are also considered unconventional, constituting a new class of unusually small “neckless” motors (Frénal et al., 2008; Heintzelman and Schwartzman, 1997; Morrissette and Sibley, 2002a). On the other hand, the subpellicular microtubules of Apicomplexa are noticeably stable and withstand the high pressure, cold and detergents traditionally used to isolate them (Morrissette et al., 1997). Gliding motility of apicomplexan zoites plays a crucial role in approaching the host cell and powers the migration of parasites across biological barriers, most likely also the active host cell entry (penetration) and egress from infected cells (Frénal et al., 2017; Meissner et al., 2013; Soldati-Favre, 2008). Several types of gliding motility have been described. For example, three types of movement were recorded in T. gondii tachyzoites i) circular gliding - parasite lies on its right side and moves in a counterclockwise manner, ii) twirling - parasite rights itself vertically, remaining attached to the substrate by its posterior end and spinning clockwise, iii) helical rotation – similar to twirling except that it occurs while the parasite is positioned horizontally, resulting in forward movement that

follows the path of a corkscrew (Håkansson et al., 1999). Similarly, wavy/flexing and rotate movements were observed in Plasmodium sporozoites attached by their apical end to the substrate (Vanderberg, 1974). This unique substrate-dependent gliding motility is thought to be facilitated by the glideosome complex consisting of adhesive proteins that are released apically and translocated to the posterior pole of the parasite by the action of an actomyosin motor localised in between the parasite plasma membrane and the IMC (Fig. 2) (Kappe et al., 2004; Keeley and Soldati, 2004; Opitz and Soldati, 2002). Gliding requires the action of the myosin motor complex (myosin A of class XIV and its myosin light chain) anchored on the IMC via gliding-associated proteins (GAP40, 45 and 50), and parasite actin that interacts with micronemal transmembrane proteins of the TRAP family via the glycolytic enzyme aldolase (Fig.2) (Frénal et al., 2010; Gaskins et al., 2004; Jewett and Sibley, 2003; Meissner et al., 2013; Sultan et al., 1997). Myosin A converts the chemical energy released by ATP hydrolysis into directed movement along actin filaments. A fast dynamic of actin polymerisation and rapid depolymerisation is required for gliding of apicomplexan zoites (Frénal et al., 2017; Opitz and Soldati, 2002). Intramembranous particles and/or a network of intermediate filament-like proteins (IMC network), connects the IMC (namely the internal cortical cytomembrane) to the subpellicular microtubules (Frénal et al., 2017; Kappe et al., 2004; Soldati and Meissner, 2004; Soldati et al., 2004) (Figs 1, 2). From opposite side of the IMC (external cortical cytomembrane) GAP45 protein recruits myosin to the cytomembrane and ensures the integrity of the pellicle during motility and appropriate spacing between the IMC and the plasma membrane (Frénal et al., 2010).

Figure 2. Model of actomyosin motor in apicomplexan zoites (taken from Soldati and Meissner 2004). Actomyosin motor is localised between the parasite plasma membrane and inner membrane complex. Interaction between the myosin A motor complex and the micronemal protein–host receptor complex is secured via aldolase/F-actin. The actomyosin motor is linked with subpellicular microtubules.

Despite verification of an important role of all of the above mentioned glideosome components for apicomplexan gliding motility, current studies have shown that motility can be preserved after the knockout of several key proteins associated with glideosome function (e.g. actin, myosin A or microneme proteins) and suggest that additional mechanisms must be involved in their motility and invasion (Egarter et al., 2014; Tardieux and Baum, 2016; Whitelaw et al., 2017). To determine the role of cytoskeletal structures in apicomplexan motility, application of drugs affecting the polymerisation of actin filaments and subpellicular microtubules has been performed in several studies (Dobrowolski and Sibley, 1996; Håkansson et al., 1999; Kováčiková et al., 2018, 2019; Morrissette and Sibley, 2002b; Münter et al., 2009; Stebbings et al., 1974; Stokkermans et al., 1996; Valigurová et al., 2017; Wetzel et al., 2003). As the apicomplexan actin filaments are extremely unstable (Morrissette and Sibley, 2002a) application of actin- modifying drugs (e.g. JAS and cytochalasin D) usually results in changes of parasite motility and/or its adhesion to host cell. In Plasmodium berghei and T. gondii, the JAS induced increase in F-actin concentration results in fast-moving zoites, indicating that

actin filaments are rate limiting (Münter et al., 2009; Wetzel et al., 2003). In contrast, treatment with cytochalasin D leads to increased adhesion dynamics and inhibits the host cell invasion and motility of parasites (Dobrowolski and Sibley, 1996; Håkansson et al., 1999; Münter et al., 2009). Despite the fact that subpellicular microtubules of Apicomplexa are considered as unusually stable it was reported that both the oryzalin and colchicine destroy the microtubules in T. gondii tachyzoites (Morrissette and Sibley, 2002b; Stokkermans et al., 1996), as well as in trophozoites/gamonts of studied blastogregarine and archigregarines (Kováčiková et al., 2019; Stebbings et al., 1974; Valigurová et al., 2017). While the shortening of the subpellicular microtubules was observed only at a high colchicine concentration (1 mM), microtubules were markedly sensitive to oryzalin and absent in oryzalin-treated parasites (de Souza and Attias, 2010; Stokkermans et al., 1996; Valigurová et al., 2017). Drug-treated replicative stages of Apicomplexa lacking subpellicular microtubules are non-polar, non-motile, and non- invasive (Stokkermans et al., 1996). Apicomplexan subpellicular microtubules have been usually investigated microscopically using the transmission electron microscopy (with or without immunogold labelling), by the indirect immunofluorescence with anti-tubulin antibodies, and/or biochemically by western blotting (Aikawa, 1971; Diakin et al., 2018; Hu et al., 2002; Kováčiková et al., 2019; Morrissette et al., 1997; Nichols and Chiappino, 1987; Ostrovska and Paperna, 1990; Paskerova et al., 2018; Schrével et al., 2016; Schwartzman et al., 1985; Simdyanov et al., 2018; Valigurová et al., 2015, 2017; Wetzel et al., 2003). The observation of actin filaments by electron microscopy is less frequent regarding to their nature; i.e. small thickness of actin filaments in general and prevailing monomeric form in T. gondii or short length in Plasmodium parasites (Schmitz et al., 2005; Wetzel et al., 2003). Nevertheless, both the actin and myosin have been frequently detected via immunofluorescence and western blotting using specific antibodies (Angrisano et al., 2012; Dobrowolski, Carruthers and Sibley, 1997; Drewry and Sibley, 2015; Endo et al., 1988; Heintzelman, 2004; Kováčiková et al., 2017, 2019; Valigurová et al., 2013, 2015, 2017; Whitelaw et al., 2017). Though phalloidin staining of filamentous actin was successful in several Apicomplexa species (Kováčiková et al., 2017; 2018, 2019; Preston and King, 1992; Valigurová, 2012; Valigurová et al. 2013, 2015, 2017), this method failed in other studies dealing with apicomplexans (even in the same gregarine species) (Dobrowolski, Niesman and Sibley, 1997; Forney et al., 1998; Heintzelman, 2004; King, 1988). The inability to stain apicomplexan microfilaments with phalloidin may be caused

by an unusually short filament length or alternatively, the phalloidin binding site may be masked by filament binding proteins, probably in combination with a weak affinity for phalloidin or its fluorescent analogues (Schüler et al., 2005). Possibly, only certain filament conformations allow phalloidin to bind on F-actin (Schmitz et al., 2010). As a novel tool for visualising F-actin dynamics in T. gondii living cells, actin-chromobody has been successfully tested (Periz et al., 2017).

3.2 Introduction to the studied groups of early emerging apicomplexans

3.2.1 Protococcidians

Protococcidiorida Kheissin, 1956 comprises parasites of marine invertebrates characterised by the absence of merogony in their life cycle (Levine, 1973; Perkins et al., 2000). Sporozoites invade the host intestinal epithelium and, depending on species, their development continues in the intestine, coelom or vascular tissues (Adl et al., 2012). Gamont development is extracellular, though the intracellular localisation of early stages has been reported in some species (Levine, 1973). Protococcidians comprise five families divided into eight genera: Angeiocystis, Coelotropha, Grellia, Eleutheroschizon, Mackinnonia, Myriosporides, Myriospora, Sawayella (Adl et al., 2012). Genus Eleutheroschizon Brasil, 1906 (Eleutheroschizonidae Chatton and Villeneuve, 1936) has been considered as enigmatic for a long time, because description of individual species, their morphology and life cycle have been reported only in few studies (Brasil, 1906; Caullery and Mesnil, 1898; Chatton and Villeneuve, 1936b; Levine, 1973; Valigurová et al., 2015). Representatives of this group are characterised by an epicellular development, with matured gamonts detaching from the host tissue and dispersing into the environment where gametogenesis and sporogenesis take place (Perkins et al., 2000; Valigurová et al., 2015). Recently, a comprehensive microscopic analysis focusing on parasite’s attachment strategy was performed on Eleutheroschizon duboscqi, a parasite of the marine polychaete Scoloplos armiger (Valigurová et al., 2015). This study showed the parasite being covered by a host-derived parasitophorous sac and attached to the host intestinal cell via a complicated attachment apparatus (consisting of lobes and filamentous fascicles organised in circles). While the motility was not confirmed in investigated developmental stages of E. duboscqi, the presence of polymerised form of

cytoskeletal proteins - actin and tubulin - was demonstrated in both the parasitophorous sac and the parasite using experimental assays and confocal microscopy. Myosin was restricted to the parasitophorous sac only and spectrin was accumulated in the caudal region of the sac (tail). Despite the positive α-tubulin labelling of the parasite surface and cytoplasm, microtubules were present only during early development and disappeared during trophozoite maturation. Preservation of tubulin in later developmental stages suggests its presence in a nonpolymerised form or in other tubulin-rich structures (Valigurová et al., 2015). Interestingly, subpellicular bands of longitudinally oriented actin-rich filaments forming beneath the IMC during trophozoite development, appear to replace the microtubules. Consistent with the disappearance of microtubules during trophozoite maturation, the invading zoites and the earliest stages, with their pointed end attached to the host epithelium, seem to exhibit oscillating movements, while the early trophozoites show only weak movement. Although a few attached advanced stages exhibited slight signs of movements and/or cytoplasmic streaming resembling metaboly, due to the intense waving and beating motion of enterocyte cilia it was not possible to determine with certainty whether or not the parasite actually moved (Valigurová, personal observation).

3.2.2 Blastogregarines

Blastogregarines (Blastogregarinida Chatton and Villeneuve, 1936) is a small group within the Apicomplexa, which taxonomic affiliation was uncertain until recently. Complex morphological and molecular phylogenetic study showed blastogregarines as a relatively isolated group of plesiomorphic apicomplexans classified as a separate class Blastogregarinea (Simdyanov et al., 2018). Blastogregarines are intestinal epicellular parasites of marine polychaetes (family Orbiniidae) and are represented by two families, Siedleckiidae Chatton et Villeneuve, 1936 and Chattonariidae, fam. nov. (Simdyanov et al., 2018) including only four species – S. nematoides, S. caullery, S. dogieli and C. mesnili (formerly S. mesnili) (Caullery and Mesnil, 1898; Chatton and Dehorne, 1929; Chatton and Villeneuve, 1936a; Simdyanov et al., 2018). Typical features of blastogregarines are persistent multinuclearity in trophozoites and gametogenesis by means of budding of the parasite posterior end (Chatton and Dehorne, 1929; Chatton and Villeneuve, 1936a; Simdyanov et al., 2018). Their life cycle is characterised by the absences of syzygy and gametocyst stages, and by pronounced anisogamy (coccidian feature) (Chatton and

Villeneuve, 1936a). In contrast, epicellular localisation of blastogregarines is similar to that observed in gregarines. The vermiform body shape of trophozoites and gamonts, their attachment to host tissue via a mucron and the typical bending/pendular motility are reminiscent of Selenidium archigregarines (plesiomorphic gregarine group) (Desportes and Schrével, 2013; Simdyanov et al., 2018; Valigurová et al., 2017). The motility of type species, S. nematoides from the intestine of marine polychaete S. armiger has been studied in detail. Attached trophozoites and gamonts perform a highly active pendular and twisting undulation and sometimes spasmodic motility, while detached cells bend from side to side. This waving movement is initiated at the apical end and proceeding distally. Experimental assays and microscopic observation revealed that the main motor driving the blastogregarines movement are the subpellicular microtubules arranged in one or several layers, which underline the pellicle and extend along the entire length of the cell (Simdyanov et al., 2018; Valigurová et al., 2017). The involvement of filamentous actin in blastogregarine movement was confirmed using cytoskeletal probes, revealing the existence of cross-linking protein complexes (presumably of actin nature) located beneath the IMC and associated with the subpellicular microtubules, and appearing to control the microtubule spacing (Valigurová et al., 2017). The positive correlation between the active motility and abundant mitochondria, located beneath the microtubules and providing the chemical energy for activity of microtubule-associated molecular motors, was proposed, similarly to that observed in archigregarines (Leander, 2006; Mellor and Stebbings, 1980; Paskerova et al., 2018; Schrével, 1971a; Simdyanov and Kuvardina, 2007; Valigurová et al., 2017).

3.2.3 Gregarines

Gregarinomorpha Grassé, 1953 is a highly diversified basal lineage of Apicomplexa widespread in marine, freshwater and terrestrial invertebrates (Schrével and Desportes, 2015). A recent study reported the occurrence of oocysts of Nematopsis temporariae in a liver of tadpoles and representing the first eugregarine observed in vertebrates (Chambouvet et al., 2016). According to the phylogenetic and morphological studies, gregarines are considered as a sister group to Cryptosporidium spp. (Adl et al., 2012; Aldeyarbi and Karanis, 2016; Barta and Thompson, 2006; Carreno et al., 1999; Leander Clopton and Keeling, 2003; Leander, Harper and Keeling, 2003; Simdyanov et al., 2017; Valigurová et al., 2007, 2008). Traditionally the gregarines are subdivided into

archigregarines, eugregarines and neogregarines. This branching is in some extent supported by molecular phylogeny, however, it is inferred mostly from gregarines life cycles (Desportes and Schrével, 2013; Leander, 2008; Schrével et al., 2016). Archigregarines are intestinal parasites of marine invertebrates characterised by the persistence of apical complex in trophozoite stage, myzocytotic feeding and the absence of septum inside the cell (Schrével and Desportes, 2015). The representatives of the most abundant and best described group, eugregarines, are widespread in intestine or body cavities of marine, freshwater and terrestrial invertebrates. Eugregarines are either aseptate or septate, with a transversal septum separating the protomerite from the deutomerite (Schrével and Desportes, 2015). In contrast to prevailing extracellular development in eugregarines, neogregarines develop intracellularly within Malpighian tubules, fat body and more rarely in intestine of insects and undergo multiple rounds of merogony after invading the host cell (Schrével and Desportes, 2015). The development of archigregarines and eugregarines takes places in different organs/tissues or body cavities (depending on group as mentioned above) of their hosts, where they are attached to the tissue or living freely in coelomic fluid (Desportes and Schrével, 2013; Valigurová, 2012; Valigurová and Koudela, 2008). In addition, an intracellular phase of early development has been observed in some species (Desportes and Schrével, 2013; Paskerova et al., 2018; Ray, 1930). These specific locations could explain the diversity in cell size, shapes and motility modes displayed by gregarines during the vegetative phase of their life cycle (Desportes and Schrével, 2013). Generally, gregarines have a monoxenous life cycle starting by sporozoites excysted from the oocyst in the lumen of host intestine. The released sporozoites invade the host tissue, hereby initiating the vegetative phase followed by a transformation of the sporozoite to the trophozoite (feeding) stage. The growing trophozoites are attached to the host tissue by a specialised apical part forming a mucron or an epimerite. After finishing the vegetative phase of their life cycle, trophozoites detach and transform into sexual stages, called the gamonts, which are usually motile (Schrével and Desportes, 2015). The most evident characteristic that differentiates gregarines from the other Apicomplexa is the sexual association between two gamonts, called the syzygy. A gametocyst wall forms around the two paired gamonts (male and female). Inside the gametocyst, the gamonts produce the gametes, which later fertilise and form a zygote. Zygote becomes enveloped by an oocyst wall and produces sporozoites by a process called sporogony (Desportes and Schrével, 2013). When the development is completed, infective oocysts are released into the

environment by gametocyst dehiscence through ducts or by direct rupture of gametocyst wall. These oocysts dispersed in the environment are ingested by a host and the life cycle repeats (Kolman et al., 2015; Peregrine, 1970) (Fig. 3 and Fig. 15 in subchapter 5.2.4).

Figure 3. The schematic diagram showing the life cycle of eugregarine Blabericola cubensis (taken from Kolman et al., 2015). a, b. Oocysts (a) with sporozoites (b). c. Sporozoites excyst from oocyst invade the host intestinal epithelial cells. d. Attached trophozoite. e, f. Detached gamonts pair together and form syzygy. g, h. Formation of a gametocyst wall surrounding the gamonts. i. Gametogony. j, k. Zygotes and oocyst formation. l. Sporogony. m. Gametocyst releasing infective oocyst into the environment by ducts formed in gametocyst wall.

The life cycle of neogregarines and some archigregarines is, in addition, characterised by a merogony (schizogony) during the vegetative growth of the trophozoite (Paskerova et al., 2018; Schrével and Desportes, 2015). The dissemination of the gregarines within the host occurs in species living in the coelomic cavities and also in

intracellular neogregarines. In such cases, necrophagy or cannibalism are considered as the potential ways of parasite transmission (Schrével and Desportes, 2015; Valigurová and Koudela, 2008). In gregarine trophozoites and gamonts, several types of motility have been described. These diverse modes of gregarine motility represent specific adaptations to parasitism in different environments (Valigurová et al., 2013). For example, archigregarines of the family Selenidiidae Brasil, 1907 exhibit pendular, coiling, rolling, bending (nematode-like) movements that appear to be facilitated by regular sets of subpellicular microtubules (Kováčiková et al., 2019; Paskerova et al., 2018; Schrével et al., 1974, 2016; Stebbings et al., 1974). In contrast, majority of intestinal eugregarines exhibit a progressive gliding with or without obvious changes in cell shape and accompanied by the secretion of mucus leaving a trail behind the gliding eugregarine (King, 1981, 1988; Kováčiková et al., 2017, 2018; Mackenzie and Walker, 1983; Valigurová et al., 2013; Walker et al., 1979). Another type of motility, exhibited by coelomic (Urosporidae Léger, 1892 and Monocystidae Bütschli, 1882) and some intestinal eugregarines (e.g. Didymophyes gigantea), is the so-called peristaltic or metabolic movement (Desportes and Schrével, 2013; Diakin et al., 2016; Hildebrand and Vinckier, 1975; Landers and Leander, 2005; Leander et al., 2006; MacMillan, 1973).

3.2.3.1 Archigregarines

Archigregarines are restricted to marine environments and are considered, on the basis of their plesiomorphic characteristics, to be the most ancestral representatives of gregarines and perhaps apicomplexans as a whole (Leander, 2008; Schrével, 1971a). Archigregarines are a key group to understand the early evolution of Apicomplexa (Paskerova et al., 2018). The life cycle of archigregarines belonging to the family Selenidiidae takes place in the intestine of polychaetes, sipunculids, and some hemichordates (Schrével and Desportes, 2015). The Selenidium trophozoites are often called “hypersporozoites” or “hypertrophied zoites” due to striking morphological similarities with the invasive apicomplexan stages (zoites) (Schrével, 1971b; Schrével and Desportes, 2015; Schrével et al., 2016). The archigregarine trophozoites are attached to the host cell via a mucron that enables the myzocytotic feeding through the well-developed organelles of the apical complex (Paskerova et al., 2018; Schrével, 1971b; Schrével and Desportes, 2015; Schrével et al., 2016; Simdyanov and Kuvardina, 2007). The typical apicomplexan three-layered

pellicle of archigregarines, consisting of plasma membrane and IMC, is covered by a glycocalyx layer (cell coat) and folded into broad and low folds separated by grooves with numerous micropores. A set of regularly spaced subpellicular microtubules, running parallel with each other along the longitudinal cell axis, is located just beneath the IMC. Several additional deeper layers of microtubules are often present, arrangement of which is irregular and rather incomplete (Paskerova et al., 2018; Schrével and Desportes, 2015). Each microtubule is surrounded by an electron-transparent hexagonal sheath (Kováčiková et al., 2019; Leander, 2007; Paskerova et al., 2018; Schrével et al., 2016). Depending on species, Selenidium archigregarines exhibit various types of movement (Desportes and Schrével, 2013; Fowell, 1936; Kováčiková et al., 2019; Leander, 2007; Leander, Harper and Keeling, 2003; Mellor and Stebbings, 1980; Paskerova et al., 2018; Rueckert and Horák, 2017; Schrével and Desportes, 2015; Schrével and Philippe, 1993; Schrével et al., 2016; Simdyanov and Kuvardina, 2007; Stebbings et al., 1974; Wakeman et al., 2014). For example, S. pendula and S. orientale exhibit pendular motility, while S. sabellariae performs coiling and S. hollandei rolling movements (Rueckert and Horák, 2017; Schrével et al., 2016; Simdyanov and Kuvardina, 2007). Slow-motion whip like movements (the propagation of bending waves) in attached trophozoites and coiling movements in detached parasites, observed in S. fallax, were preceded by a slight longitudinal contraction of the cell (Mellor and Stebbings, 1980; Stebbings et al., 1974). Bending, twisting, and contracting movements are performed by S. serpulae and S. sabellae (Leander, 2007; Rueckert and Horák, 2017). Bending motility of the entire cell, usually with one bend along the cell is typical for S. pherusae (Paskerova et al., 2018). Trophozoites and gamonts of S. pygospionis exhibit a bending (nematode- like) motility with a coiling of their anterior end (Kováčiková et al., 2019; Paskerova et al., 2018). Similarly, nematode-like bending movement combined with twisting was observed in S. terebellae (Leander, Harper and Keeling, 2003; Wakeman et al., 2014). In contrast to the active movement in above mentioned Selenidium species, S. spiralis and S. opheliae show only weak bending movement, while S. melogena is completely immobile (Rueckert and Horák, 2017; Wakeman et al., 2014). This bending/rolling/pendular motility of Selenidium spp. is reminiscent of movement reported in apicomplexan sporozoites (Leander, 2008) and blastogregarines (Simdyanov et al., 2018; Valigurová et al., 2017). Movements enable archigregarines to find syzygy partners, invade adjacent epithelial cells, and to move nutrients across their cell surface within the intestinal lumen (Leander, 2007).

