Keywords: Microsporidia, C. elegans, diversity, host-pathogen interaction, host specificity, sensitivity, natural variation

Microsporidia are fungi-related intracellular pathogens that infect a great variety of animals, including the nematode Caenorhabditis elegans. The first microsporidia isolated from wild C. elegans was named Nematocida parisii in 2008. C. elegans and N. parisii have been used as a powerful model for the study of host-pathogen interactions. However, it was unclear how widespread and diverse microsporidia infections are in C. elegans or other related nematodes in the wild.

By sampling rhabditid nematodes worldwide, we established a collection of 47 nematodes that displayed putative microsporidia infections. We characterized molecularly these infections and determined that N. parisii (or N. ironsii) is the most common microsporidia infecting C. elegans in the wild. We further described and named six new Nematocida species. In addition, we defined two new genera of nematode-infecting microsporidia, named Enteropsectra and Pancytospora, which are genetically distinct from Nematocida. Further investigations showed that these microsporidia are diverse in terms of spore size and shape, host tissue tropism, host cell intracellular localization, cellular exit route, host specificity pattern, etc. Overall, these findings illustrate the widespread and diverse microsporidia infections in C. elegans and related nematodes in the wild.

We further assayed the natural variation of C. elegans in sensitivity to N. ausubeli infection, by comparing 10 C. elegans strains using food consumption tests. Two C. elegans strains, JU1249 and JU2825, displayed the largest sensitivity differences, which were suggested to be a result of the different tolerance between the two strains.

These two strains are proven to be good candidates for future studies on the genetic loci associated with C. elegans sensitivity variation to microsporidian infections. Furthermore, I observed an exciting effect of host-pathogen interaction. Microsporidia infection is able to suppress the progressive decline in fertility in some C. elegans with the mortal germline phenotype (Mrt).

RESUME

Les microsporidies sont des pathogènes intracellulaires obligatoires apparentés aux champignons. Elles infectent de nombreux animaux, dont le nématode Caenorhabditis elegans. La première microsporidie isolée d'une souche de C. elegans sauvage a été nommée Nematocida parisii. L'interaction entre N. parisii et C. elegans est devenue un puisant modèle pour l'étude des interactions hôte-pathogène.

Cependant, ce modèle a été récemment découvert et de nombreux détails sur son écologie et sa biologie restaient inconnus. Notamment, nous ignorions l'incidence et la diversité des infections microsporidiennes chez C. elegans et autres nématodes dans la nature.

A partir d'une collection de nématodes, de la famille des Rhabditidae, échantillonnés dans le monde entier, j'ai recensé un panel de 47 nématodes présentant des symptômes d'infection par des microsporidies. J'ai caractérisé moléculairement la diversité de ce parasite infectant ces nématodes et déterminé que N. parisii est la microsporidie la plus souvent responsable des infections chez C. elegans dans la nature. J'ai également décrit et nommé six nouvelles espèces de Nematocida. Au cours de mes travaux, j'ai aussi défini deux nouveaux genres de microsporidies génétiquement distincts de Nematocida, appelés Enteropsectra et Pancytospora. Mes travaux ont de plus détaillé la diversité qui existe chez les microsporidies parasites de nématodes. Ces microsporidies présentent des différences en terme de taille et forme de leurs spores, de leur tropismes tissulaire et intracellulaire chez l'hôte, de leur voie de sortie des cellules hôtes mais aussi de spectre d'hôtes. Mes résultats ont démontré que, dans la nature, les infections de C. elegans et autres nématodes par les microsporidies sont répandues et diverses.

De plus, j'ai estimé la variation naturelle pour la sensibilité de C. elegans à l'infection par N. ausubeli. J'ai notamment comparé 10 souches naturelles de C. elegans en utilisant des tests de consommation alimentaire. Deux souches de C. elegans, JU1249 et JU2825, présentaient des niveaux contrastés de sensibilité, ce que j'ai interprété comme étant une différence de niveau de tolérance aux infections. Ces deux souches se sont révélées être de bons candidats pour une future caractérisation des loci génétiques associés à la variation de sensibilité de C. elegans aux infections microsporidiennes. Enfin, j'ai observé un effet surprenant de l'infection de C. elegans par les microsporidies. En effet, la présence du pathogène est capable de supprimer le