A mechanism based on cooperation between the pellicle (IMC) and longitudinally oriented subpellicular microtubules, acting like an analogue of the musculocuticular system of nematodes, was proposed for archigregarine motility. In this hypothesis, the pellicle plays a role of skeletal unit, while the subpellicular microtubules are the motile unit (Leander, 2007; Paskerova et al., 2018; Stebbings et al., 1974). Meanwhile, the sliding of microtubules likely provided by microtubule-associated molecular motors (MAPs) such as kinesins and dyneins, comparable to the system occurring in a ciliary axoneme, was suggested as the mechanism for the propagation of bending waves in Selenidium spp. (Mellor and Stebbings, 1980; Stebbings et al., 1974). Presumably, the electron-transparent sheaths surrounding the individual microtubules play an important role in microtubular sliding (Leander, 2007; Mellor and Stebbings, 1980; Stebbings et al., 1974). Association between the numerous mitochondria (providing the chemical energy in ATP for MAPs activity) and subpellicular microtubules appears necessary to ensure functional cell motility (Leander, 2006; Mellor and Stebbings, 1980; Schrével, 1971a). Indeed, the ectoplasm of actively moving representatives of Selenidiidae comprises higher number of subpellicular microtubules compared to slow-moving species, and their abundant mitochondria are concentrated in the cortical zone under the microtubules (Paskerova et al., 2018; Schrével, 1971a; Simdyanov and Kuvardina, 2007). As an evidence, the immotile species S. melongena lacking assembly of subpellicular microtubules can be mentioned (Wakeman et al., 2014). The rate of movement of Selenidium species has been also proposed to correlate with the availability of oxygen in their environment (Schrével, 1971b). In addition, the involvement of the axial streak together with putative microfilaments was suggested to participate in archigregarine motility (Fowell, 1936; Paskerova et al., 2018). Previous studies testified the role of subpellicular microtubules in the motility of archigregarines using the colchicine, blocking the assembly of microtubules subunits and urea, inducing the depolymerisation of microtubules (Schrével et al., 1974; Stebbings et al., 1974). The cessation of bending movements in S. fallax attached trophozoites was detected after the application of colchicine in concentrations of 0.1 – 2% (Stebbings at al., 1974). The number of microtubules in colchicine-treated archigregarines was significantly reduced and the remaining microtubules appeared to be destroyed. In addition, the pattern of pellicular folding was altered compared to native parasites (Stebbings et al., 1974). Similarly, the incubation of S. hollandei trophozoites in seawater containing 0.6 - 1.0 M urea resulted in depolymerisation of subpellicular microtubules accompanied by blocking

of the rolling motility (Desportes and Schrével, 2013; Schrével et al., 1974). Accordingly, the motility of S. pygospionis ceased after treatment with microtubule-destroying drugs – colchicine and oryzalin (Kováčiková et al., 2019). Incubation of parasites with colchicine caused their rigidity proceeding from posterior end of the cell to the anterior end, and fading of microtubules in the outermost layer. The fluorescent α-tubulin staining of colchicine-treated S. pygospionis revealed the preservation of the longitudinal arrangement of microtubules, but these were destroyed (i.e. the stained lines corresponding to microtubules were interrupted) and small tubulin clusters formed. Treatment of S. pygospionis with oryzalin resulted in significant reduction of microtubule numbers (calculated per 1 µm of pellicle length) and formation of distinct α-tubulin clusters (Kováčiková et al., 2019). The motility and the microtubule polymerisation recovered after returning the S. hollandei and S. pygospionis to the seawater (Desportes and Schrével, 2013; Kováčiková et al., 2019; Schrével et al., 1974). The involvement of actin in archigregarine motility has been also tested using the actin-modifying drugs – JAS and cytochalasins. The incubation with cytochalasin B resulted in inhibition of S. pendula pendular movement (Ghazali and Schrével, 1995) and the treatment of S. pygospionis with JAS and cytochalasin D caused movement suppression or complete cessation of bending, indicating the role of F-actin in archigregarines motility (Kováčiková et al., 2019). Overall, the subpellicular microtubules appear to be the main motor in archigregarine motility, while the actin filaments provide rather supportive force, considering potential microtubule-membrane interactions ensured via actin filaments (Ghazali and Schrével, 1995; Kováčiková et al., 2019; Stebbings et al., 1974). Support for this theory can be found in a study on blastogregarine S. nematoides, which revealed the existence of cross-linking protein complexes, presumably of actin nature, consisting of proteins embedded in the IMC and the network surrounding the individual microtubules and anchoring them to the IMC hereby controlling the spacing of subpellicular microtubules (Valigurová et al., 2017).

3.2.3.2 Eugregarines

Eugregarines parasitise a wide range of aquatic and terrestrial hosts and their ultrastructure, morphology, phylogeny and motility have been studied most intensively within known gregarine groups. The three-layered pellicle, consisting of the plasma membrane and IMC, creates numerous epicytic folds arranged in longitudinal lines

separated by grooves (Schrével and Desportes, 2015). The density and dimensions of the folds depend on i) the degree of the trophozoite maturation, ii) the anterior-posterior gradient of the cell surface; and iii) the sexual differentiation of the gregarine (Schrével and Desportes, 2015). The epicytic folds of some eugregarines show an undulating pattern, usually with alternating undulated and straight folds (Valigurová and Koudela, 2008; Valigurová et al., 2013; Vávra and Small, 1969; Vivier, 1968). These lateral undulations were suggested to provide force for eugregarine gliding (Schrével and Phillipe, 1993; (Vávra and Small, 1969; Vivier 1968), possibly mediated by an actomyosin system (Desportes and Schrével, 2013). The usually dilated tips of epicytic folds comprise the so called 12-nm filaments exhibiting the properties of intermediate filaments and the rippled dense structures, both thought to have a supportive scaffolding function or possibly representing a component of the gliding motility (Desportes and Schrével, 2013; Kováčiková et al., 2017; Schrével et al., 1983; Valigurová et al., 2013; Walker et al., 1984). Potential connections between the 12-nm apical filaments and intramembranous particles (IMPs) anchoring these filaments to the IMC (internal cytomembrane) has been discussed as a system facilitating the eugregarine gliding (Dallai and Talluri, 1983; Walker et al., 1984). Moreover, links were observed between the external cytomembrane of IMC and plasma membrane; on the basis of these observations involvement of 12-nm filaments-rippled dense structures complex in eugregarines movement was proposed via undulation of epicytic folds (Desportes and Schrével, 2013). It has been shown that the number of 12-nm filaments does not influence the gliding rate, but rather seems to control the direction of the movement (i.e. gliding path of species equipped with a low number of 12-nm filaments was rather semi-circular than linear) (Kováčiková et al., 2017; Valigurová et al., 2013). Recent study using cytoskeletal drugs, however, contradicts the expectation that the lateral undulation of epicytic folds provides the main force behind eugregarine gliding (Kováčiková et al., 2018). Experimental assays revealed that the wavy pattern of epicytic folds in G. garnhami is similar in treated and non-treated parasites (even in eugregarines incubated with cytochalasin D, where the gliding motility was completely blocked) and demonstrated the role of ectoplasmic network and myonemes on eugregarines motility (Kováčiková et al., 2018). The mucus trail left behind gliding eugregarines was previously investigated as a potential lubricant for gliding locomotion (Schwiakoff, 1894; Valigurová et al., 2013; Walker et al., 1979). Although mucus flow accompanies gliding, there is a lack of evidence to show whether it is the cause or a consequence of gliding movement

(Mackenzie and Walker, 1983). It has previously been suggested that the secreted mucus pushes the eugregarine forward passively (Desportes and Schrével, 2013). Nevertheless, another study showed the translocation of concanavalin-A beads posteriorly along the gregarine surface at rates that are similar to gliding rates, and that the large beads comparable in mass to the eugregarine itself can be actively translocated (King, 1981). These experimental data pointed that eugregarines generate substantial locomotory forces and hence, it is more likely that the mucus acts as a lubricant. Accordingly, the presence of mucus trails behind gliding G. garnhami in non-treated and experimentally affected parasites indicates this mucosubstance to have a supportive function, maintaining an environment suitable for gregarine movement on a solid surface. It can be expected that the cell coat covering the entire surface of the gregarine is continuously reformed by the secretion of its components from the cell (Kováčiková et al., 2018). It has also been shown that the increased load of mucus in eugregarine cytoplasm is correlated with the gliding rate (Valigurová et al., 2013). In addition, the site of the eugregarine pellicle, where the superfolds (individual epicytic folds clustering on several projections distributed regularly throughout the cell periphery) are grouped together, exhibited a denser secretion and accumulation of mucus, suggesting this part to be the gliding side (Kováčiková et al., 2018) (Fig. 4).

Figure 4. Schematic diagram illustrating a possible function of cortical filaments (myonemes, ectoplasmic network), facilitating the gliding motility (taken form Kováčiková et al. 2018). a. G. garnhami in non-gliding phase. The myonemes and the ectoplasmic network are evenly distributed around the ectoplasm. b. G. garnhami gamonts during gliding on solid substrate. Note the presence of protruding superfolds grouping together caused by the contraction of myonemes along with more compact ectoplasmic network and denser accumulation of mucus in this region. black arrow – mucus drops, ecn – ectoplasmic network, s – superfolds, white arrowhead – myonemes, white asterisk – cytoplasm.

For a deeper understanding of the actomyosin system in eugregarines, mostly Gregarina species were studied using biochemical and molecular approaches (Baines and King, 1989; Ghazali and Schrével, 1993; Ghazali et al., 1989; Heintzelman, 2004;

Heintzelman and Mateer, 2008; Kováčiková et al., 2018; Philippe et al., 1982; Valigurová, 2012; Valigurová et al., 2013). A layer consisting of fibrils joining together by connectives forming a network, called the myocyte, encircles the eugregarines. The myocyte is supposed to be formed by contractile elements that are responsible for eugregarine motility and bending (Crawley, 1905). In more recent publications (Beams et al., 1959; Hildebrand, 1980; Valigurová and Koudela, 2008; Valigurová et al., 2013; Walker et al., 1979) these filaments in eugregarines were divided into two types depending on their thickness and position relative to the gregarine axis: i) an anastomosing ectoplasmic network of intermediate filamentous proteins underlying the IMC, and ii) annular rib-like myonemes located deeper in the edge of the ectoplasm and endoplasm. In Gregarina representatives, the myonemes and ectoplasmic network are considered to be the major contractile elements providing the driving force for cell locomotion and/or protomerite bending (Beams et al., 1959; Kováčiková et al., 2018; Valigurová et al., 2013) (Fig. 4). The function of these structures was verified by experimental assays using JAS and cytochalasins, which demonstrated their significant impact on eugregarines motility (King, 1988; Kováčiková et al., 2018; Valigurová et al., 2013; Walker et al., 1979). Several (immuno)fluorescent and biochemical studies were performed to analyse the nature of these cytoskeletal filaments. For example, a study, in which actin was specifically labelled with antibody generated to G. polymorpha actin, revealed its bilaminar staining pattern (Heintzelman, 2004). The outer layer of actin was organised into longitudinal lines, copying the arrangement of epicytic folds, while the inner layer corresponding to the rib-like myonemes was oriented perpendicular to the longitudinal cell axis (Heintzelman, 2004). The same bilaminar pattern, present after fluorescent staining with phalloidin in G. garnhami gamonts, indicates that the myonemes and ectoplasmic network are composed of actin filaments (F-actin). Moreover, the application of cytochalasin D, resulting in rapid motility cessation in G. garnhami and vanishing of myonemes (caused by drug’s depolymerising effect on actin filaments) proved that myonemes are essential for gamonts gliding (Kováčiková et al., 2018). The association of actin filaments with different types of myosins (A, B and F) appears to be likely, because myosins were immunoflurescently detected in other Gregarina spp. and in C. cf. communis, in similar localisation (i.e. restricted to epicytic folds and/or annular myonemes) and organisation pattern as F-actin in G. garnhami (Heintzelman, 2004; Heintzelman and Mateer, 2008; Kováčiková et al., 2017, 2018; Valigurová et al., 2013).

Actin and myosin participation in rapid gliding motility, as well as myonemes-mediated bending was proposed for Gregarina polymorpha (Heintzelman, 2004). Recent studies suggest, that the dynamic process of actin polymerisation, proposed as glideosome in apicomplexan zoites, is not essential for gliding in eugregarines, but only the polymerised form of actin seems to be the main leading motor structure responsible for their motility. The incubation of eugregarines with JAS, inducing further stabilisation of F-actin, did not significantly change the motility of gamonts, while the treatment with cytochalasin D almost immediately blocked eugregarines motility due to depolymerisation of existing actin filaments (i.e. due to degradation of ectoplasmic network and myonemes complex under the pellicle) (Kováčiková et al., 2018). Hence, the gliding motility in eugregarines differs from substrate-dependent gliding based on glideosome described for apicomplexans zoites. This is further supported by the fact that in eugregarine species used for investigation of the gliding motility no subpellicular microtubules (essential components in apicomplexan glideosome machinery) were observed (Kováčiková et al., 2017, 2018; Valigurová et al., 2013).

4 Material and methods

4.1 Material collection and identification

Several representatives of basal apicomplexans were collected from marine and terrestrial invertebrate hosts. This chapter includes only the procedures performed by the author. Summarisation of all apicomplexan species discussed in presented thesis is listed in Table 1.

4.1.1 Apicomplexan parasites from marine polychaete, Scoloplos armiger

 protococcidian Eleutheroschizon duboscqi Brasil, 1906  blastogregarine Siedleckia nematoides Caullery et Mesnil, 1898

The polychaetes Scoloplos armiger (Müller, 1776) (Polychaeta, Orbiniidae) were collected at the silty-sand littoral zone near the White Sea Biological Station of Lomonosov Moscow State University (Velikaja Salma, Kandalaksha Bay, White Sea, 66°33.190′ N, 33°06.550′ E) in Russia during the years 2014-2016. Both apicomplexan species were collected from the host intestine in Millipore (0.22 µm) filtered seawater using the entomological needles and subsequently transferred with a hand-drawn glass pipettes to embryo dishes with a 30 mm cavity for careful washing. Species were then separated. The host dissection and manipulation with parasites were performed using a stereomicroscope MBS-1 (LOMO, Russia). Protococcidian E. duboscqi was identified according to the original description by Brasil (1906) and later publication showing its life cycle stages by Chatton and Villeneuve (1936b). Blastogregarine S. nematoides was identified based on original descriptions by Caullery and Mesnil (1898).

4.1.2 Apicomplexan parasites from marine polychaete, Pygospio elegans

 archigregarine Selenidium pygospionis Paskerova et al. 2018  eugregarine Polyrhabdina sp.

The host organisms, polychaetes Pygospio elegans Claparède, 1863 (Polychaeta, Spionidae) were collected at the silty-sand intertidal zone near the White Sea Biological Station of Lomonosov Moscow State University (Velikaja Salma, Kandalaksha Bay,

White Sea, 66°33.190′ N, 33°06.550′ E) in Russia during the years 2014-2016. Prior to the dissection, the polychaetes were stored in small containers in the fridge at +10 °C with periodically changed seawater. Both the archigregarines and eugregarines were isolated from the host intestine in Millipore (0.22 µm) filtered seawater with the help of entomological needles. Subsequently, the parasites were transferred with a hand-drawn glass pipettes to embryo dishes with a 30 mm cavity for careful washing with filtered seawater and species separation. The host dissection and manipulation with parasites were performed using a stereomicroscope MBS-1 (LOMO, Russia). Individuals of archigregarine S. pygospionis were described as new species by Paskerova et al. (2018) according to morphological characteristics and phylogenetic analyses based on the SSU rDNA. Two manuscripts dealing with the description and motility of Polyrhabidna sp. are planned to be written in the near future (author of this thesis is co-author in one of them).

4.1.3 Apicomplexan parasites from marine polychaete, Travisia forbesii

 eugregarine Urospora travisiae Dogiel, 1910  eugregarine U. ovalis Dogiel, 1910

The host species Travisia forbesii Johnston, 1840 (Polychaeta, Opheliidae) was collected from upper sublittoral zone in the vicinity of White Sea Biological Station of Lomonosov Moscow State University (Velikaja Salma, Kandalaksha Bay, White Sea, 66°33.190′ N, 33°06.550′ E) in Russia and kept at temperature of +4 °C. After animal dissection, the brown bodies were collected from the host coelom with the help of entomological needles in Millipore (0.22 µm) filtered seawater using a stereomicroscope MBS-1 (LOMO, Russia). U. travisiae and U. ovalis were identified according to the original description by Dogiel (1910) and based on morphological and phylogenetical study published by Diakin et al. (2016).

4.1.4 Apicomplexan parasites from marine barnacle, Balanus balanus

 eugregarine Cephaloidophora cf. communis Mawrodiadi, 1908

Marine barnacles Balanus balanus Linnaeus, 1758 (Crustacea, Cirripedia), arctic boreal deep-water species, were collected during the years 2013-2015 from subtidal zone close

to the White Sea Biological Station of Lomonosov Moscow State University (Velikaja Salma, Kandalaksha Bay, White Sea, 66°33.190′ N, 33°06.550′ E) in Russia. Barnacles were afterwards dissected and eugregarines Cephaloidophora cf. communis were collected from host intestine using the entomological needles and hand-drawn glass pipettes under an MBS-1 stereomicroscope (LOMO, Russia). Eugregarines were transferred to embryo dishes with a 30 mm cavity and rinsed in Millipore (0.22 µm) filtered seawater for further procedures. Eugregarine specimens found in the examined barnacles were identified as C. cf. communis based on morphological characteristics described by Mawrodiadi (1908) and Simdyanov et al. (2015)1.

4.1.5 Apicomplexan parasites from desert locust, Schistocerca gregaria

 eugregarine Gregarina garnhami (Canning, 1956)

Host insects, Schistocerca gregaria (Forskål, 1775) (Orthoptera, Acrididae), were bred in colonies under laboratory conditions. After narcosis of insects with chloroform, followed by their decapitation and dissection, the eugregarines were isolated from locust mid-gut and caeca in Ringer’s saline solution using glass pipettes. Collected parasites were subsequently transferred to embryo dishes with a 30 mm cavity and were carefully washed from detritus and host remnants with Ringer’s saline solution. The manipulations and observations of the parasites were performed using an Olympus SZX7 stereomicroscope (Olympus, Japan). Trophozoites and gamonts of eugregarine G. garnhami were identified based on morphological descriptions published in studies by Canning (1956) and Valigurová and Koudela (2008).

4.1.6 Apicomplexan parasites from tropical cockroach, Blaberus dicoidalis

 eugregarine Blabericola (syn. Gregarina) cubensis (Peregrine, 1970)  eugregarine Protomagalhaensia granulosae Peregrine, 1970

1 The identification of Cephaloidophora cf. communis studied in Simdyanov et al. (2015) and Kováčiková et al. (2017) is questionable because individuals were collected from the different locality (White Sea) and the different host (Balanus balanus) than the species in original description. The first description was performed from Balanus improvisus, B. eburneus, and B. amphitrite from Black Sea by Mawrodiadi in 1908. However, gregarines isolated from B. balanus conformed in morphology to the original description of C. communis. 27

The host insects, Blaberus discoidalis Serville, 1838 (Dictyoptera, Blaberidae), were gained from laboratory bred colonies. Both of the eugregarine species were collected from mid-gut after host narcosis with chloroform, decapitation and dissection. The eugregarines were transferred by glass pipettes to embryo dishes with a 30 mm cavity for washing in Ringer’s saline solution and species separation under an Olympus SZX7 stereomicroscope (Olympus, Japan). Eugregarine Blabericola cubensis was identified based on characteristics described in original publication by Peregrine (1970) and revision of Blabericola spp. by Clopton (2012a). Protomagalhaensia granulosae was identified according to the description by Peregrine (1970) and redescription by Clopton (2012b).

Ringer’s saline solution (pH = 7.2):

H2O 1000 ml NaCl 7.5 g KCl 0.35 g

CaCl2 0.21 g

Table 1. Summarisation of investigated representatives of early emerging apicomplexans

Host Parasite location Host Parasite Locality References habitat within the host Eleutheroschizon duboscqi Brasil, 1906 White Sea, Valigurová et al., 2015 Scoloplos cf. armiger (Protococcidia,Eleutheroschizonidae) Kandalaksha Bay, intestinal lumen Müller, 1776 marine Siedleckia nematoides Caullery Russia (Polychaeta, Orbiniidae) Valigurová et al., 2017; et Mesnil, 1898 Simdyanov et al., 2018 (Blastogregarinida, Siedleckiidae) Orbinia (Syn. Aricia) latreilli Chattonaria mesnili Audouin & H. Milne Edwards, Chatton & Dehorne, 1929 English Channel marine intestinal lumen Simdyanov et al., 2018 1833 (Blastogregarinida, Chattonariidae) France (Polychaeta, Orbiniidae) Selenidium pygospionis Paskerova et al., 2018; Pygospio elegans Paskerova et al. 2018 White Sea, Kováčiková et al., 2019 Claparède, 1863 (Archigregarinida, Selenidiidae) marine Kandalaksha Bay, intestinal lumen (Polychaeta, Spionidae) Polyrhabdina sp. Russia material in preparation (Eugregarinida, Lecudinidae) Pherusa plumosa Selenidium pherusae White Sea, (Müller, 1776) Paskerova et al. 2018 marine Kandalaksha Bay, intestinal lumen Paskerova et al., 2018 Polychaeta, Flabelligeridae) (Archigregarinida, Selenidiidae) Russia Travisia forbesii Urospora travisiae Dogiel, 1910 White Sea, Diakin et al., 2018 Johnston, 1840 U. ovalis Dogiel, 1910 marine Kandalaksha Bay, coelom (article in preparation (Polychaeta, Opheliidae) (Eugregarinida, Urosporidae) Russia published in Ph.D. thesis) Cephaloidophora cf. communis Balanus balanus White Sea, Mawrodiadi, 1908 Linnaeus, 1758 marine Kandalaksha Bay, intestinal lumen Kováčiková et al., 2017 (Eugregarinida, (Crustacea, Cirripedia) Russia Cephaloidophoridae) Schistocerca gregaria Gregarina garnhami laboratory intestinal lumen (Forskål, 1775) (Canning, 1956) terrestrial Kováčiková et al., 2018 breeding (midgut and caeca) (Orthoptera, Acrididae) (Eugregarinida, Gregarinidae) Blabericola cubensis (Peregrine, 1970) Blaberus discoidalis (Eugregarinida, Blabericolidae) laboratory intestinal lumen Unpublished results Serville, 1838 terrestrial Protomagalhaensia granulosae breeding (midgut) (subchapter 5.2) (Dictyoptera, Blaberidae) Peregrine, 1970 (Eugregarinida, Blabericolidae)

4.2 Experimental assays and light microscopy

4.2.1 Experiments with cytoskeletal drugs

For experimental assays, parasites were divided equally into several embryo dishes with a 30 mm diameter cavity. The membrane-permeable cytoskeletal drugs were used to investigate the involvement of actin filaments and microtubules in parasites motility:  Jasplakinolide (JAS; Cat. No. J7473, Invitrogen, Czech Republic) - a cytotoxic natural reagent inducing actin polymerisation and stabilising existing actin filaments.  Cytochalasin D (Cat. No. PHZ1063, Invitrogen, Czech Republic) - a cell- permeable fungal metabolite binding to actin, causing the disruption of actin filaments and inhibiting the actin polymerisation.  Oryzalin (Cat. No. 36182, Sigma-Aldrich, Czech Republic) - a dinitroaniline herbicide depolymerising existing microtubules and prevents the polymerisation of new ones.  Colchicine (Cat. No. C3915, Sigma-Aldrich, Czech Republic) - a toxic alkaloid extracted from a plant (meadow saffron) inhibiting the polymerisation of

microtubules.

Jasplakinolide, cytochalasin D and oryzalin were reconstituted in dimethyl sulfoxide (DMSO) to prepare a 1 mM stock solution. Drugs were afterwards diluted in filtered seawater for experiments with marine apicomplexans or Ringer’s saline solution for gregarines from terrestrial hosts, to prepare a working solutions at the concentration 10 µM or 30 µM (alternatively 5 µM concentration was used). Colchicine was reconstituted directly in filtered seawater/Ringer’s saline solution to the concentration 10 and/or 100 mM. The initial concentrations of all drugs used in assays were set on the basis of experimental data published in Valigurová et al. (2013, 2017). Controls were performed in filtered seawater/Ringer’s saline solution with concentration of DMSO corresponding to the concentration of used drug. In set time intervals (depending on species and applied probe), a portion of parasites from each experimental assay was transferred to the microscopic glass slide for observations of their motility and video-/photo documentation using Leica DM 2000 light microscope connected to a DFC 420 digital camera (Leica Microsystems, Germany) or Olympus BX61 microscope equipped with a DP71 digital

camera (Olympus, Japan). Alternatively, gregarines were monitored under an Olympus SZX7 stereomicroscope (Olympus, Japan). At the end of each experiment, parasites from each assay were equally divided and fixed for further microscopic analyses (scanning and transmission electron microscopy, confocal laser scanning microscopy).

4.2.2 Mucus shedding in drug-treated eugregarines

Living trophozoites and gamonts of G. garnhami (around 50 cells for each experimental assay) were treated for 7 hours with 30µM JAS and 30µM cytochalasin D. Corresponding portion of control eugregarines was incubated in Ringer’s saline solution and 30 µM DMSO for the same period. Parasites were then transferred to the microscopic slides covered by a thin layer of microbiological LB agar (LB Broth with agar, Lennox; Sigma- Aldrich, Czech Republic) and slightly moistened with Ringer’s solution. The mucus shedding by gregarines gliding on agar was monitored under an Olympus BX51 microscope equipped with a DP70 digital camera, phase contrast and an ND25 filter (Olympus, Japan). The presence and the form of gliding path was documented and compared between controls and drug-treated eugregarines.

4.2.3 Experiments with artificial seawater with changed ionic composition

For study of potential effect of single ions constituting together the composition of seawater, an artificial seawater was prepared with laboratory available salts. Salinity of artificial seawater was set up to 25‰ according to White Sea salinity measured in Kandalaksha Bay. Archigregarines Selenidium pygospionis and eugregarines Polyrhabdina sp. were after washing in filtered natural seawater, put to the embryo dish with artificial seawater with ionic composition simulating the natural seawater (control) and to dishes containing artificial seawaters with changed concentration of ions. The pH of artificial seawater was in all cases adapted to 8.1. Saline composition of the White Sea was identified based on publication by Bruevitch (1960).

The composition of used artificial seawater (simulating the natural seawater):

H2O 1000 ml NaCl 18.78 g (321.36 mM) KCl 0.53 g (7.1 mM)

CaCl2 0.7 g (6.36 mM)

MgCl2.6H2O 4.35 g (21.4 mM)

MgSO4.7H2O 2.81 g (11.4 mM)

Four variants of seawater with changed ionic composition were prepared as follows: 1. Proportion and molarity of NaCl and KCl was exchanged (= the concentration of Na+ and K+ ions in final solution was affected): NaCl=0.42g/l (7.1 mM); KCl=23.96 g/l (321.36 mM); salinity = 24‰.

2. The amount of MgCl2.6H2O and MgSO4.7H2O was doubled (= the concentration of Mg2+ ions increased) in experimental solution compared to the control artificial

seawater: MgCl2.6H2O=8.7 g/l (42.8 mM); MgSO4.7H2O=5.62 g/l (22.8 mM); salinity = 27‰.

3. CaCl2 was omitted from final solution (= the concentration of Ca2+ ions decreased): salinity = 24‰.

4. The amount of CaCl2 was doubled (= the concentration of Ca2+ ions increased) in

experimental solution compared to the control artificial seawater: CaCl2 =1.4 g/l (12.72 mM); salinity = 25‰.

From each experimental assay portion of gregarines was transferred to the microscopic glass slide to observe and document their motility using a Leica DM 2000 light microscope connected to a DFC 420 digital camera (Leica Microsystems, Germany). Motility index was calculated in 25 individuals of S. pygospionis and 25 individuals of Polyrhabdina sp. from controls (natural and artificial seawater) and each experiment in set time intervals (30, 60, 240, 480 and 960 min) under an MBS-1 stereomicroscope (LOMO, Russia), according to King and Lee (1982):

number of motile gregarines in time t Motility index = x 100 number of motile gregarines in time 0

4.3 Electron microscopy

4.3.1 Transmission electron microscopy (TEM)2

Samples consisting of detached individuals and/or parasites attached to the host tissue were fixed in an ice bath in 2.5% (v/v) glutaraldehyde in cacodylate buffer (0.05 M; pH = 7.4) or in filtered seawater. To process marine specimens, the osmolality of the fixative was set to 750 mOsm by the addition of 1.28% sodium chloride (NaCl) (according to the actual salinity of the host habitat). Some specimens were fixed with 3% glutaraldehyde– ruthenium red [0.15% (w/v) stock solution in Milli-Q water] in cacodylate buffer (0.1 M; pH = 7.4) or with 2.5% glutaraldehyde–alcian blue [1% (w/v) stock solution in Milli-Q water] in filtered seawater. Gregarines from terrestrial hosts were fixed in 2.5% (v/v) glutaraldehyde in phosphate buffered saline (PBS). Fixed samples were then washed 3 x for 15-20 min and post-fixed in 1-2% (w/v)

OsO4 for 2 h in the same buffer as used for fixation and rinsed 3x for 15-20 min in the washing buffer (according to the fixative used). A portion of the samples fixed in 2.5% (v/v) glutaraldehyde in filtered seawater was rinsed 3 x for 15 min in cacodylate buffer

(0.2 M; pH = 7.4), then post-fixed in 1% (w/v) OsO4 for 2 h. Specimens were then dehydrated in an acetone or ethanol series (30%, 50%, 70%, 90% and 100%) and embedded in Epon (Polybed 812). Cell suspensions of some species were before dehydration procedure embedded in agar [2% (w/v) in distilled water; 62 °C]. Ultrathin sections were obtained with diamond knives using a Leica EM UC6 ultramicrotome (Leica Microsystems, Germany) and stained with uranyl acetate and lead citrate according to the standard protocols. Sections were examined under a TEM-1010 (JEOL, Japan).

4.3.2 Scanning electron microscopy (SEM)2

Specimens were fixed in 2.5-5% (v/v) glutaraldehyde in cacodylate buffer (0.2 M; pH = 7.4) or in PBS, washed 3 x for 15 min in the same buffer as used for fixation, post-fixed

2 The majority of samples was processed by staff of the Laboratory of Electron Microscopy, University of South Bohemia, Faculty of Science and Biology Centre of the Czech Academy of Sciences, Institute of Parasitology, České Budějovice, Czech Republic. Remaining samples were fixed and partially processed directly in field or laboratory by author. Semithin sections were prepared by author with glass knives using a Leica EM UC7 ultramicrotome (Leica Microsystems, Germany) and observed under an Olympus CX31 microscope (Olympus, Japan).

in 1-2% (w/v) OsO4 for 2 h in cacodylate buffer or in PBS, and finally washed 3 x for 15 min in the washing buffer (according to the fixative used). After dehydration in an acetone or ethanol series (30%, 50%, 70%, 90% and 100%), parasites were critical point-dried with CO2, coated with gold, and observed using a JSM-7401F (JEOL, Japan).

4.3.3 Freeze-etching (FE TEM)3

For freeze fracture, the suspensions of eugregarine B. cubensis gamonts or trophozoites attached to host tissue were fixed overnight in 2.77% (v/v) glutaraldehyde in 0.2 M cacodylate buffer (pH = 7.4), then washed for 1 h in PBS and saturated with 20% glycerol (w) overnight in a refrigerator for cryprotection. Prior to further processing, gregarines suspensions were concentrated on a clock glass and the dense pellet was placed using ceramic tweezers on a gold carrier. Parts of tissue with attached gregarines were transferred on a gold carrier by tweezer directly from the tube. Specimens were frozen in liquid nitrogen (-210 °C), mounted on a gold holder, and then processed in a BAF 060 freeze-etching system (BAL-TEC). Inside the chamber, the temperature was set from – 100 °C to –120 °C and the pressure was 10-5 Pa. Ice crystals from the surface of the sample were sublimated for 5-13 min, samples were fractured with a microtome knife and etched for 2-20 min. The surfaces of the fractured structures were coated with layers of platinum (2.4 nm at an angle 45°) and carbon (22.8 nm at an angle 90°). Afterwards, the specimens were removed from the BAF 060 device and melted at room temperature. Replicas were cleaned with 5% sodium hypochlorite (NaClO) and 70% sulfuric acid (H2SO4). Alternatively the rests of tissue from the replicas were remove by combination of 50-70% chromsulfuric acid (K2Cr2O7 +H2SO4 +H2O), 2-10% sodium hypochloride (NaClO) and 5-10% sulfuric acid. Finally, the specimens were washed in distilled water and transferred to copper grids. Replicas were examined using a Morgagni 268 D (FEI) transmission electron microscope. Statistical evaluation of intramembranous particles (IMP) per unit area (1 µm2) was performed in ImageJ software. The nomenclature follows terminology proposed in Branton et al. (1975) and used in Schrével et al. (1983) and Valigurová et al. (2013, 2017).

3 Freeze-etching procedure, preparation and statistical evaluation of replicas in publications included in this thesis were performed by Mgr. Naděžda Vaškovicová Ph.D. from Institute of Scientific Instruments of the Czech Academy of Sciences, Brno, Czech Republic. Author was partially involved in this methodology and in further evaluation of replicas by TEM. 34

4.4 Confocal laser scanning microscopy (CLSM)

Specimens were fixed for 45-60 min in freshly prepared 4% paraformaldehyde (PFA) in 0.1 M PBS at room temperature and then washed 3 x for 15 min in 0.1 M PBS before further processing. Parasites incubated with JAS were briefly rinsed before fixation to prevent the competitive inhibition of phalloidin binding. Specimens were transferred and stored in 0.01% sodium azide (NaN3) in PBS. Before staining procedure, samples were permeabilised with 0.3 or 0.5% Triton X-100 (Sigma-Aldrich, Czech Republic) in the period of 15-60 min, depending on species. The direct fluorescence of filamentous actin (F-actin) was performed using the phalloidin-tetramethylrhodamine B isothiocyanate (phalloidin-TRITC; Cat. No. P1951, Sigma-Aldrich, Czech Republic) wherein samples were incubated overnight at room temperature and then washed 3 x for 10 min in 0.1 M PBS. Controls were incubated using the same protocol but without phalloidin. For indirect immunofluorescence, the samples were incubated overnight at 4 °C with the following primary antibodies in PBS with 0.1% BSA:  mouse monoclonal IgG anti-actin raised against Dictyostelium actin, recognising the actin in Toxoplasma gondii and Plasmodium sp. (provided by Prof. Dominique Soldati-Favre, University of Geneva)  mouse monoclonal anti-α-tubulin, clone B-5-1-2 (dilution 1:1000; Cat. No. T5168, Sigma-Aldrich, Czech Republic)  rabbit anti-myosin (smooth and skeletal) (dilution 1:5; Cat. No. M7648, Sigma- Aldrich, Czech Republic),  rabbit anti-chicken spectrin (dilution 1:40; Cat. No. S1390, Sigma-Aldrich, Czech Republic),  rabbit anti-human spectrin (dilution 1:800; Cat. No. S1515, Sigma-Aldrich, Czech Republic). Specimens were afterwards washed 3 x for 10 min in 0.1 M PBS and incubated at 37 °C for 4 h with the following secondary antibodies in PBS with 1 % BSA:  FITC-conjugated anti-mouse (dilution 1:125; Cat. No. F1010, Sigma-Aldrich, Czech Republic)  FITC-conjugated anti-rabbit IgG (dilution 1:40; Cat. No. F9887 Sigma-Aldrich,

Czech Republic)  TRITC-conjugated anti-rabbit IgG (dilution 1:200; Cat. No. T6778, Sigma- Aldrich, Czech Republic) Samples were then washed again 3 x for 10 min in 0.1 M PBS. Controls were incubated with a secondary antibody alone, i.e. without primary antibody. Some specimens stained either for direct or indirect fluoresce, were counterstained to localise cell nuclei with 4′,6-diamidino-2-phenylindol (DAPI; Cat. No. D9542, Sigma- Aldrich, Czech Republic) for 10 min or with Hoechst 33342 (Cat. No. H1399, Molecular probes, Czech Republic) for 1 h and then washed 3 x for 10 min in 0.1 M PBS. Preparations were mounted in VECTASHIELD Hard Set Mounting Medium (Cat. No. H- 1400; Vector laboratories, USA) and examined under an Olympus IX81 FVBF-2 microscope, equipped with a laser-scanning Fluo View 500 confocal unit (Fluo View 3.4 software; Olympus, Japan) and DP70 digital camera. Fluorescence was visualised using TRITC (phalloidin/544 nm), FITC (anti-actin, anti-α-tubulin/457-515 nm) and UV (DAPI, Hoechst/405 nm) lasers sets. Micrographs of all specimens (= particular staining and controls) were obtained under identical image capture conditions (filters, laser intensity). Some micrographs were processed using Fiji software (an image-processing package based on ImageJ developed at the National Institutes of Health).

4.5 Western blot

Western blot analysis required a large amount of examined material, therefore only eugregarines from laboratory bred insect hosts were used. Either gamonts/syzygies or mature gametocysts with oocysts of G. garnhami, B. cubensis and P. granulosae were collected. Samples were washed at least 3 x in Ringer’s saline solution containing the antibiotics (penicillin and streptomycin, or ampicillin). Afterwards, liquid was removed by mechanical pipette and gregarines were frozen at -80 °C overnight. For proteins isolation, a determined volume of 0,1 mM Tris (depending on the amount of sample) with protease inhibitor cocktail (Cat. No. 05892791001; Roche: cOmplete ULTRA Tablets, Mini, EASYpack) was added to the sample. The cells were subsequently either mechanically processed by plastic homogenisator or were damaged by repetition of heat (37 °C) and cold (-80 °C) shocks. Alternatively, eugregarines were broken by sonication (power 30/ pulse 30/ 1 min - repetition 3x) with or without previous freezing procedure. After centrifugation (10 min/4 °C/13 200 rpm) supernatant was

separated from the pellet and transferred to a new tube. Pellet was dissolved in 0.1 mM Tris containing 1 % SDS and protease inhibitor. Before electrophoresis, the samples were kept for 5-10 min on 100 °C in thermostat. Specimens were mixed with 8 µl Laemmli protein sample buffer (Cat. No. 1610747; Bio- Rad, Czech Republic) and loaded into 4-15% Mini-PROTEAN TGX™ precast protein gels (Cat. No. 4561083; Bio-Rad) with molecular weight (ladder) Spectra Multicolor Broad Range Protein Ladder (Cat. No. 26634; Thermo Scientific). Electrophoresis was performed in tris-glycine-SDS (TGS) running buffer and set up to 120V for 60 min. After electrophoresis, the gel was immersed to commasie blue solution for protein staining and let on shaker at room temperature overnight. The gel was then washed 3 x in commmasie blue destaining solution.

Commasie blue staining solution: MeOH 250 ml ddH2O 200 ml Acetic acid 50 ml Commasie blue 0.5 g

Destaining solution: MeOH 250 ml

ddH2O 200 ml Acetic acid 50 ml

Semi-dry blotting in Bio-Rad Trans-Blot turbo Transfer system – 690BR010832 was used to transfer proteins from the gel to a nitrocellulose membrane. Blotting sandwich was prepared from two extra thick Wathmann papers (Cat. No. 1703965; Bio-rad, Czech Republic) dipped in Towbin blotting buffer, between which the gel containing samples and molecular ladder, and nitrocellulose membrane (previously activated by distilled water and afterwards incubated for 5 min in Towbin buffer) were inserted. Semi-dry transfer was set up for 7-10 min on 25V.

Towbin (blotting) buffer: ddH2O 280 ml 10X Tris + Glycin 40 ml MeOH 80 ml 10% SDS 0.8 ml

Before further processing of the nitrocellulose membrane with selected antibodies, the ladder was cut off from the membrane. Afterwards the membrane was transferred to a clean box with blocking solution composed of PBS buffer containing 0.1% Tween 20 (PBST) and 3% or 5% BSA. The membranes were then incubated in blocking buffer on shaker overnight at 4 °C to prevent the nonspecific binding of antibodies. The blocking buffer was removed and a buffer containing PBST with 1% or 3% BSA, and primary antibody was added. For western blot analysis, the following primary antibodies were used:  mouse monoclonal IgG anti-actin raised against Dictyostelium actin, recognising actin in Toxoplasma gondii and Plasmodium sp. (provided by Prof. Dominique Soldati-Favre, University of Geneva)  mouse monoclonal anti-α-tubulin, clone B-5-1-2 (dilution 1:1000; Cat. No. T5168, Sigma-Aldrich, Czech Republic)  rabbit anti-chicken spectrin (dilution 1:800; Cat. No. S1390, Sigma-Aldrich, Czech Republic)  rabbit anti-human spectrin (dilution 1:800; Cat. No. S1515, Sigma-Aldrich, Czech Republic)  mouse anti-myosin antibody [C5C.S2] (dilution 1:2500; Cat. No. AB24648, Abcam, Czech Republic)

Samples were incubated with primary antibodies for 2 h at room temperature and then washed 3 x for 10 min in PBST, both on shaker. After careful washing, the membranes were dipped to PBST with 1% or 3% BSA and containing the following secondary antibodies:  goat anti-mouse IgG (GAM) (dilution 1:5000; Cat. No. 31320, Thermo Scientific)  goat anti-rabbit IgG (GAR) (dilution 1:5000; Cat. No. 31340, Thermo Scientific)

The membranes were incubated with secondary antibodies on shaker for 1.5 – 2 h at room temperature then washed 3 x for 10 min in PBST and transferred to reveal buffer for next 10 min.

Reveal buffer (pH=9): ddH2O 100 ml NaCl 22.25 g Tris-HCl 30.25 g

MgCl2.6H2O 2.53 g

Finally, the membranes were covered with 10 ml of reveal buffer containing 66 µl of nitro-blue tetrazolium chloride (NBT) and 33 µl of 5-bromo-4-chloro-3'- indolyphosphate p-toluidine salt (BCIP) and left to reveal as long as necessary. Providing, all current bands were visualised, process of revealing was stopped by a transfer of the membrane to distilled water at least for 10 min.

4.6 Molecular analysis

For DNA isolation, MasterPureTM Complete DNA and RNA Purification Kit (Cat. No. MC85200 and MC89010; Epicentre an Illumina company, USA) was used. Collected gregarine trophozoites and/or gamonts (S. pygospionis, Polyrhabdina sp. and G. garnhami) or mature gametocysts with oocysts (G. garnhami, B. cubensis) were carefully washed from host debris and transferred to T&C Lysis solution. Alternatively, the specimen were fixed in 96% ethanol or RNA later, washed 2 x with T&C Lysis solution and then transferred to 300 µl T&C Lysis solution and processed according to kit protocol. Sequences of SSU rDNA, actin and α-tubulin from investigated species were amplified using a total volume of 25 µl PCR sample with Taq DNA Polymerase (Cat. No. 10342020; Thermo Scientific, Czech Republic) kits and following primers:

SSU (Q5-Q39Api): Forward: 5'-GTATCTGGTTGATCCTGCCAGT-3' Reverse: 5'-GATCCTTCTGCAGGTTCACCTAC-3'

Actin 1 (ActF5-ActR5) (according to Leander and Keeling, 2004): Forward: 5'-GAGAAGATGACNCARATHATGTTYGA-3' Reverse: 5'-GGCCTGGAARCAYTTNCGTRGNAC-3'

Actin 2 (actR-actS) (designed by Prof. Isabelle Florent): Forward: 5'-AAGGCWGGWKTGGCCGGNGATGCNCC-3' Reverse: 5'-TCNTCGTAYTCYTTKGTKATCCACAT-3'

α-tubulin (atubF-atubR) (designed by Prof. Isabelle Florent): Forward: 5'-GGGAGCTNTWYTGYYTNGARCA-3' Reverse: 5'-GCATNCCYTCNCCNACRTACCA-3'

The amplification process in thermocycler, starting at 95 °C for 2-5 min for the initial denaturation, was followed by 35-39 cycles of denaturation at 95 °C for 40-45 s, anneal at 45-54 °C for 40-45 s, extension at 72 °C for 2 min and final extension at 72 °C for 7-10 min. PCR products with amplified fragments were loaded to 0.5 or 1% agarose gel with GeneRulerTM 1kb DNA Ladder (Cat. No. SM0312; Thermo Scientific) for electrophoresis (30-50 min, 50 V) and examined using Syngene G:Box.

5 Results

5.1 Published data

The data presented in this subchapter are taken from manuscripts that have already been published or accepted (in press) in scientific journals. The papers are listed chronologically.

Paper I Valigurová A., Paskerova G.G., Diakin A., Kováčiková M., Simdyanov T.G. (2015). Protococcidian Eleutheroschizon duboscqi, an unusual apicomplexan interconnecting gregarines and cryptosporidia. PLoS One 10 (4): e0125063.

The attachment strategy, subcellular structure and host-parasite interactions of the protococcidian Eleutheroschizon duboscqi, parasitising the intestine of marine polychaete Scoloplos armiger were studied using the light, electron and confocal microscopic methods. Epicellularly developing stages of E. duboscqi were covered by a host cell- derived, two-membrane parasitophorous sac forming a caudal tipped appendage (tail). The parasite pellicle was organised in broad folds with grooves in between and covered by a dense fibrous glycocalyx coat. The attachment apparatus of E. duboscqi consisted of lobes arranged in one (trophozoites) or two (gamonts) circles, crowned by a ring of filamentous fascicles. The pellicle of E. duboscqi also appears unique, in that it seems to re-build or reorganise during the parasite development. Detached parasites were enveloped by a parasitophorous sac covering their distal area above the attachment site, hereby suggesting that after detachment from the host tissue they preserve this envelope of host origin, providing them ongoing protection in a potentially unfriendly surrounding environment. For more accurate visualisation of cytoskeletal elements parasites were incubated with cytoskeletal probes (jasplakinolide, cytochalasin D and oryzalin at a concentration 10 and 30 µM). Confocal laser scanning microscopic observations confirmed the presence of actin and tubulin polymerised forms in both the parasitophorous sac and the parasite, while the myosin labelling was restricted to the sac only. Spectrin appeared to be dispersed in low concentrations in the parasite cytoplasm, while the tail of parasitophorous sac stained with a strong intensity, suggesting the accumulation of plasma membrane proteins in this zone. While the earliest developmental stages attached to host tissue had typical subpellicular microtubules, during maturation, the trophozoites underwent changes

leading to a loss of subpellicular microtubules in mature trophozoites and later endogenous stages. Nevertheless, the preservation of tubulin either in its non-polymerised form or its presence in other tubulin-rich structures was confirmed by positive α-tubulin labelling of the parasite surface and cytoplasm. The second option appears more likely, as the incubation with oryzalin resulted in the vanishing of fluorescent labelling from the parasite periphery (the putatively unpolymerised α-tubulin was more dispersed throughout the cytoplasm) along with a frequent detachment of parasites with their sacs from host epithelium. Positive labelling of the parasite surface for filamentous actin supported by the effect of actin-modifying drugs on its staining pattern suggest that longitudinally oriented filaments, located beneath the parasite IMC are actin rich and play the role of the parasite cytoskeleton instead of subpellicular microtubules lacking in mature stages. Importantly, the attachment strategy of E. duboscqi shares features with that of cryptosporidians and gregarines, i.e. the parasite itself conspicuously resembles an epicellularly located gregarine, while the parasitophorous sac develops in a similar manner to that in cryptosporidians. Detailed re-evaluation of epicellular development in other apicomplexans showed that E. duboscqi also shares similarities with certain eimeriid coccidians from poikilotherms. All these parasites stimulate additional growth and fusion of host cell microvilli along with modifications to the host cell plasma membrane, leading to a formation of parasitophorous sac, hereby enclosing them in the cavity of a host- derived envelope that separates them from the host internal environment.

Paper II Kováčiková M., Simdyanov T.G., Diakin A., Valigurová A. (2017): Structures related to attachment and motility in the marine eugregarine Cephaloidophora cf. communis (Apicomplexa). European Journal of Protistology 59: 1-13.

Present study deals with ultrastructural and immunological analyses of structures that are expected to be involved in parasite attachment to host tissue and the unique motility mode displayed by the trophozoites and gamonts of eugregarine Cephaloidophora cf. communis, parasite of the barnacle Balanus balanus (Crustacea, Cirripedia). The attachment apparatus, the epimerite, possessed numerous irregularly distributed pores in its plasma membrane, supposed to release adhesives presumably produced by the microneme-like structures found in the eugregarine apical end and facilitating stronger attachment to the host tissue (i.e. represented the chemical way of attachment strategy). In addition, the tiny 42

long protrusion rising from the epimerite centre observed in some SEM micrographs, likely ensure a more stable, mechanical anchoring to host intestine. The epimerite was separated from the protomerite by a septum consisting of tubulin-rich filamentous structures. Another accumulation of α-tubulin, visible in some parasites after immunofluorescent labelling, was detected in a form of a funnel-like structure extending from the epimerite centre and ending at the septum. Detached trophozoites and gamonts showed highly active gliding motility, enriched by jumping and rotational movements with rapid changes in gliding direction. Moreover, a flexion in the area of the septum separating the protomerite from the deutomerite and in the first third of the deutomerite was observed during their gliding. In some cases, the protomerite partially retracted into the deutomerite. Reverse movement was also observed. Florescent phalloidin labelling revealed the presence of filamentous actin dispersed throughout the entire eugregarine with a slightly increasing intensity around both septa and in the area of the nucleus. In addition, specific antibody labelling confirmed the presence of actin forming abundant clusters in the eugregarine cytoplasm, occurring especially in the epimerite-protomerite region. Myosin was restricted exclusively to the gregarine cortex with an organisation following the pattern of longitudinal epicytic folds. Moreover, clusters of putatively unpolymerised α-tubulin were dispersed in the cytoplasm of some individuals. Despite the presence of the basic motor proteins in C. cf. communis, i.e. actin (prevailing in polymerised form) and myosin, the exact position of actin microfilaments as well as the role of both these proteins in motility of marine eugregarines remain to be elucidated. Nevertheless, the so far obtained data indicate that the motility in C. cf. communis differs from the substrate-dependent gliding described for apicomplexan zoites. This outcome is further supported by the absence of subpellicular microtubules, which are considered essential components in apicomplexan zoite motor machinery. Furthermore, significant changes in cell shape during the gliding locomotion indicate that additional structures must be involved in motility of C. cf. communis trophozoites and gamonts. The unique architecture of epicytic folds (comprising two 12-nm filaments and internal lamina with an extraordinary dentate appearance) altogether with an intensive secretion of mucopolysaccharides densely coating the entire gregarine surface, are proposed as auxiliary elements responsible for the variable modes of gliding motility in this eugregarine.

Paper III Valigurová A., Vaškovicová N., Diakin A., Paskerova G.G., Simdyanov T.G., Kováčiková M. (2017): Motility in blastogregarines (Apicomplexa): Native and drug- induced organisation of Siedleckia nematoides cytoskeletal elements. PLoS One 12 (6): e0179709.

Based on experimental data evaluated by a combined microscopic approach, this study focuses on the motility and cytoskeleton of the blastogregarine Siedleckia nematoides parasitising the intestine of marine polychaete Scoloplos armiger. Parasites moved independently on a solid substrate and showed no signs of gliding motility. While the pendular movement was only barely visible in the earliest trophozoite stages of S. nematoides, it was recognisably observed in maturing trophozoites. Mature trophozoites and gamonts exhibited highly active pendular/twisting motility with typically waves originating in the proximal region of the cell (just behind the attachment area) and proceeding to its distal end, while the last third of the cell appeared more rigid with a limited mobility. Experimental motility assays using cytoskeletal drugs affecting the de- /polymerisation of cytoskeletal proteins revealed that the subpellicular microtubules, organised in several layers under the blastogregarine pellicle, appear to be the leading motor structures. The treatment with microtubule destroying agents (10 mM and 100 mM colchicine or 10 µM and 30 µM oryzalin) confirmed the gradual disruption of microtubules, correlating with the increasing drug concentrations and prolongation of the incubation period and resulted in the complete blocking of parasite motility. The drug- induced cell rigidity first appeared in the posterior half of the parasite (similar to that observed in Selenidium archigregarines). Ultrathin sections confirmed complete loss of a portion of microtubules from the otherwise continuous outermost microtubule layer, corresponding to an overall decrease of the fluorescent signal for α-tubulin labelling observed under CLSM. The phalloidin labelling confirmed the presence of actin filaments in the S. nematoides cortex and cytoplasm and showed that the majority of actin was already present in a polymerised form and appeared to be located beneath the inner membrane complex. Incubation with drugs influencing actin polymerisation (10 µM and 30 µM JAS or 10 µM and 30 µM cytochalasin D) caused gradual motility cessation and amplification of the fluorescence signal in JAS-treated blastogregarines labelled by phalloidin. In addition, the filamentous structures associated with subpellicular microtubules observed in ultrathin sections of control blastogregarines (i.e. cross-linking 44

protein complexes consisting of proteins embedded in the IMC and the network around subpellicular microtubules) reacted to the JAS and cytochalasin D treatment, resulting in changes in spacing of subpellicular microtubules, hereby indicating their actin nature. As already proposed for motility in Selenidium archigregarine, data presented herein suggest that the subpellicular microtubules organised in several layers are the real leading motor structures. If the axoneme-like sliding mechanism of microtubules is applicable for motility of S. nematoides, the putative actin cytoskeleton (located beneath the IMC) might associate lengthwise with subpellicular microtubules to position them. Otherwise, the actin filaments may force the synchronised bending of microtubules in some cell regions and this way generate the undulating motility. Despite the presence of key glideosome components such as three-layered apicomplexan pellicle, actin (including its filamentous form), myosin restricted to the cell cortex, subpellicular microtubules, numerous micronemes and prominent glycocalyx layer (where adhesins might be located), the motility mechanism of S. nematoides differs from the glideosome machinery. Nevertheless, experimental assays using cytoskeletal probes proved that the polymerised forms of actin and tubulin (forming subpellicular microtubules) play an essential role in the S. nematoides movement.

Paper IV Simdyanov T.G., Paskerova G.G., Valigurová A., Diakin A., Kováčiková M., Schrével, J., Gillou L., Dobrovolskij A.A., Aleoshin V.V. (2018): First ultrastructural and molecular phylogenetic evidence from the blastogregarines, an early branching lineage of plesiomorphic Apicomplexa. Protist 169 (5): 697-726.

The affiliation of blastogregarines has been for a long time uncertain and they were considered as an intermediate apicomplexan lineage between gregarines and coccidians, or an isolated group of eukaryotes altogether. In the present study we report the ultrastructure of two blastogregarines, Siedleckia nematoides and Chattonaria mesnili, and provide the first molecular data on their phylogeny based on SSU, 5.8S, and LSU rDNA sequences. Morphological analysis revealed plesiomorphic status of blastogregarines, which possess both the gregarine and coccidian characteristics. Blastogregarines share many ultrastructural features especially with archigregarines – they possess longitudinally folded or smooth pellicle lined by longitudinal subpellicular microtubules. Also, mucron with apical complex, dedicated for myzocytotic 45

feeding, does not disappear in the trophozoite stage in both groups. These traits shared with archigregarines likely represent the ancestral states of the corresponding cell structures for Apicomplexa. The mucronal complex of S. nematoides is well developed and equipped with apical organelles, the conoid, internal and external polar rings, numerous rhoptries and putative micronemes, and a mucronal vacuole. The mucron of C. mesnili is strongly modified lacking the conoid and rhoptries. Subpellicular microtubules is S. nematoides lying just beneath the three-layered pellicle (consisting of the plasma membrane with a well-developed glycocalyx, and the inner membrane complex) were arranged regularly in a single layer with few additional microtubules located deeper within the cytoplasm. These microtubules arose from the anterior polar ring, which represent microtubule-organising center (MTOC, giving the rise to the subpellicular microtubules) and passed along the entire cell length. The typical feature of blastogregarines is bending motility (similar to that observed in archigregarines); S. nematoides performed highly active bending, twisting, and squirming movements in comparison to the weak motility with slow and intermittent bending observed in C. mesnili. These differences in motility can be attributed to specific cortex modifications. According to molecular phylogenetic analyses performed in this study, blastogregarines represent a separate class of Apicomplexa as an independent, early diverging lineage.

Paper V

Paskerova G.G., Miroliubova T.S., Diakin A., Kováčiková M., Valigurová A., Gillou L., Aleoshin V.V., Simdyanov T.G. (2018): Fine structure and molecular phylogeny of two marine gregarines, Selenidium pygospionis n. sp. and S. pherusae n. sp., with notes on the phylogeny of Archigregarinida (Apicomplexa). Protist 169 (6): 826–852.

The morphological, ultrastructural, and molecular phylogenetic evidence from two archigregarine species: Selenidium pygospionis sp. n. and S. pherusae sp. n., parasitising intestine of marine polychaetes were reported in this study. Both species exhibited the typical features of archigregarines, which are considered a key group for understanding the early evolution of Apicomplexa. The phylogenetic analyses based on LSU (28S) rDNA and near-complete ribosomal operon (concatenated SSU, 5.8S, LSU rDNAs) sequences were performed, including S. pygospionis sequences. Although being preliminary, these phylogenetic analyses revealed gregarines as a monophyletic group.

Available SSU (18S) rDNA sequences of archigregarines formed four major lineages fitting the taxonomical affiliations of their hosts, but not the morphological/biological features used for the taxonomical revision by Levine (1971). The SSU rDNA-based phylogenies showed archigregarines as a paraphyletic group, although their deep branching remained unresolved and their monophyly was not rejected by topology tests. Despite their molecular-phylogenetic heterogeneity, subcellular organisation of archigregarines exhibits extremely conservative plesiomorphic features. Interestingly, intracellular stages of S. pygospionis localised within parasitophorous vacuoles were found in the epithelium of host intestine. This observation may point to the initial intracellular localisation in the earliest phase of trophozoite development. Both studied species of Selenidium move by forming one (S. pherusae) or up to four (S. pygospionis) bending sections along their body but never contract. Observations on flattened trophozoites of S. pygospionis revealed that their bending motility is generated only in one cell plane (the flattened sides act antagonistically in the bending sections forming in a single plane of the cell), therefore it was proposed to refer to this type of motility as a nematode-like bending (postulating that the three-membrane pellicle and the longitudinal subpellicular microtubules are skeletal and motile units representing together a unicellular analogue of the musculocuticular system of nematodes). Observed motility considerably differs from the bending motility exhibited by other Selenidium spp., where bends generated in different cell planes, are often combine with contraction and twisting of the cell in different cell sections. The dynamic motility of studied archigregarines presumably correlates with the number of mitochondria located directly beneath the subpellicular microtubules. To note, the intracellular axial streak extending from the anterior end to the posterior end and forming an expansion around the nucleus, together with putative microfilaments may also be involved in Selenidium spp. motility. Based on the presence of a series of connected vacuoles arranged along the S. pygospionis cell axis and around the nucleus, was proposed that the axial streak is a system of vacuoles (possibly digestive) originating in the mucron during the myzocytotic feeding and transporting nutrients from the anterior to the posterior end of archigregarine.

Paper VI Kováčiková M., Vaškovicová N., Nebesářová J., Valigurová A. (2018): Effect of jasplakinolide and cytochalasin D on cortical elements involved in the gliding motility of the eugregarine Gregarina garnhami (Apicomplexa). European Journal of Protistology 66: 97-114.

Eugregarine Gregarina garnhami, parasitising the digestive tract of the desert locust Schistocerca gregaria (Orthoptera, Acrididae), was investigated to determine the involvement of cytoskeletal elements (the ectoplasmic network and myonemes) and the secretion of mucosubstances during its gliding motility. A freeze-etching analysis of pellicle membranes revealed the size, density, and arrangement of intramembranous particles along with the distribution and size of numerous pores and ducts found to interrupt the pellicle. Electron and confocal microscopic analyses of eugregarines treated with actin-modifying drugs (30 µM JAS and 30 µM cytochalasin D) showed the importance of the myonemes (occurring at the border between the ectoplasm and endoplasm) and ectoplasmic network (situated between the myonemes and the IMC) in G. garnhami motility. After the application of 30 µM JAS the gamonts were able to glide until the end of experiment (7 h) without obvious cell deformations or gliding deceleration. The dense ectoplasmic network and bundles of annular myonemes were detected under the pellicle of JAS-treated eugregarines in ultrathin sections. Intensive labelling of F-actin with phalloidin revealed that numerous actin filaments running perpendicular to the longitudinal cell axis, close to each other and linked lengthwise, correspond in their organisation and localisation to the myonemes detected in ultrathin sections. We attribute the weak effect of JAS on G. garnhami gamonts to the assumption that the majority of actin in this species is already present in polymerised form (even before incubation with the actin stabilising JAS). In contrast, application of 30 µM cytochalasin D induced the vanishing (less evident presence) of myonemes and ectoplasmic network, resulting in eugregarines motility cessation almost immediately after drug application. After washing this drug out, the gliding motility recovered, confirming that the cytochalasin D-treated eugregarines were alive, but had suffered complete immobility. Lower cytochalasin D concentrations of 5 and 10 µM, applied to compare the effect of various drug concentrations on G. garnhami motility, caused no significant changes. The gliding of drug-treated eugregarines on agar was examined to verify the presence of secreted mucus trail. Clearly evident long and regular mucous paths 48

left behind gliding gamonts were seen in all experimental groups; however, the paths of drug-treated gregarines were less noticeable. All microscopic and experimental data indicate that annular myonemes consist of bundles of actin filaments. The organisation and density of myonemes and ectoplasmic network, changing during the experiments with actin-modifying drugs in accordance with gamonts gliding activity, suggest that these structures might serve as essential contractile elements facilitating the gliding motility in G. garnhami. Moreover, this study indicates that the dynamic process of actin polymerisation and subsequent rapid depolymerisation, proposed for glideosome-based gliding of apicomplexan zoites, is not essential for gliding motility in studied eugregarine. In contrast, only the polymerised form of actin seems to be the main leading motor structure, as showed the treatment with JAS, not significantly affecting the motility of G. garnhami. On the contrary, treatment with cytochalasin D almost immediately blocked eugregarine motility due to depolymerisation of existing actin filaments. The contraction of myonemes along with more compact ectoplasmic network likely caused that superficially protruding superfolds grouped together on one site of gregarine and exhibited a denser secretion and accumulation of mucus, suggesting this part to be the eugregarine gliding side.

Paper VII Kováčiková M., Paskerova G.G., Diakin A., Simdyanov T.G., Vaškovicová N., Valigurová A. (2019): Motility and cytoskeletal organisation in the archigregarine Selenidium pygospionis (Apicomplexa): observations on native and experimentally affected parasites. Parasitology Research 'in press'.

The study focuses on the movement of archigregarine Selenidium pygospionis, a parasite from the intestine of Pygospio elegans (Polychaeta, Spionidae). A combination of light, electron, and confocal laser scanning microscopy, supplemented by experimental motility assays with cytoskeletal probes applied on living S. pygospionis trophozoites and gamonts, enabled to verify the fundamental role of actin and tubulin in the archigregarine motility. Archigregarines detached from the host tissue exhibited a regular bending or nematode-like movement with the beating frequency of 0.35 ± 0.04 beats/s (with the beat to beat interval of 2.87 ± 0.33 s). The three-layered pellicle, consisting of a plasma membrane underlain by two cortical cytomembranes (the IMC), was coated by a thin

glycocalyx layer. The cytoplasm was differentiated into an ectoplasm, a narrow cortical region containing the subpellicular microtubules (21.9 ± 0.1 nm in diameter) and various vesicles, and a granular endoplasm with large nucleus and cytoplasmic inclusions. The subpellicular microtubules were divided into an outermost continuous set and additional clusters of single microtubules located deeper within the ectoplasm. An electron-lucent hexagonal sheath (35.3 ± 0.1 nm in diameter) surrounded each microtubule. The distance between two adjacent microtubules was 38 ± 0.3 nm, and the number of microtubules was 26.6 ± 0.4 per 1 µm of pellicle length in control parasites incubated in seawater. Based on experimental assays using the cytoskeletal drugs affecting the polymerisation of tubulin (10 and 30 µM oryzalin and 100 mM colchicine) and actin (30 µM JAS and 30 µM cytochalasin D), two main elements responsible for bending motility in S. pygospionis were identified. The subpellicular microtubules appear to be the main motor component. The motility of most archigregarines treated with 10 µM oryzalin ceased after 13 hours, with significant reduction in the number of subpellicular microtubules in their outermost layer (14.9 ± 1.3 per 1 µm of pellicle length). Immunofluorescent analysis showed a complete destruction of microtubules and formation of noticeable α-tubulin clusters under the pellicle. In colchicine-treated S. pygospionis, the rigidity of archigregarines (staring at posterior end and proceeding gradually anteriorly along the cell) caused the complete movement stoppage in 90 minutes, suggesting that microtubule depolymerisation begins in the caudal region and continues towards the apical polar ring (= MTOC, where microtubules initially originate). Though the commonly continuous outermost microtubular layer of colchicine-treated S. pygospionis showed some empty regions, the number of microtubules differed only moderately from controls because the remnants of microtubules were often still preserved in the subpellicular layer. The fluorescent staining of colchicine-treated parasites revealed the preservation of the longitudinal arrangement of α-tubulin corresponding to microtubules, which were, however, interrupted and small tubulin clusters had formed. The experiments with actin-modifying drugs demonstrated the role of actin filaments in S. pygospionis motility; changes in motility and its blocking occurred along with an increase in polymerised actin, indicating that actin filaments rather act as a supportive component for subpellicular microtubules. The movement of JAS- treated archigregarines ceased in most individuals after 9 hours, while cytochalasin D suppressed the motility but did not stop it. Organisation of cytoskeletal elements did not show significant changes compared to controls at ultrastructural level. However, the phalloidin labelling revealed fluorescence signal multiplication in JAS-treated

archigregarines, while those incubated with cytochalasin D showed only very weak signal. Despite the morphological similarity of archigregarine trophozoites (considered to be overgrown apicomplexan zoites = hypersporozoites) to apicomplexan zoites, the bending motility of S. pygospionis obviously differs from their substrate-dependent gliding facilitated by the glideosome. We believe that this bending motility predominates in Selenidium archigregarines and most likely replaces the substrate-dependent gliding during transformation of their sporozoite into the trophozoite stage.

5.2 Unpublished data

First subchapter includes the summary accompanied with figure plates taken from prepared manuscript dealing with the morphology of urosporid eugregarines. Following subchapters include the partial data from experiments testing the influence of specific ionic composition of seawater on motility of marine gregarines, and results of Western blot analyses together with related morphological observations on eugregarines from laboratory insect colonies that were not included in publications. Next subchapters deal with so far unpublished data obtained on marine eugregarine Polyrhabdina sp. and with evaluation of molecular methods performed by the thesis author. In addition, it is important to note that motility research is largely dependent on video documentation, which represents a significant part of this work, but could not be included in presented thesis.

5.2.1 Manuscript in preparation

Diakin A., Vaškovicová N., Paskerova G.G., Kováčiková M., Nebesářová J., Valigurová A. Gametocysts, oocysts and sporozoites morphology of urosporid gregarines with remarks on molecular phylogeny as inferred from rDNAs. A manuscript draft published In Diakin A. (2018): Biology of marine early emerging apicomplexans. Ph.D. thesis, Masaryk University, Brno, p. 106-130.

Presented study focuses on brown bodies, occurring in the body cavity of marine polychaete Travisia forbesii, and forming as a result of immune reaction of host coelomocytes, which incorporated gametocysts produced by the coelomic gregarines Urospora ovalis and/or U. travisiae. Under the gametocyst envelope, a great number of

spindle shaped heteropolar oocysts could be observed. With a high probability, investigated oocysts belong to species U. travisiae. Under the SEM and on freeze-etching replicas evaluated under TEM, the outer surface of these oocysts appeared smooth or fine- grained. The oocyst wall was built of two envelopes, an 8 nm thick exospore and 30-45 nm thick endospore. One pole of the oocyst bore a long thin, narrowing tail that was either straight or curved. The opposite pole of the oocyst was equipped by a transparent cylindrical to conical funnel, presumably dedicated for excystation of mature sporozoites. Under the oocysts wall, the residual body and 4-8 sporozoites could be found. The spindle/banana-shaped sporozoites were organised in four at both poles inside the oocyst, with their apical ends oriented towards the oocyst poles and posterior ends interdigitated in the middle region of the oocyst. They were covered with a typical three-layered apicomplexan pellicle consisting of the plasma membrane and the IMC, underlain by 20 longitudinal subpellicular microtubules arising from an apical polar ring and proceeding towards the posterior end of sporozoite. The large roundish nucleus, localised in the middle of the cell or slightly posteriorly, often occupied the entire transversal section and showed the characteristics typical for apicomplexans zoites. The anterior third of sporozoite cytoplasm was packed with specialised secretory organelles, several club- shaped rhoptries and numerous micronemes. Phylogenetic analysis of the LSU and SSU+LSU sequences showed that Urospora spp. formed a robust long branch with a basal position to the clades comprising crustacean gregarines, eugregarines of the genus Gregarina, and Ancora sgittata.

Figure 5. Morphology of brown bodies from coelom of Travisia forbesii. A. General view of brown body. LM. B. Histological section of brown body. H&E, LM. C. Crashed brown body full of oocysts. SEM. D. Transversal section of coelomocytes layer surrounding gametocyst. TEM. E. Fracture of coelomocytes layer. FE TEM. g – gametocyst, h – host cells, oo – oocysts.

Figure 6. Morphology of gametocyst wall. A. General view of gametocyst. DIC. B-C. Cross-section through the lamellar structures of the gametocyst wall. TEM. D. Layer of coelomocytes and gametocyst at lower magnification. Black rectangles demark position of Fig. 2F and Fig. 2G. TEM. E. Fractured gametocyst with oocysts and layer of host coelomocytes. FE TEM. F. Detailed view of outer surface of gametocyst wall containing fibrous material. TEM. G. Anastomoses between host cells and fibrous layer on the wall. TEM. H. Replica showing the outer surface of gametocyst wall with membrane remnants. FE TEM. black arrows – lamellae of gametocyst wall, es – empty space between lamellae or gametocyst wall and layer of coelomocytes, fib – fibrillar material on outer surface of the wall, gw – the wall of gametocyst, h – host cells, oo – oocysts.

Figure 7. General morphology of oocysts and oocyst walls. A. Living oocysts. DIC. B. Excystation of sporozoites caused by pressing. DIC. C. General view of oocyst surface with a funnel and tail. SEM. D-E. The view of the outer (D) and inner (E) oocyst’s surface. Note the presence of individual sporozoites inside the oocyst. SEM. F. Detailed view of three oocysts walls. SEM. G. General view of oocyst with multilayered envelope. SEM. H. Cross-sectioned oocyst with sporozoites. TEM. I-J. Details of oocyst wall forming “slender processes” (J). TEM. K. General view of fractured oocysts. FE TEM. L. Replica showing two layers of the wall. FE TEM. M. Details of hair-like ornamentation of the exospore shown on Fig. 3I–J. FE TEM. black arrow – ridges on the oocyst surface, f – funnel, hl – hair-like ornamentation of the exospore, oo – oocysts, ow1–3 – walls of three oocysts, paired black arrowheads – lamellae of endospore, r – residuum, sp – sporozoites, t – tail, white arrowhead – exospore.

Figure 8. Morphology of sporozoites. A-B. General view of oocyst in longitudinal (A) and transversal (B) section; I-III sporozoites sectioned in their anterior half, IV-VII sporozoites sectioned in their posterior half. TEM. C. Detail of sporozoite pellicle underlined with subpellicular microtubules. TEM. D. General view of tangentially sectioned sporozoites. Inset and white rectangle show bimembrane vesicle associated with pellicle. TEM. E. Cross-sectioned sporozoite approximately in the middle region of the cell. TEM. F. Sporozoite apical end in detail in longitudinal section. TEM. G-H. Sections through the anterior half of sporozoites. TEM. I. Superficial view on sporozoites’ apical end; the furrow and burgeon are marked by black circles. SEM. J. Multimembrane structures (scale bar = 100 nm is equal for all frames). TEM. black arrow – endospore, black arrowheads – plasma membrane, double arrowheads – polar ring, Ga – stalk of Golgi apparatus dictyosomes, m – mitochondrion, mi – micronemes, n – nucleus, paired black

arrowheads – membranes of IMC, r – residuum, rh – rhoptries, sl – sac-like structure, sp1- 4 – sporozoites, white arrow – exospore, white arrowheads – subpellicular microtubules.

Figure 9. Morphology of solitary sporozoites and zygote. A-B. Longitudinally (A) and cross (B) sectioned sporozoite. TEM. C. Cross section of zygote. TEM. D. Periphery of zygote in detail. TEM. ag – amylopectin granules, black arrow – endospore, Ga – Golgi apparatus, mi – micronemes, n – nucleus, white arrow – exospore.

Figure 10. SSU rDNA phylogenetic analysis of Lecudinoidea and related sequences (55 OTUs, alignment of 1,673 sites). The Bayesian tree was constructed under GTR+Г+I model. Numbers at the nodes denote Bayesian posterior probabilities (numerator) and ML bootstrap percentage (denominator). Black disks on the branches indicate the Bayesian posterior probabilities and the bootstrap percentages equal to or more than 0.95 and 95%, respectively.

Figure 11. Bayesian inference trees of alveolates obtained using the GTR+Г+I model for 62 sequences. A. LSU rDNA (alignment of 2,916 sites). B. Concatenated SSU and LSU (alignment of 4,524 sites). Numbers at the nodes indicate Bayesian posterior probabilities (numerator) and ML bootstrap percentage (denominator). Black dots on the branches indicate Bayesian posterior probabilities and bootstrap percentages of at least 95 and 95%, respectively. The newly obtained LSU sequences of Urospora travisiae and U. ovalis are indicated by a black rectangle. Accession numbers in the tree B are arranged in following order: SSU rDNA, LSU rDNA. The sequences of Babesia bigemina were obtained from the Sanger Institute genome project (www.sanger.ac.uk/Projects/B_bigemina/). Asterisks mark partial LSU rDNA sequences of small size (300-700 bp).

5.2.2 The effect of seawater ionic composition on motility of marine gregarines

For this experiment, the archigregarine Selenidium pygospionis and eugregarine Polyrhabdina sp., isolated from the intestine of the same host marine polychaete (Pygospio elegans) and exhibiting different modes of motility (bending in S. pygospionis trophozoites vs. gliding in Polyrhabdina sp. trophozoites/gamonts), were used. The motility index was calculated for S. pygospionis (n=25) and for Polyrhabdina sp. (n=25) during the experimental assays in artificial seawater with changed ionic composition (i.e. the proportion of individual salts was amended). The impact of specific composition of ions on motility was more pronounced in S. pygospionis incubated in seawater with doubled calcium or magnesium salts (= doubling of Ca2+ and Mg2+ ions), which was reflected in a lowest value of its motility index. However, comparing all the experimental and control groups, the changes in gregarines motility were negligible and therefore no clear conclusions could be drawn (Table 2). These preliminary data show that both of these marine gregarines are extremely tolerant to the changes of seawater ionic composition, which would make this environment unsuitable for the survival of other organisms.

Table 2. Motility index in Selenidium pygospionis and Polyrhabdina sp. incubated in seawater with changed ionic composition. Time (in minutes) from the start of the experiment / motility index Seawater Species 0 30 60 240 480 960 S 100 100 96 96 96 72 Naturalcomposition SW P 100 100 100 100 96 92 Artificial SW S 100 96 96 96 96 80 P 100 96 96 96 96 96 S 100 96 96 96 96 92 NaCl ↔ KCl P 100 100 100 96 92 92

2x MgCl2.6H2O, S 100 100 96 88 88 64

MgSO4.7H2O P 100 100 100 92 84 76 S 100 100 100 96 88 72 Ø CaCl2 P 100 96 96 96 96 88 S 100 96 92 88 88 68 2x CaCl2 P 100 100 100 100 96 88 Abbreviations: SW – seawater, S – S. pygospionis, P – Polyrhabdina sp.

5.2.3 Western blot

Western blot analysis was used to detect proteins that are the main components of various cytoskeletal structures in Apicomplexa (e.g. α-tubulin as a protein structure of microtubules), which are believed to be responsible for various types of cell motility. For this part of the study, commercially available primary antibodies against the α-tubulin, myosin, and spectrin, along with the specific mouse monoclonal IgG anti-actin antibody raised against Dictyostelium actin and recognising the actin in T. gondii and Plasmodium sp. were used (for more information see Material and methods chapter). Western blot analysis showed to be highly limited by the sample size, therefore it was not possible to use species from marine hosts (due to limited sampling period). Hence, model terrestrial eugregarines (species known to reach large dimensions and occur in high numbers within a single host) from laboratory kept insect colonies were used to test and standardise the protocols for the study needs. In addition, commercially available antibodies, despite numerous repetitions, often showed no or not reliable results. Multiple bands detected in some gels may indicate that used antibodies bound non-specifically. This situation was often observed in samples analysed for the presence of spectrin. On the other hand, the lack of bands after incubation with anti-myosin antibody (but in some cases also in samples tested for actin and α-tubulin) was possibly caused by low concentration of target protein in the sample. Only immunoblots shown below with anti-actin, anti-α-tubulin, and anti-spectrin antibodies gave positive results, although the specificity of reactions shown is not satisfactory (Figs 12, 13). The preliminary data obtained by Western blot analysis shown in this chapter are sketchy and hence were not used for publication. To increase the reliability of obtained data, production and application of species-specific reagents will be needed.

Figure 12A-B. Detection of actin (42 kDa) (A) and α-tubulin (50 kDa) (B) proteins in Gregarina garnhami. L – ladder (molecular weight) kDa. Abbreviations: HT – host tissue (control), GG – G. garnhami gamonts and syzygies, GO – G. garnhami mature gametocysts with oocysts in samples divided to supernatant (s) and pellet (p) part.

Figure 13A-B. Detection of α-tubulin (50 kDa) (A) and spectrin-like (260/240 kDa) (B) proteins in Blabericola cubensis and Protomagalhaensia granulosae. L – ladder (molecular weight) kDa. Abbreviations: BC – B. cubensis, PG – P. granulosae gamonts and syzygies in samples divided to supernatant (s) and pellet (p) part.

5.2.4 Morphological observations on terrestrial eugregarines

The data on model terrestrial eugregarines from laboratory kept insect hosts (including their general morphology, ultrastructural organisation of cell cortex and fluorescent visualisation of selected cytoskeletal proteins) were collected to compare target structures observed by the microscopic analyses with the detection of specific proteins by Western blot. The bands for actin revealed by Western blot (Fig. 12A) most likely correlated with cytoskeletal filamentous structures (myonemes and ectoplasmic network) observed in ectoplasm of Gregarina garnhami by different microscopic approaches (Kováčiková et al., 2018). Similarly, these structures were found in ectoplasm of B. cubensis and P. granulosae (Figs 14, 16). Microtubules, on the contrary, were not confirmed in studied eugregarines from insect hosts, although the presence of protein α-tubulin was detected by immunoblotting (Figs 12B, 13A). It is supposed that α-tubulin is present in eugregarines in unpolymerised form. Spectrin with different organisation pattern (depending on type of antibody used) was observed in B. cubensis gamonts under CLSM (Fig. 18), however Western blot did not show satisfying results (Fig. 13B).

Figure 14. Motility and organisation of cell cortex in eugregarine Protomagalhaensia granulosae. A. A motile syzygy. Note the obvious change in cell shape during the gliding indicating the flexibility of gamonts. LM, transmitted light. B. The syzygy formed by a single anterior primite and three small posterior satellites. LM, transmitted light. C. Tangential section of the eugregarine posterior end showing the pellicle organised in long wavy epicytic folds. TEM. D. The detail of cytoplasm. Note the presence of transversal myonemes organised in bundles running in rings perpendicular to the longitudinal cell axis. TEM. E. Longitudinal section of the epicytic folds. TEM. F. A detail of cross- sectioned micropores consisting of central duct bounded by a prominent collar. TEM. am – amylopectin, black arrow – epicytic fold, double white arrowhead – micropore, ec – ectoplasm, en – endoplasm, g – glycocalyx, imc – inner membrane complex, n – nucleus, p – primite, plm – plasma membrane, s – satellite, white arrow – myonemes, white arrowhead – ectoplasmic network.

Figure 15. Life cycle stages of the eugregarine Blabericola cubensis. A. Two gamonts joined in a syzygy. B. Formation of gametocyst wall around the paired gamonts. C. Formation of sporoducts in maturing gametocyst. D. Gametocyst dehiscence by sporoducts after 48 h of incubation in a wet chamber. E. Detail of protruded sporoduct with few attached oocysts. F. Oocysts released in chains from gametocyst through sporoduct to the external environment. G. Individual barrel-shaped oocysts with truncated ends. Inset shows an oocyst after sonication with released sporozoites. LM, transmitted light (A-G) and phase contrast (G inset). black arrow – oocyst, black arrowhead – sporoduct, white arrow – sporozoites.

Figure 16. Motility and organisation of cell cortex in eugregarine Blabericola cubensis. A. A mucous trail left behind gliding gamonts. LM, phase contrast. B. Various stages of trophozoites attached to the host tissue. SEM. C. Organisation of epicytic folds at higher magnification. SEM. D. General view of a cross-sectioned eugregarine showing the organisation of epicytic folds and differentiation of cytoplasm into the ectoplasm and granular endoplasm. Inset shows cross-sectioned micropore, obliquely sectioned duct and ectoplasmic network in detail. TEM. E. Cross-sectioned epicytic folds. TEM. F. Fractured epicytic fold. FE TEM. G. Fractured IMC showing the base of grooves between the epicytic folds, micropores and medium-sized pores. FE TEM. am – amylopectin, asterisk – cytoplasm, black arrow – ectoplasmic network, black arrowhead – pore, d – duct, double black arrowhead – gliding path, double white arrowhead – micropore, ec – ectoplasm, ee – EF of the external cytomembrane, ei – EF of the internal cytomembrane, en – endoplasm, ep – EF of the plasma membrane, fil – 12-nm apical filaments, g – glycocalyx, imc – inner membrane complex, lb – lamina basalis, n – nucleus, pe – PF of the external

cytomembrane, pi – PF of the internal cytomembrane, plm – plasma membrane, pp – PF of the plasma membrane, rds – rippled dense structures, white arrow – epicytic fold, white arrowhead – medium-sized pores

Figure 17. Localisation and organisation of actin filaments in Blabericola cubensis gamont. A. A composite view on eugregarine cortex created by flattening of all optical sections. B. A view of the eugregarine middle region created by flattening a series of optical sections. CLSM, phalloidin-TRITC. B - CLSM in a combination with transmission light.

Figure 18. Localisation and organisation of spectrin-like proteins in Blabericola cubensis gamont. A. Cortically localised spectrin (chicken) following the pattern of longitudinal epicytic folds in a composite view created by flattening a series of optical sections. B. Clusters of spectrin (human) regularly distributed in the deutomerite ectoplasm. CLSM, IFA. B – CLSM in a combination with transmission light.

Figure 19. Fluorescent labelling of Blabericola cubensis oocysts. A-B. Localisation of actin (IFA-FITC) on both oocyst poles and Hoechst bound to the DNA of sporozoites. C. Co-staining of phalloidin (phalloidin-TRITC) and Hoechst staining the nuclei of sporozoites. CLSM (A, C) and CLSM in a combination with transmitted light (B). Composite views created by flattening of all optical sections.

5.2.5 Morphological observations on marine eugregarine Polyrhabdina sp4.

Figure 20. Organisation of cell cortex in eugregarine Polyrhabdina sp.. A. general

4 Results from ultrastructural and CLSM observations of marine eugregarine Polyrhabdina sp. are shown only partially, considering unpublished re-description by Paskerova et al. Moreover, experimental assays with video documentation, electron microscopic and CLSM evaluation will be published in manuscript by Valigurová et al. focusing on motility of this species (author will be co-author in this paper).

view of gamont showing the position of nucleus. LM. B. A general view of gamont showing the organisation of epicytic folds. SEM. C. Cross-section showing the cytoplasm and arrangement of epicytic folds. TEM. D-E. Epicytic folds and subpellicular structures in detail. TEM. es – external space of epicytic folds, imc – inner membrane complex, is – internal space of epicytic fold, n – nucleus, plm – plasma membrane, white arrow – epicytic fold, white arrowhead – microtubules running perpendicular to longitudinal cell axis.

Figure 21. Localisation and organisation of cytoskeletal proteins in eugregarine Polyrhabdina sp.. A. Co-localisation of actin (IFA-FITC) dispersed throughout the entire cell, and myosin (IFA-TRITC) restricted to the cell cortex. B. Co-localisation of actin (IFA-FITC) and F-actin (phalloidin-TRITC). C-D. Organisation and localisation of F- actin (phalloidin-TRITC) (C) and actin (IFA-FITC) (D). Note the more intense staining of F-actin in eugregarine periphery and in the nucleus area. CLSM. Composite views created by flattening of all optical sections. ec – ectoplasm, en – endoplasm, imc – inner membrane complex, n – nucleus, plm – plasma membrane.

5.2.6 Molecular analysis

While analyses of SSU rDNA obtained from marine and terrestrial apicomplexans examined in this study were successfull, the sequences of actin and tubulin were not obtained, despite repeated use of primers designed by Leander and Keeling (2004) or newly designed degenerate primers for actin and α-tubulin. The problem of unsuccessful amplification could be linked to the incorrect primers sequences of not being complementary to the template, nonspecific binding to other template sequences or thermocycler parameters, while the first option seems to be most likely. Also low concentration of target DNA in the samples and/or samples contamination were considered according to Nanodrop measurements. In addition, besides cytoplasmic bacteria-like inclusion observed under TEM in some eugregarines (Mackenzie and Walker, 1979), archigregarine S. pygospionis and eugregarine Polyrhabdina sp. may be hyperparasitised by microsporidians (Paskerova et al., 2016; Sokolova et al., 2013). Thus, the bands with an expected size for the actin gene obtained by PCR from genomic DNA in eugregarine Polyrhabdina sp. (Fig. 22) most likely correspond to the actin of microsporidium Metchnikovella incurvata frequently parasitising its cytoplasm (Sokolova et al., 2013), as also confirmed by TEM and CLSM observations (Fig. 23). The fluorescent labelling of Polyrhabdina sp. actin revealed clusters of actin within eugregarine cytoplasm. These clusters correspond to the structures intensively labelled with DNA dye Hoechst, which represent the nuclei of microsporidians.

Figure 22. PCR from genomic DNA for actin gene in eugregarine Polyrhabdina sp.. Bands with expected size for actin gene (primers: actR-actS) are marked in black ellipse.

Figure 23. Localisation of DNA and actin in gamonts of eugregarine Polyrhabdina sp. parasitised by microsporidia. A. Hoechst bound to the DNA corresponding to the parasites in the eugregarine cytoplasm. Note that the nuclear stain did not label the eugregarine nucleus5. B. Accumulation of F-actin (phalloidin-TRITC). C. Accumulation of actin (IFA-FITC). CLSM in a combination with transmitted light (A) and CLSM (B- C). Composite views created by flattening of all optical sections. n – nucleus, white arrow – some of the intracellularly located microsporidia.

5 It is important to emphasize that it is extremely difficult to stain nuclei in eugregarines and archigregarines with fluorescent nuclear dyes such as DAPI or Hoechst (proved by testing of several protocols). The penetration of tested nuclear dyes through extremely resistant trimembrane pellicle and their pass to the nucleus is usually possible only after strong membrane permeabilisation resulting in obvious pellicle damage (undesirable for purposes of this study). 71

6 Discussion

Apicomplexans are a diverse group of unicellular parasitic protists that are responsible for widespread diseases in humans and domestic animals. Most likely the ancestral apicomplexans, parasitising marine annelids, spread to other marine invertebrates, and their radiation continued through freshwater and terrestrial invertebrates to vertebrate hosts (Cox, 1994). Their representatives exhibit enormous diversity in the motility modes and strategies utilized for host cell invasion and/or attachment strategies. However apicomplexans share one typical feature, namely the presence of apical complex at least at some stage of their life cycle. The apical complex is a unique invasion apparatus comprised of specialised secretory organelles (rhoptries, micronemes), polar ring(s) with associated subpellicular microtubules, and a conoid, and it is initially linked with a myzocytotic mode of feeding (sucking out the host cell cytoplasm), occurring in colpodellids, blastogregarines and archigregarines (Leander, 2008; Paskerova et al., 2018; Schrével and Desportes, 2015; Simdyanov and Kuvardina, 2007; Simdyanov et al., 2018; Wakeman and Horiguchi, 2017). A mucron derived from the apical complex, comprising the typical set of apical organelles and a mucronal vacuole, was observed in this study in the blastogregarine S. nematoides and in archigregarines S. pygospionis and S. pherusae (Paskerova et al., 2018; Simdyanov et al., 2018; Valigurová et al., 2017). The polar ring in these species gives rise to regularly spaced subpellicular microtubules. In contrast, the mucron of blastogregarine C. mesnili is strongly modified, lacking the conoid and rhoptries, and the present subpellicular microtubules are not connected by a polar ring. The myzocytosis appears not to be typical feeding strategy of protoccocidian E. duboscqi, in which the apical complex of organelles and subpellicular microtubules are absent during the endogenous phase of life cycle (Valigurová et al., 2015). Similarly in eugregarines, the apical complex and subpellicular microtubules disappear during the course of transformation of the sporozoite into the trophozoite stage attached to the host cell via an attachment organelle – epimerite, mucron or modified protomerite (Desportes and Schrével, 2013; Valigurová et al., 2007). Interestingly, in eugregarine C. cf. communis the structural organisation of the attachment apparatus (lenticular epimerite) exhibits a very specific pattern with the epimerite plasma membrane interrupted by numerous irregularly distributed pores. These pores are thought to release adhesives, most likely produced by the abundant microneme-like structures found in the parasite apical end and facilitating its adhesion to the host tissue. Attachment strategy in C. cf. communis appears

to combine the chemical fixation and mechanical anchoring to the host tissue provided by tiny long protrusion rising from the epimerite centre (Kováčiková et al., 2017). In the detached eugregarine stages (gamonts) or when the parasites are developing freely in the host cavities, the feeding likely occurs through cell surface nutrition via micropores (Schrével and Desportes, 2015), frequently documented between the epicytic folds. Micropores were also observed in C. cf. communis, G. garnhami, B. cubensis and P. granulosae investigated in this study. However, the specific structure of an epimerite in C. cf. communis indicate that modified apical region could be eventually dedicated to feeding in gregarines lacking a typical epimerite, as already proposed in G. cuneata from mealworms and actinocephalid gregarines (Cook et al., 2001; Valigurová, 2012). In archigregarine S. pygospionis the translucent vesicular structures were connected to the micropores. Besides that, globular and drop-shaped cytoplasmic vesicles, connected to the pellicle via ducts and comprising multi-membranous whorls, were suggested to have an additional role in surface-mediated nutrition, accessory to myzocytotic feeding (Kováčiková et al., 2019; Paskerova et al., 2018). Similar vesicular structures were also observed in other Selenidium species (Schrével et al., 2016; Wakeman and Horiguchi, 2017). The gliding motility is ensuring the migration of apicomplexan invasive stages (sporozoites, merozoites, tachyzoites, etc.) to the appropriate location within the host organism, and host cell invasion, and is likely generated by the glideosome (first announced for T. gondii and later for Plasmodium spp.) (Kappe et al., 2004; Keeley and Soldati, 2004; Opitz and Soldati, 2002). In this model, the actomyosin motor is expected to be embedded between the parasite plasma membrane and the IMC, and to require a stable subpellicular network of microtubules (Dubremetz et al., 1998; Kappe et al., 2004; Keeley and Soldati, 2004; Matuschewski and Schüler, 2008). Recent studies showed that the glideosome concept requires re-evaluation as new factors were discovered and numerous glideosome components (including actin and myosin) can be knocked out without complete blocking of motility (Egarter et al., 2014; Whitelaw et al., 2017), but its principal mechanism still appears largely to be valid (Heintzelman, 2015). Our studies revealed that different mechanisms are involved in motility displayed by representatives of early emerging groups of apicomplexans, despite the presence of key glideosome components such as actin, myosin and α-tubulin (as a protein structure of microtubules). It seems that, basal apicomplexans exhibit several mechanisms of cell motility that correlate with certain modifications of their cortex with associated cytoskeleton occurring

during the transformation of the invasive sporozoite into the vegetative trophozoite (Diakin et al., 2016; Kováčiková et al., 2019; Landers and Leander, 2005; Leander, 2008; MacMillan, 1973; Miles, 1968; Paskerova et al., 2018; Valigurová et al., 2017). Generally, the pellicle of apicomplexan parasites is formed by a plasma membrane underlain by a closely apposed IMC. The pellicle of parasites investigated in this study was either smooth (blastogregarine S. nematoides and archigregarine S. pherusae) or formed numerous longitudinal shallow folds (archigregarine S. pygospionis and protococcidian E. duboscqi), high folds with flattened tops (blastogregarine C. mesnili) or typical epicytic folds of the eugregarines separated by grooves and exhibiting specific structures (12-nm filaments, rippled dense structures and basal lamina) (Kováčiková et al., 2017, 2018, 2019; Paskerova et al., 2018; Simdyanov et al., 2018; Valigurová et al., 2015, 2017). Selenidium archigregarines exhibit pendular, rolling, coiling, and bending or so- called nematode-like movement (Fowell, 1936; Kováčiková et al., 2019; Leander, 2007; Leander, Harper and Keeling, 2003; Paskerova et al., 2018; Rueckert and Horák, 2017; Schrével, 1971a; Schrével and Desportes, 2015; Schrével and Phillipe, 1993; Schrével et al., 2016; Simdyanov and Kuvardina, 2007). Similarly, blastogregarine S. nematoides performs highly active pendular or twisting movements (Valigurová et al., 2017). In contrast, blastogregarine C. mesnili shows only weak motility with slow and intermittent bending movements, attributed to specific cortex modifications (Simdyanov et al., 2018). These movements in both the Selenidium archigregarines and blastogregarines are supposed to be facilitated by regular sets of subpellicular microtubules, which are usually organised in several layers (Schrével et al., 1974; Stebbings et al., 1974; Valigurová et al., 2017). The role of these microtubules was examined by experimental assays using the microtubule-destroying drugs and urea (Kováčiková et al., 2019; Schrével et al., 1974; Stebbings et al., 1974; Valigurová et al., 2017). Experimental motility assays revealed that depolymerisation of subpellicular microtubules in studied parasites begins in the caudal region and processes to the apical end towards the apical polar ring (MTOC) (Kováčiková et al., 2019; Valigurová et al., 2017). The subpellicular microtubules in majority of apicomplexan zoites end freely in the region behind the nucleus (de Souza and Attias, 2010; Morrissette and Sibley, 2002a). In blastogregarine and archigregarine trophozoites and gamonts, despite their resemblance to the hypertrophic zoite, microtubules run parallel with each other and extend over the entire length of the parasite (Paskerova et al., 2018; Schrével et al., 2016; Simdyanov et al., 2018; Valigurová et al., 2017). A mechanism based on the cooperation between the pellicle (IMC) and longitudinal subpellicular

microtubules, acting like an analogue of the musculocuticular system of nematodes, was proposed for archigregarine motility (Leander, 2007; Paskerova et al., 2018; Stebbings et al., 1974). Meanwhile, the sliding of microtubules provided by microtubule-associated molecular motors (MAPs) was suggested as the mechanism for the propagation of bending waves in Selenidium spp. and S. nematoides (Mellor and Stebbings, 1980; Stebbings et al., 1974; Valigurová et al., 2017). Association between mitochondria (providing chemical energy in ATP for MAPs activity) and subpellicular microtubules is important to ensure functional cell motility (Leander, 2006; Mellor and Stebbings, 1980; Schrével, 1971a; Valigurová et al., 2017). A correlation was confirmed between the dynamic motility in studied archigregarines and blastogregarines and the number of cortical mitochondria (Paskerova et al., 2018; Valigurová et al., 2017). Accordingly, the ectoplasm of actively moving Selenidium spp. comprises a higher number of subpellicular microtubules compared to slow-moving species (Schrével, 1971a; Simdyanov and Kuvardina, 2007). The involvement of the axial streak together with putative microfilaments was also discussed regarding the S. pygospionis motility (Fowell, 1936; Paskerova et al., 2018). Although the polymerised form of α-tubulin was demonstrated in protococcidian E. duboscqi, its mature trophozoites and gamonts lack subpellicular microtubules (Valigurová et al., 2015). The presence of actin filaments was confirmed in all basal apicomplexans investigated during this study: blastogregarine S. nematoides, archigregarine S. pygospionis, eugregarines C. cf. communis, Polyrhabdina sp., G. garnhami and B. cubensis, and protococcidian E. duboscqi. The filamentous actin in blastogregarine S. nematoides forms the cross-linking protein complexes, consisting of proteins embedded in the IMC and the network around subpellicular microtubules, that are thought to be associated with microtubules and to influence their spacing (Valigurová et al., 2017). According to experimental assays, the involvement of actin filaments in archigregarines motility is rather supportive (Kováčiková et al., 2019). In E. duboscqi, the subpellicular bands of longitudinally oriented actin-rich filaments forming beneath the IMC during trophozoite maturation, appear to substitute the subpellicular microtubules in later stages (Valigurová et al., 2015). For eugregarine free trophozoites, gamonts and syzygies gliding is a typical motility mode, performed with or without obvious changes in their cell shape (King, 1981, 1988; Kováčiková et al., 2017, 2018). More continuous gliding with only slight changes in cell shape, direction or speed was exhibited by G. garnhami, B. cubensis and

Polyrhabdina sp. It is hypothesised that in the course of environmental adaptations, the cortex of eugregarines became very rigid and, hence, they lost the wriggling ability typical of ancestral archigregarines (Valigurová et al., 2013). However, considering the flexible movements documented in some eugregarine species the cellular plasticity must be relatively high (Valigurová et al., 2013). As an example of this plasticity can be mentioned the progressive gliding enriched by jumping and rotational movements with rapid changes in gliding direction, cell flexions and even reverse movement displayed by the marine eugregarine C. cf. communis (Kováčiková et al., 2017). Similarly, P. granulosae, G. polymorpha and G. cuneata show significant cell flexions during movement, especially in their protomerite regions (Valigurová et al., 2013) or in both, the protomerite and the deutomerite regions (Kováčiková, personal observations). Eugregarines gliding is enriched by additional but prominent motility modes, including bending, curving or shortening of the longitudinal cell axis and intense movements of the protomerite, which are attributed to the presence of contractile elements previously entitled ‘myocyte’, (Crawley, 1905) corresponding to the ectoplasmic network and rib-like myonemes described in more recent studies (Beams et al., 1959; Heintzelman, 2004; Hildebrand, 1980; Kováčiková et al., 2018; Valigurová and Koudela, 2008; Valigurová et al., 2013; Walker et al., 1979). Contractile function of these structures is further supported by observation of stiff gamonts of G. steini, lacking the myonemes and showed no cell shape changes during their rapid gliding (Valigurová et al., 2013). To elucidate the possible role of actomyosin system in eugregarine gliding several biochemical and molecular investigations have been performed (Baines and King, 1989; Ghazali and Schrével, 1993; Ghazali et al., 1989; Heintzelman, 2004; Heintzelman and Mateer, 2008; Kováčiková et al., 2017, 2018; Philippe et al., 1982; Valigurová, 2012; Valigurová et al., 2013). The presence of actin predominantly polymerised into microfilaments was confirmed in eugregarine cell cortex (Kováčiková et al., 2017, 2018; Valigurová et al., 2013). The experimental study on motility of G. garnhami confirmed a dense layer of annular myonemes, running perpendicular to the longitudinal cell axis and forming visible bundles in ultrathin sections, to consist of actin filaments, and along with a network of ectoplasmic filaments having a fundamental role in gamonts motility (Kováčiková et al., 2018). The contraction of myonemes along with more compact ectoplasmic network likely results on grouping of superficially protruding superfolds together on one site of gregarine. This is supplemented with a denser secretion and accumulation of mucus, suggesting this part to be the eugregarine gliding side

(Kováčiková et al., 2018). Rib-like myonemes forming bundles perpendicular to the cell axis and ectoplasmic network were observed also in eugregarines P. granulosae and B. cubensis; in the latter also supported by positive phalloidin labelling confirming the presence of filamentous actin. Similar to G. garnhami, the rapid decline in motility was observed in both these eugregarines from cockroaches after application of cytochalasin D, hereby proving important role of actin filaments on gliding motility. The association of actin filaments with different types of myosins (A, B and F) appears to be likely, because these myosins were immunoflurescently detected in various Gregarina spp. in a similar localisation and organisation pattern (Heintzelman, 2004; Heintzelman and Mateer, 2008; Valigurová et al., 2013) as F-actin in G. garnhami (Kováčiková et al., 2018). The myosin in eugregarines from mealworms and C. cf. communis is restricted to the cortical region of the cell, following the pattern of the epicytic folds (Kováčiková et al., 2017; Valigurová et al., 2013). Also the spectrin-like proteins (considered to stabilise the eugregarine cortex and possibly affecting the cell motility) were immunofluorescently visualised in ectoplasm and cortex of B. cubensis and detected by immunoblotting. The eugregarine gliding movement is, in addition, facilitated by the unique architecture of epicytic folds and is supported by the intense secretion of mucopolysaccharides, densely coating the entire gregarine surface (Kováčiková et al., 2017, 2018; Valigurová et al., 2013). Importantly, the presence of subpellicular microtubules was not confirmed in investigated eugregarines (Kováčiková et al., 2017, 2018; Valigurová et al., 2013). However, the presence of tubulin-rich filamentous structures was detected in the area of septum separating the epimerite and the protomerite region along with α-tubulin clusters immunofluorescently localised within the cytoplasm of C. cf. communis, indicating its presence in unpolymerised form (Kováčiková et al., 2017). The occurrence of unpolymerised α-tubulin in eugregarines is further supported by results of Western blot analysis of G. garnhami, B. cubensis and P. granulosae. Despite the absence of longitudinally oriented subpellicular microtubules in later stages of eugregarine development (trophozoites, gamonts, syzygies), their presence was confirmed in sporozoites stage of herein studied urosporid eugregarines. The pellicle of spindle-shaped Urospora sp. sporozoites is underlain by 20 longitudinal microtubules that arise from the apical polar ring and proceed towards the posterior end (Diakin et al., 2018). Overall, for all investigated groups of early emerging apicomplexans it can be concluded that despite the presence of the basic motor proteins (actin, myosin, and α- tubulin), the principles of their motility differ from the glideosome concept described in

apicomplexan zoites. In spite of the striking morphological similarity of blastogregarine and archigregarine trophozoites, considered to be overgrown apicomplexan zoites (hypersporozoites), their motility obviously differs from the substrate-dependent gliding in Toxoplasma or Plasmodium invasive stages (Håkansson et al., 1999; Münter et al., 2009). Hypothetically, this bending motility predominates and possibly replaces the substrate-dependent gliding during the transformation of sporozoite into the trophozoite stage. In addition, both the archigregarine S. pygospionis and blastogregarine S. nematoides continue to bend even after detachment from the host tissue (Kováčiková et al. 2019; Valigurová et al., 2017). Eugregarines perform a progressive gliding motility and in most cases they indeed glide contacting the substrate, however they lack one of the essential components involved in glideosome motor, the subpellicular microtubules (Kováčiková et al., 2017, 2018; Valigurová et al., 2013). In addition, the results of this study indicate that the dynamic process of actin polymerisation and subsequent rapid depolymerisation, essential for apicomplexan glideosome mechanism (Opitz and Soldati, 2002), is not required for gliding motility in eugregarines. In contrast, only the polymerised form of actin seems to be the main leading motor structure responsible for gliding in representatives of investigated eugregarines.

7 Conclusions and perspectives

According to main aims of this study, the results are summarised in following points:  Field collection and processing of material were performed in different environmental conditions (examination of marine and terrestrial apicomplexan species). In addition, the type of material fixation was conditioned by individual studied group or even by the parasite species and therefore standardisation of fixation protocols for each species was required.  Observation and photo/video documentation of cell motility in selected apicomplexans in vitro revealed the existence of various movement modes in representatives of phylogenetically distinct groups. Characteristic progressive gliding motility with slight or intensive changes in cell shape, speed or direction was observed in gamonts of studied eugregarines. Non-progressive bending, twisting or pendular movement was, in contrast, performed by archigregarines and blastogregarines.  Experimental motility assays using various cytoskeletal drugs affecting the parasite motility revealed extreme tolerance and survival of studied apicomplexans after their exposure to these drugs at high concentration. In addition, different reactions to applied drugs were observed between individual species, even between the species collected from the same host organism. Incubations of parasites in media with different ionic composition indicate that marine gregarines are extremely tolerant to the changed ionic composition of the seawater.  Comparative morphological analysis of structures responsible for motility revealed two main motors elements. First, in species possessing longitudinally folded (archigregarine S. pygospionis and blastogregarine C. mesnili) or smooth (archigregarine S. pherusae and blastogregarine S. nematoides) pellicle underlined by the subpellicular microtubules, and performing typical bending/pendular/twisting movements, the main motility motor appears to be constituted by the subpellicular microtubules while the actin filaments have rather supportive function. Second, in eugregarines with pellicle folded into numerous longitudinal epicytic folds and exhibiting gliding motility, myonemes and ectoplasmic network formed by actin filaments represent the main component responsible for movement and cell flexions. Engagement of epicytic folds into gliding motility is also supposed, however the exact mechanism remains unclear. 79

Intensive secretion of mucopolysaccharides coating the surface of gliding eugregarines represents additional element facilitating the gliding movement. Importantly, presented study revealed that principles of motility in archigregarines, blastogregarines and eugregarines differ from the glideosome concept, despite the presence of the basic motor proteins, actin (predominantly occurring in polymerised form), myosin and α-tubulin in these organisms.  Biochemical analysis of target proteins was partially successful in the determination of proteins forming the cytoskeletal structures responsible for motility. Western blot analysis confirmed the presence of actin and revealed α- tubulin in investigated eugregarine species. This method, however, turned out to be limited by sample size (proteins concentration), therefore did not allow examination of apicomplexans from marine hosts (due to limited sampling period). Despite repeated testing of a primer for α-tubulin and various primers for actin used in molecular analyses in other similar studies, this study failed to molecularly identify the cytoskeletal proteins of the gregarine species investigated herein.

The main success of present thesis is markedly prolonged incubation and survival of marine and terrestrial apicomplexans during experiments with different cytoskeletal drugs. Therefore, further development of experimental assays on representatives of basal apicomplexan lineages is highly prospective for understanding the principles and evolution of motility in Apicomplexa and to define the main motor structures responsible for movement inside individual groups. In addition, comprehensive biochemical and molecular research performed on model basal apicomplexans is essential. For immunoblotting and/or immunofluorescence analyses it is important to develop, produce and apply species-specific antibodies, since application of commercially available antibodies may lead to their non-specific binding. In molecular studies, further testing of various primers and standardisation of protocols is needed to obtain required results. In addition, only hyperparasite-free cells should be used for modular biological analysis. In this regard, preliminary microscopic (TEM, CLSM) detection of potential intracellular bacteria or microsporidia in apicomplexan samples taken for molecular analysis would be beneficial.

8 References

Adl S.M., Simpson A.G, Lane C.E, Lukeš L., Bass D., Bowser S.S., Brown M.V., et al. (2012). The Revised Classification of Eukaryotes. J Eukaryot Microbiol 59 (5): 429– 493. Aikawa M. (1971). Plasmodium: The fine structure of malarial parasites. Exp Parasitol 30 (2): 284–320. Aldeyarbi H.M., Karanis P. (2016). The ultra-structural similarities between Cryptosporidium parvum and the gregarines. J Eukaryot Microbiol 63 (1): 79–85. Angrisano F., Riglar D.T., Sturm A., Volz J.C., Delves M.J., Zuccala E.S., Turnbull L., et al. (2012). Spatial localisation of actin filaments across developmental stages of the malaria parasite. PLoS One 7 (2): e32188. Baines I., King C.A. (1989). Demonstration of actin in the protozoon — Gregarina. Cell Biol Int Rep 13 (8): 679–686. Barta J.R., Thompson R.C.A. (2006). What is Cryptosporidium? Reappraising its biology and phylogenetic affinities. Trends Parasitol 22 (10): 463–468. Bartošová-Sojková P., Oppenheim R.D, Soldati-Favre D., Lukeš J. (2015). Epicellular apicomplexans: Parasites ‘On the way in’. PLoS Pathogens 11 (9): e1005080. Beams H.W., Tahmisian T.N., Devine R.L., Anderson E. (1959). Studies on the fine structure of a gregarine parasitic in the gut of the grasshopper, Melanoplus differentialis. J Protozool 6 (2): 136–146. Branton D., Bullivant S., Gilula N.B., Karnovsky M.J., Moor H., Muhlethaler K., Northcote D.H., et al. (1975). Freeze-etching nomenclature. Science 190 (4209): 54– 56. Brasil L. (1906). Eleutheroschizon duboscqi, sporozoaire nouveau parasite de Scoloplos armiger O.F. Müller. Arch Zool Exp Gen 4: 17–22. Bruevitch S.V. (1960). Hydrochemical studies of the White Sea. In: Trudy instituta okeanologii imeni P. P. Shirshova Akademii nauk SSSR, 42: 199-254. Canning E.U. (1956). A new eugregarine of locusts, Gregarina garnhami n.sp., parasitic in Schistocerca gregaria Forsk. J Protozool 3 (2): 50–62. Carreno R.A., Martin D.S., Barta J.R. (1999). Cryptosporidium is more closely related to the gregarines than to coccidia as shown by phylogenetic analysis of apicomplexan parasites inferred using small-subunit ribosomal RNA gene sequences. Parasitol Res 85 (11): 899–904.

Carruthers V.B., Giddings O.K., Sibley L.D. (1999). Secretion of micronemal proteins is associated with Toxoplasma invasion of host cells. Cell Microbiol 1 (3): 225–235. Caullery M., Mesnil F. (1898). Sur un Sporozoaire aberrant (Siedleckia n. g.). C R Soc Biol 5: 1093–1095. Chambouvet A., Valigurová V., Pinheiro L.M., Richards T.A., Jirků M. (2016). Nematopsis temporariae (Gregarinasina, Apicomplexa, Alveolata) is an intracellular infectious agent of tadpole livers. Environ Microbiol Rep 8 (5): 675–679. Chatton E., Dehorne L. (1929). Observations sur les Sporozoaires du genre Siedleckia. S. mesnili, n. sp. et S. dogieli n.sp. Arch Anat Microsc 25: 530–543. Chatton E., Villeneuve F. (1936a). La sexualite et le cycle évolutif des Siedleckia d’apres l’étude de S. caulleryi, n. sp. Hologrégarines et Blastogrégarines. Sporozoaires Hologamétogènes et Blastogamétogènes. CR Acad Sci D Nat 203: 505–508. Chatton E., Villeneuve F. (1936b). Le cycle évolutif de l’Eleutheroschizon duboscqui Brasil. Preuve expérimentale de l’absence de schizogonie chez cette forme et chez la Siedleckia caulleryi Ch. et Vill. CR Acad Sci D Nat 203: 834–837. Chobotar B., Scholtyseck E. (1982). Ultrastructure. In: P.L. Long (Ed.). The biology of the coccidia, Edward Arnold Press, London, 101-165. Clopton R.E. (2012a). Synoptic revision of Blabericola (Apicomplexa: Eugregarinida: Blabericolidae) parasitizing blaberid cockroaches (Dictyoptera: Blaberidae), with comments on delineating gregarine species boundaries. J Parasitol 98 (3): 572–583. Clopton R.E. (2012b). Redescription of Protomagalhaensia blaberae Peregrine, 1970 (Apicomplexa: Eugregarinida: Blabericolidae) parasitizing the bolivian cockroach, Blaberus boliviensis (Dictyoptera: Blaberidae). Comp Parasitol 79 (2): 182–191. Cook T.J.P., Janovy J. Jr., Clopton R.E. (2001). Epimerite-host epithelium relationships among eugregarines parasitizing the damselflies Enallagma civile and Ischnura verticalis. J Parasitoly 87 (5): 988–996. Cox F.E. (1994). The evolutionary expansion of the sporozoa. Int J Parasitology 24 (8): 1301–1316. Crawley H. (1905). The movements of gregarines. Proc Acad Nat Sci Philadelphia 57: 89–99. Dallai R., Talluri M.V. (1983). Freeze-fracture study of the gregarine trophozoite: I. the top of the epicyte folds. Boll Zool 50 (3–4): 235–244. De Souza W., Attias M. (2010). Subpellicular microtubules in Apicomplexa and Trypanosomatids. In: W. de Souza (Ed.). Structures and organelles in pathogenic

protists, Springer-Verlag, Berlin, Heidelberg, 27–62. Desportes I., Schrével J. (2013). The gregarines, the early branching Apicomplexa (2 vols). In: Treatise on zoology - Anatomy, taxonomy, biology. BRILL, Leiden – The Netherlands, 781 pp. Diakin A., Paskerova G.G., Simdyanov T.G., Aleoshin V.V, Valigurová A. (2016). Morphology and molecular mhylogeny of coelomic gregarines (Apicomplexa) with different types of motility: Urospora ovalis and U. travisiae from the polychaete Travisia forbesii. Protist 167 (3): 279–301. Diakin A., Vaškovicová N., Paskerova G.G., Kováčiková M., Nebesářová J., Valigurová A. Gametocysts, oocysts and sporozoites morphology of urosporid gregarines with remarks on molecular phylogeny as inferred from rDNAs. A manuscript draft published In: Diakin A. (2018). Biology of marine early emerging apicomplexans. Ph.D. thesis, Masaryk University, Brno, p. 106-130. Dobrowolski J.M., Carruthers V.B, Sibley L.D. (1997). Participation of myosin in gliding motility and host cell invasion by Toxoplasma gondii. Mol Microbiol 26 (1): 163– 173. Dobrowolski J.M., Niesman I.R., Sibley L.D. (1997). Actin in the parasite Toxoplasma gondii is encoded by a single copy gene, act1 and exists primarily in a globular form. Cell Motil Cytoskeleton 37 (3): 253–262. Dobrowolski J.M., Sibley L.D. (1996). Toxoplasma invasion of mammalian cells is powered by the actin cytoskeleton of the parasite. Cell 84 (6): 933–939. Dogiel V. (1910). Beiträge zur Kenntnis der Gregarinen. IV. Callynthrochlamys phronimae Frenz. u. a. m. Arch Protistenkd 20: 60–78. Drewry L.L., Sibley L.D. (2015). Toxoplasma actin is required for efficient host cell invasion. MBio 6 (3): e00557-15. Dubremetz J.F., Garcia-Réguet N., Conseil V., Fourmaux M.N. (1998). Apical organelles and host-cell invasion by Apicomplexa. Int J Parasitol 28 (7): 1007–1013. Egarter S., Andenmatten N., Jackson A.J., Whitelaw J.A, Pall G., Black J.A., Ferguson D.J., Tardieux I., Mogilner A., Meissner M. (2014). The Toxoplasma acto-myoA motor complex is important but not essential for gliding motility and host cell invasion. PLoS ONE 9 (3): e91819. Endo T., Yagita K., Yasuda T., Nakamura T. (1988). Detection and localization of actin in Toxoplasma gondii. Parasitol Res 75 (2): 102–106. Forney J.R., Vaughan D.K., Yang S., Healey M.C. (1998). Actin-dependent motility in

Cryptosporidium parvum sporozoites. J Parasitol 84 (5): 908–913. Fowell R.R. (1936). The fibrillar structures of Protozoa, with special reference to schizogregarines of the genus Selenidium. J Roy Micr Soc 56: 12–28. Frénal K., Dubremetz J.F., Lebrun M., Soldati-Favre D. (2017). Gliding motility powers invasion and egress in Apicomplexa. Nat Rev Microbiol 15 (11): 645–660. Frénal K., Foth B.J., Soldati D. (2008). Myosin class XIV and other myosins in protists. In: L.M. Coluccio (Ed.). Myosins. Proteins and cell regulation (vol. 7). Springer, Dordrecht, 421–440. Frénal K., Polonais V., Marq J.B., Stratmann R., Limenitakis J., Soldati-Favre D. (2010). Functional dissection of the apicomplexan glideosome molecular architecture. Cell Host Microbe 8 (4): 343–357. Gaskins E., Gilk S., DeVore N., Mann T., Ward G., Beckers C. (2004). Identification of the membrane receptor of a class XIV myosin in Toxoplasma gondii. J Cell Biol 165 (3): 383–393. Ghazali M., Philippe M., Deguercy A., Gounon P., Gallo J.M., Schrével J. (1989). Actin and spectrin-like (Mr=260-240 000) proteins in gregarines. Biol Cell 67 (2): 173– 184. Ghazali M., Schrével J. (1993). Myosin like protein (Mr 175,000) in Gregarina blaberae. J Eukaryot Microbiol 40 (3): 345–354. Ghazali M., Schrével J. (1995). Identification and localization of proteins in gregarines that are immunologically related to smooth muscle α-actinin. Eur J Protistol 31 (3): 292–301. Håkansson S., Morisaki H., Heuser J., Sibley L.D. (1999). Time-lapse video microscopy of gliding motility in Toxoplasma gondii reveals a novel, biphasic mechanism of cell locomotion. Mol Biol Cell 10 (11): 3539–3547. Heintzelman M.B. (2004). Actin and myosin in Gregarina polymorpha. Cell Motil Cytoskeleton 58 (2): 83–95. Heintzelman M.B. (2015). Gliding motility in apicomplexan parasites. Semin Cell Dev Biol 46: 135–142. Heintzelman M.B., Mateer M.J. (2008). GpMyoF, a WD40 repeat-containing myosin associated with the myonemes of Gregarina polymorpha. J Parasitol 94 (1): 158– 168. Heintzelman M.B, Schwartzman J.D. (1997). A novel class of unconventional myosins from Toxoplasma gondii. J Mol Biol 271 (1): 139–146.

Hildebrand H.F. (1980). Elektronenmikroskopische Untersuchungen an den Entwicklungsstadien des Trophozoiten von Didymophyes gigantea (Sporozoa, Gregarinida). III. Die Feinstruktur des Epizyten mit besonderer Berücksichtigung der kontraktilen Elemente. Z Parasitenkd 64: 29–46. Hildebrand H.F., Vinckier D. (1975). Nouvelles observations sur la Grégarine Didymophyes gigantea Stein. J Protozool 22 (2): 200–213. Hu K., Roos D.S, Murray J.M. (2002). A novel polymer of tubulin forms the conoid of Toxoplasma gondii. J Cell Biol 156 (6): 1039–1050. Jewett T.J, Sibley L.D. (2003). Aldolase forms a bridge between cell surface adhesins and the actin cytoskeleton in apicomplexan parasites. Mol Cell 11 (4): 885–894. Jirků M., Modrý D., Šlapeta J., Koudela B., Lukeš J. (2002). The phylogeny of Goussia and Choleoeimeria (Apicomplexa; Eimeriorina) and the evolution of excystation structures in coccidia. Protist 153 (4): 379–390. Kappe S.H.I., Buscaglia C.A., Bergman L.W., Coppens I., Nussenzweig V. (2004). Apicomplexan gliding motility and host cell invasion: overhauling the motor model. Trends Parasitol 20 (1): 13–16. Keeley A., Soldati D. (2004). The glideosome: a molecular machine powering motility and host-cell invasion by Apicomplexa. Trends Cell Biol 14 (10): 525–528. King C.A. (1981). Cell surface interaction of the protozoan Gregarina with concanavalin A beads - implications for models of gregarine gliding. Cell Biol Int Rep 1 (3): 279- 305. King C.A. (1988). Cell Motility of sporozoan protozoa. Parasitol Today 4 (11): 315–319. King C.A., Lee K. (1982). Effect of trifluoperazine and calcium ions on gregarine gliding. Experientia 38: 1051–1052. Kolman J., Clopton R., Clopton D.T. (2015). Effects of developmental temperature on gametocysts and oocysts of two species of gregarines Blabericola migrator and Blabericola cubensis (Apicomplexa: Eugregarinida: Blabericolidae) parasitizing blaberid cockroaches (Dictyoptera: Blaberidae). J Parasitol 101 (6): 651–657. Kono M., Herrmann S., Loughran N.B., Cabrera A., Engelberg K., Lehmann C., Sinha D., et al. (2012). Evolution and architecture of the inner membrane complex in asexual and sexual stages of the malaria parasite. Mol Biol Evol 29 (9): 2113–2132. Kováčiková M., Paskerova G.G., Diakin A., Simdyanov T.G., Vaškovicová N., Valigurová A. (2019). Motility and cytoskeletal organisation in the archigregarine Selenidium pygospionis (Apicomplexa): observations on native and experimentally

affected parasites. Parasitol Res 'in press'. https://doi.org/10.1007/s00436-019-06381-z Kováčiková M., Simdyanov T.G., Diakin A., Valigurová A. (2017). Structures related to attachment and motility in the marine eugregarine Cephaloidophora cf. communis (Apicomplexa). Eur J Protistol 59: 1–13. Kováčiková M., Vaškovicová N., Nebesářová J., Valigurová A. (2018). Effect of jasplakinolide and cytochalasin D on cortical elements involved in the gliding motility of the eugregarine Gregarina garnhami (Apicomplexa). Eur J Protistol 66: 97–114. Kumpula E.P., Pires I., Lasiwa D., Piirainen H., Bergmann U., Vahokoski J., Kursula I. (2017). Apicomplexan actin polymerization depends on nucleation. Sci Rep 7 (1): 12137. Landers S.C., Leander B.S. (2005). Comparative surface morphology of marine coelomic gregarines (Apicomplexa, Urosporidae): Pterospora floridiensis and Pterospora schizosoma. J Eukaryot Microbiol 52 (1): 23–30. Leander B.S. (2006). Ultrastructure of the archigregarine Selenidium vivax (Apicomplexa) – A dynamic parasite of sipunculid worms (host: Phascolosoma agassizii). Mar Biol Res 2 (3): 178–190. Leander, B.S. (2007). Molecular phylogeny and ultrastructure of Selenidium serpulae (Apicomplexa, Archigregarinia) from the calcareous tubeworm Serpula vermicularis (Annelida, Polychaeta, Sabellida). Zool Scr 36 (2): 213–227. Leander B.S. (2008). Marine gregarines: evolutionary prelude to the apicomplexan radiation? Trends Parasitol 24 (2): 60–67. Leander B.S, Clopton R.E., Keeling P.J. (2003). Phylogeny of gregarines (Apicomplexa) as inferred from small-subunit rDNA and beta-tubulin. Int J Syst Evol Microbiol 53 (Pt 1): 345–354. Leander B.S, Harper J.T., Keeling P.J. (2003). Molecular phylogeny and surface morphology of marine aseptate gregarines (Apicomplexa) Selenidium spp. and Lecudina spp. J Parasitol 89 (6): 1191–1205. Leander B.S., Keeling P.J. (2004). Early evolutionary history of dinoflagellates and apicomplexans (Alveolata) as inferred from Hsp90 and actin phylogenies. J Phycol 40 (2): 341–350. Leander B.S, Lloyd S.A.J., Marshall W., Landers S.C. (2006). Phylogeny of marine gregarines (Apicomplexa) — Pterospora, Lithocystis and Lankesteria — and the

origin(s) of coelomic parasitism. Protist 157 (1): 45–60. Levine N.D. (1971). Taxonomy of the Archigregarinorida and Selenidiidae (Protozoa, Apicomplexa). J Protozool 18 (4): 704–717. Levine N.D. (1973). Grellia gen. n. for Eucoccidium of Grell (1953) Preoccupied*. J Protozool 20 (5): 548–549. Mackenzie C., Walker M.H. (1979). Bacteria-like structures in the endoplasm of Gregarina garnhami (Eugregarinida, Protozoa). Cell Tissue Res 202: 33–39. Mackenzie C., Walker M.H. (1983). Substrate contact, mucus, and eugregregarine gliding. J Protozool 30 (1): 3–8. MacMillan, W.G. (1973). Conformational changes in the cortical region during peristaltic movements of a gregarine trophozoite*. J Protozool 20 (2): 267–274. Matuschewski K., Schüler H. (2008). Actin/myosin-based gliding motility in apicomplexan parasites. In: B.A Burleigh, D. Soldati-Favre (Eds.). Molecular mechanisms of parasite invasion. Subcell Biochem, 110–120. Mawrodiadi P.A. (1908). The barnacles of the Black Sea and their gregarine parasites. Preliminary note. Mem Soc Nat Nouvelles Odessa 32: 101–132. Meissner M., Ferguson D.J.P, Frischknecht F. (2013). Invasion factors of apicomplexan parasites: essential or redundant? Curr Opin Microbiol 16 (4): 438–444. Mellor J.S., Stebbings H. (1980). Microtubules and the propagation of bending waves by the archigregarine, Selenidium fallax. J Exp Biol 87: 149–161. Miles H.B. (1968). The fine structure of the epicyte of the acephaline gregarines Monocystis lumbriciolidi, and Nematocystis magna: observations by electron microscope. Rev Ibér Parasitol 28: 455–465. Morrissette N.S., Murray J.M., Roos D.S. (1997). Subpellicular microtubules associate with an intramembranous particle lattice in the protozoan parasite Toxoplasma gondii. J Cell Sci 110 (Pt 1): 35–42. Morrissette N.S., Sibley L.D. (2002a). Cytoskeleton of apicomplexan parasites. Microbiol Mol Biol Rev 66 (1): 21–38. Morrissette N.S., Sibley L.D. (2002b). Disruption of microtubules uncouples budding and nuclear division in Toxoplasma gondii. J Cell Sci 115 (Pt 5): 1017–1025. Münter S., Sabass B., Selhuber-Unkel C., Kudryashev M., Hegge S., Engel U., Spatz J.P., Matuschewski K., Schwarz U.S., Frischknecht F. (2009). Plasmodium sporozoite motility is modulated by the turnover of discrete adhesion sites. Cell Host Microbe 6 (6): 551–562.

Nichols B., Chiappino M.L. (1987). Cytoskeleton of Toxoplasma gondii. J Protozool 34 (2): 217–226. Opitz C., Soldati D. (2002). The Glideosome: A dynamic complex powering gliding motion and host cell invasion by Toxoplasma gondii. Mol Microbiol 45 (3): 597– 604. Ostrovska K., Paperna I. (1990). Cryptosporidium sp. of the starred lizard Agama stellio: ultrastructure and life cycle. Parasitol Res 76: 712–720. Paskerova G.G., Frolova E.V., Kováčiková M., Panfilkina T.S., Mesentev Y.S., Smirnov A.V., Nassonova E.S. (2016). Metchnikovella dogieli sp. n. (Microsporidia: Metchnikovellida), a parasite of archigregarines Selenidium sp. from polychaetes Pygospio elegans. Protistology 10 (4): 148-157. Paskerova G.G., Miroliubova T.S., Diakin A., Kováčiková M., Valigurová A., Guillou L., Aleoshin V.V., Simdyanov T.G. (2018). Fine structure and molecular phylogeny of two marine gregarines, Selenidium pygospionis n. sp. and S. pherusae n. sp., with notes on the phylogeny of Archigregarinida (Apicomplexa). Protist 169 (6): 826– 852. Peregrine P.C. (1970). Gregarines found in cockroaches of the genus Blaberus. Parasitology 61 (1): 135–151. Periz J., Whitelaw J., Harding C., Gras S., Del Rosario Minina M.I., Latorre-Barragan F., Lemgruber L., et al. (2017). Toxoplasma gondii F-actin forms an extensive filamentous network required for material exchange and parasite maturation. Elife 6: e24119. Perkins F.O., Barta J., Clopton R., Peirce M.A., Upton S.J. (2000). Phylum Apicomplexa. In: J.J. Lee, G.F. Leedale and P. Bradbury (Eds). The illustrated guide to the Protozoa (2nd Ed). Society of Protozoologists, Lawrence, KS, USA, 190-370. Philippe M., Vinckier D., Dubremetz J.F., Schrével J. (1982). The three cortical membranes of the gregarines (parasitic protozoa). III. Comparative studies of the membrane proteins among different sporozoan species during their vegetative phase. J Protozool 29 (3): 424–430. Preston T.M., King C.A. (1992). Evidence for the expression of actomyosin in the infective stage of the sporozoan protist Eimeria. Cell Biol Int Rep 16 (4): 377–381. Ray H.N. (1930). Studies on some sporozoa in polychaete worms: I. Gregarines of the genus Selenidium. Parasitology 22 (3): 370–398. Rueckert S., Horák A. (2017). Archigregarines of the english channel revisited: new

molecular data on Selenidium species including early described and new species and the uncertainties of phylogenetic relationships. PLoS ONE 12(11): e0187430. Russell D.G., Burns R.G. (1984). The polar ring of coccidian sporozoites: a unique microtubule-organizing centre. J Cell Sci 65 (1): 193–207. Schmitz S., Grainger M., Howell S., Calder L.J., Gaeb M., Pinder J.C., Holder A.A., Veigel C. (2005). Malaria parasite actin filaments are very short. J Mol Bio 349 (1): 113–125. Schmitz S., Schaap I.A., Kleinjung J., Harder S., Grainger M., Calder L., Rosenthal P.B., Holder A.A., Veigel C. (2010) Malaria parasite actin polymerization and filament structure. J Biol Chem 285 (47): 36577–36585. Scholtyseck E., Mehlhorn H. (1970). Ultrastructural study of characteristic organelles (paired organelles, micronemes, micropores) of sporozoa and related organisms. Z Parasitenkd 34 (2): 97–127. Schrével J. (1971a). Contribution à l’étude des Selenidiidae parasites d'annélides polychètes. II. Ultrastructure de quelques trophozoïtes. Protistologica 7: 101–130. Schrével J. (1971b). Observations biologiques et ultrastructurales sur les Selenidiidae et leurs conséquences sur la systématique des Grégarinomorphes. J Protozool 18 (3): 448–470. Schrével J., Buissonnets S., Metais M. (1974). Action de l'urée sur la motilité et les microtubules sous pelliculaires du protozoaire Selenidium hollandei. C R Acad Sci Paris 278: 2201–2204. Schrével J., Caigneaux E., Gros D., Philippe M. (1983). The three cortical membranes of the gregarines. I. Ultrastructural organization of Gregarina blaberae. J Cell Sci 61: 151–174. Schrével J., Desportes I. (2015). Gregarines. In: H. Mehlhorn (Ed.). Encyclopedia of Parasitology. Springer-Verlag, Berlin, Heidelberg, 47 pp. Schrével J., Philippe M. (1993). The Gregarines. In: J.P. Kreier, J.R. Baker (Eds.). Parasitic Protozoa (2nd Ed). Academic Press, 133-245. Schrével J., Valigurová A., Prensier G., Chambouvet A., Florent I., Guillou L. (2016). Ultrastructure of Selenidium pendula, the type species of archigregarines, and phylogenetic relations to other marine Apicomplexa. Protist 167 (4): 339–368. Schüler H., Mueller A.K., Matuschewski K. (2005). Unusual properties of Plasmodium falciparum actin: new insights into microfilament dynamics of apicomplexan parasites. FEBS Lett 579 (3): 655–660.

Schwartzman J.D., Krug E.C., Binder L.I., Payne M.R. (1985). Detection of the microtubule cytoskeleton of the coccidian Toxoplasma gondii and the hemoflagellate Leishmania donovani by monoclonal antibodies specific for β-tubulin. J Protozool 32 (4): 747–749. Schwiakoff W. (1894). Uber die Ursache der fortschreitenden Bewegung der Gregarinen. Z Wiss Zool 58: 340–354. Shaw M.K., Tilney L.G. (1999). Induction of an acrosomal process in Toxoplasma gondii: visualization of actin filaments in a protozoan parasite. Proc Natl Acad Sci U S A 96 (16): 9095–9099. Simdyanov T.G., Diakin A., Aleoshin V.V. (2015). Ultrastructure and 28S rDNA phylogeny of two gregarines: Cephaloidophora cf. communis and Heliospora cf. longissima with remarks on gregarine morphology and phylogenetic analysis. Acta Protozool 54 (3): 241–262. Simdyanov T.G., Guillou L., Diakin A.Y., Mikhailov K.V., Schrével J., Aleoshin V.V. (2017). A new view on the morphology and phylogeny of eugregarines suggested by the evidence from the gregarine Ancora sagittata (Leuckart, 1860) Labbé, 1899 (Apicomplexa: Eugregarinida). PeerJ 5: e3354. Simdyanov T.G., Kuvardina O.N. (2007). Fine structure and putative feeding mechanism of the archigregarine Selenidium orientale (Apicomplexa: Gregarinomorpha). Eur J Protistol 43 (1): 17–25. Simdyanov T.G., Paskerova G.G., Valigurová A., Diakin A., Kováčiková M., Schrével J., Gillou L., Dobrovolskij A.A., Aleoshin V.V. (2018). First ultrastructural and molecular phylogenetic evidence from the blastogregarines, an early branching lineage of plesiomorphic Apicomplexa. Protist 169 (5): 697–726. Siński E., Behnke M.J. (2004). Apicomplexan parasites: environmental contamination and transmission. Pol J Microbiol 53: 67–73. Sokolova Y.Y., Paskerova G.G., Rotari Y.M., Nassonova E.S., Smirnov A.V. (2013). Fine structure of Metchnikovella incurvata Caullery and Mesnil, 1914 (Microsporidia), a hyperparasite of gregarines Polyrhabdina sp. from the polychaete Pygospio elegans. Parasitology. 140: 855–867. Soldati-Favre D. (2008). Molecular dissection of host cell invasion by the apicomplexans: the glideosome. Parasite 15 (3): 197–205. Soldati D., Foth B.J., Cowman A.F. (2004). Molecular and functional aspects of parasite invasion. Trends Parasitol 20 (12): 567–574.

Soldati D., Meissner M. (2004). Toxoplasma as a novel system for motility. Curr Opin Cell Biol 16 (1): 32–40. Stebbings H., Boe G.S., Garlick P.R. (1974). Microtubules and movement in the archigregarine Selenidium fallax. Cell Tissue Res 348 (3): 331–345. Stokkermans T.J., Schwartzman J.D., Keenan K., Morrissette N.S., Tilney L.G., Roos D.S. (1996). Inhibition of Toxoplasma gondii replication by dinitroaniline herbicides. Exp Parasitol 84 (3): 355–370. Sultan A.A., Thathy V., Frevert U., Robson K.J., Crisanti A., Nussenzweig V., Nussenzweig R.S., Ménard R. (1997). TRAP is necessary for gliding motility and infectivity of Plasmodium sporozoites. Cell 90 (3): 511–522. Tardieux I., Baum J. (2016). Reassessing the mechanics of parasite motility and host-cell invasion. J Cell Biol 214 (5): 507–515. Toso M.A., Omoto C.K. (2007). Gregarina niphandrodes may lack both a plastid genome and organelle. J Eukaryot Microbiol 54 (1): 66–72. Valigurová A. (2012). Sophisticated adaptations of Gregarina cuneata (Apicomplexa) feeding stages for epicellular parasitism. PLoS One 7 (8): e42606. Valigurová A., Hofmannová L., Koudela B., Vávra J. (2007). An ultrastructural comparison of the attachment sites between Gregarina steini and Cryptosporidium muris. J Eukaryot Microbiol 54 (6): 495–510. Valigurová A., Jirků M., Koudela B., Gelnar M., Modrý D., Šlapeta J. (2008). Cryptosporidia: Epicellular parasites embraced by the host cell membrane. Int J Parasitol 38 (8-9): 913–922. Valigurová A., Koudela B. (2008). Morphological analysis of the cellular interactions between the eugregarine Gregarina garnhami (Apicomplexa) and the epithelium of its host, the desert locust Schistocerca gregaria. Eur J Protistol 44 (3): 197-207. Valigurová A., Michalková V., Koudela B. (2009). Eugregarine trophozoite detachment from the host epithelium via epimerite retraction: Fiction or fact? Int J Parasitol 39 (11): 1235-1242. Valigurová A., Paskerova G.G., Diakin A., Kováčiková M., Simdyanov T.G. (2015). Protococcidian Eleutheroschizon duboscqi, an unusual apicomplexan interconnecting gregarines and cryptosporidia. PLoS One 10 (4): e0125063. Valigurová A., Vaškovicová N., Diakin A., Paskerova G.G., Simdyanov T.G., Kováčiková M. (2017). Motility in blastogregarines (Apicomplexa): native and drug- induced organisation of Siedleckia nematoides cytoskeletal elements. PLoS ONE 12

(6): e0179709. Valigurová A., Vaškovicová N., Musilová N., Schrével J. (2013). The enigma of eugregarine epicytic folds: where gliding motility originates? Front Zool 10 (1): 57. Vanderberg J.P. (1974). Studies on the motility of Plasmodium sporozoites. J Protozool 21: 527–537. Vávra J., Small B.E. (1969). Scanning electron microscopy of gregarines (Protozoa, Sporozoa) and its contribution to the theory of gregarine movement. J Protozool 16 (5): 745-757. Vivier E. (1968). L’Organisation ultrastructurale corticale de la gregarine Lecudina pellucida; ses rapports avec l'alimentation et la locomotion. J Protozool 15 (2): 230- 246. Votýpka J., Modrý D., Oborník M., Šlapeta J., Lukeš J. (2017). Apicomplexa. In: J.M. Archibald et al. (Eds.). Handbook of the Protists. Springer International Publishing, 567–624. Wakeman K.C., Heintzelman M.B., Leander B.S. (2014). Comparative ultrastructure and molecular phylogeny of Selenidium melongena n. sp. and S. terebellae Ray 1930 demonstrate niche partitioning in marine gregarine parasites (Apicomplexa). Protist 165 (4): 493–511. Wakeman K.C., Horiguchi T. (2017). Morphology and molecular phylogeny of the marine gregarine parasite Selenidium oshoroense n. sp. (Gregarina, Apicomplexa) isolated from a northwest pacific hydroides Ezoensis okuda 1934 (Serpulidae, Polychaeta). Mar Biodiv 48: 1498. Walker M.H., Lane L., Lee W.M. (1984). Freeze-fracture studies on the pellicle of the eugregarine, Gregarina gamhami (Eugregarinida, Protozoa). J Ultrastruct Res 88 (1): 66-76. Walker M.H., Mackenzie C., Bainbridge S.P., Orme C. (1979). A study of the structure and gliding movement of Gregarina garnhami. J Protozool 26 (4): 566-574. Wetzel D.M., Hakansson S., Hu K., Roos D., Sibley L.D. (2003). Actin filament polymerization regulates gliding motility by apicomplexan parasites. Mol Biol Cell 14: 396–406. Whitelaw J.A., Latorre-Barragan F., Gras S., Pall G.S., Leung J.M., Heaslip A., Egarter S. et al. (2017). Surface attachment, promoted by the actomyosin system of Toxoplasma gondii is important for efficient gliding motility and invasion. BMC Biol 15: 1.

Zhu, G., Marchewka M.J., Keithly J.S. (2000). Cryptosporidium parvum appears to lack a plastid genome. Microbiology 146 (2): 315–321.

9 Original publications and author contribution

This Ph.D. thesis is based on seven manuscripts of the thesis author (Magdaléna Kováčiková, MK). As the original papers listed below are protected by copyright, they are presented in full version in the printed thesis only. The electronic (publicly/freely available) version of the thesis contains links to the papers available on-line.

Paper I Valigurová, A., Paskerova, G.G., Diakin, A., Kováčiková, M., Simdyanov, T.G. (2015). Protococcidian Eleutheroschizon duboscqi, an unusual apicomplexan interconnecting gregarines and cryptosporidia. PLoS One 10 (4): e0125063. DOI: 10.1371/journal.pone.0125063. Publication is available at: https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0125063 MK contributed to the light and electron microscopic analyses and data evaluation, and assisted in processing of CLSM samples.

Paper II Kováčiková M., Simdyanov T.G., Diakin A., Valigurová A. (2017). Structures related to attachment and motility in the marine eugregarine Cephaloidophora cf. communis (Apicomplexa). European Journal of Protistology 59: 1-13. DOI: 10.1016/j.ejop.2017.02.006. Publication is available at: https://www.sciencedirect.com/science/article/pii/S0932473916301559 MK performed the video documentation, light microscopic, electron microscopic and CLSM analyses and data evaluation, and wrote the manuscript.

Paper III Valigurová A., Vaškovicová N., Diakin A., Paskerova G.G., Simdyanov T.G., Kováčiková M. (2017). Motility in blastogregarines (Apicomplexa): Native and drug- induced organisation of Siedleckia nematoides cytoskeletal elements. PLoS One 12 (6): e0179709. DOI: 10.1371/journal.pone.0179709. Publication available at: https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0179709 MK contributed to the material collection, experimental assays, video documentation, light and transmission electron microscopic analyses and data evaluation, and assisted in processing of CLSM samples.

Paper IV Simdyanov T.G., Paskerova G.G., Valigurová A., Diakin A., Kováčiková M., Schrével, J., Gillou L., Dobrovolskij A.A., Aleoshin V.V. (2018). First ultrastructural and molecular phylogenetic evidence from the blastogregarines, an early branching lineage of plesiomorphic Apicomplexa. Protist 169 (5): 697-726. DOI: 10.1016/j.protis.2018.04.006. Publication is available at: https://www.sciencedirect.com/science/article/pii/S1434461018300300 MK contributed to the transmission electron microscopic analysis.

Paper V Paskerova G.G., Miroliubova T.S., Diakin A., Kováčiková M., Valigurová A., Gillou L., Aleoshin V.V., Simdyanov T.G. (2018). Fine structure and molecular phylogeny of two marine gregarines, Selenidium pygospionis n. sp. and S. pherusae n. sp., with notes on the phylogeny of Archigregarinida (Apicomplexa). Protist 169 (6): 826–852. DOI: 10.1016/j.protis.2018.06.004. Publication is available at: https://www.sciencedirect.com/science/article/pii/S143446101830066X MK contributed to the material collection and data evaluation.

Paper VI Kováčiková M., Vaškovicová N., Nebesářová J., Valigurová A. (2018). Effect of jasplakinolide and cytochalasin D on cortical elements involved in the gliding motility of the eugregarine Gregarina garnhami (Apicomplexa). European Journal of Protistology 66: 97-114. DOI: 10.1016/j.ejop.2018.08.006. Publication is available at: https://www.sciencedirect.com/science/article/pii/S0932473918300403 MK performed experimental assays and video documentation, light microscopic, electron microscopic and CLSM analyses and data evaluation, and wrote the manuscript.

Paper VII Kováčiková M., Paskerova G.G., Diakin A., Simdyanov T.G., Vaškovicová N., Valigurová A. (2019). Motility and cytoskeletal organisation in the archigregarine Selenidium pygospionis (Apicomplexa): observations on native and experimentally affected parasites. Parasitology Research 'in press'. DOI: 10.1007/s00436-019-06381-z. Research 'in press'. DOI: 10.1007/s00436-019-06381-z. Publication is available at: http://link.springer.com/article/10.1007/s00436-019-06381-z MK performed experimental assays and video documentation, light microscopic, electron microscopic and CLSM analyses and data evaluation, and wrote the manuscript.