Introduction

Organisms are driven to evolve continuously by selective pressures from the everchanging environment [START_REF] Darwin | On the origins of species by means of natural selection[END_REF]. The natural environment includes abiotic and biotic factors. Abiotic factors refer to physical and chemical parameters, such as light, water, temperature. Biotic factors describe all the intraspecific and interspecific interactions between organisms, including those interacting preys and predators, hosts and pathogens, competitors, mutualists. [START_REF] Woolhouse | Biological and biomedical implications of the co-evolution of pathogens and their hosts[END_REF]Morgan and Koskella, 2011). Though both abiotic and biotic factors could provide selective pressures on species, the latter were highlighted as a fundamental driver of the perpetual evolutionary change [START_REF] Van Valen | A new evolutionary law[END_REF][START_REF] Brockhurst | Running with the Red Queen: the role of biotic conflicts in evolution[END_REF]). The interacting species in biotic interactions may keep adapting to each other and coevolve antagonistically. Among the diverse biotic interactions, host-pathogen interactions probably provide the best case to study counter-adaptations and antagonistic coevolution.

Several host model organisms, such as mice, zebrafish, Arabidopsis thaliana, Caenorhabditis elegans, Drosophila melanogaster have been used to study the evolutionary consequences, the mechanisms and the molecular basis of host-pathogen interactions. Among these model organisms, the nematode C. elegans has its unique advantages to facilitate studies on hosts and pathogens. Major attributes of C. elegans include the small size, the simple growth conditions, the rapid generation time, the genetic tractability and the availability of a great number of genetic mutants [START_REF] Sulston | The embryonic cell lineage of the nematode Caenorhabditis elegans[END_REF][START_REF] Kamath | Systematic functional analysis of the Caenorhabditis elegans genome using RNAi[END_REF]Stiernagle, 2006). In the last 50 years, C. elegans has proven to be a powerful tool for studies on basic biological processes, as well as for investigations on host-pathogen interactions [START_REF] Kurz | Caenorhabditis elegans for the study of host-pathogen interactions[END_REF][START_REF] Zhang | Host-Microbe Interactions in Caenorhabditis elegans[END_REF][START_REF] Cohen | Microbial pathogenesis and host defense in the nematode C. elegans[END_REF].

C. elegans has been found susceptible to a wide variety of pathogens, including several human pathogens. The first identified natural intracellular pathogen of C. elegans is Nematocida parisii, which defined a new genus and species of microsporidia (Troemel et al. 2008). The Microsporidia are obligate pathogens with over 1,400 described species that can infect about half of the metazoan phyla [START_REF] Vavra | Microsporidia and "The Art of Living Together[END_REF]. Microsporidian infections were found quite prevalent in wild Caenorhabditis and related nematodes (Troemel et al. 2008;[START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF].

Together, C. elegans and microsporidia provide an ideal model for the study of hostpathogen interactions and coevolution.

In the Introduction, first, I shall briefly present the origin, morphology and life cycle of microsporidia; In the second part, I will present the natural history of C. elegans and related nematodes, and the interactions of C. elegans with diverse microbes in the natural habitat; third, I will describe the previous and recent studies on the interactions between C. elegans and Nematocida parisii; in the last part, I will present the background of my PhD project.

I. What are Microsporidia?

The Microsporidia are obligate intracellular parasites that are related to fungi. As a large group of strictly intracellular pathogens, microsporidia species have evolved to be highly adapted to the parasitic life style. They are highly reduced in morphology, metabolism and genome [START_REF] Vavra | Structure of Microsporidia[END_REF]. Within the host cytoplasm (or the nuclear), microsporidia proliferate rapidly and generate large numbers of infectious spores. Spores are the life stage that survives outside hosts, even in harsh conditions. Microsporidia are typically transmitted horizontally, while other transmission modes also exist [START_REF] Cali | Developmental morphology and life cycles of the microsporidia[END_REF]. Microsporidia use the polar tube, a unique highly specialized invasion structure inside the spore, to inject their sporoplasms (the cytoplasmic content and the nucleus of the spore) into the intracellular milieu of their hosts [START_REF] Vavra | Structure of Microsporidia[END_REF]. Microsporidia are found to be pathogens to a vast variety of hosts, ranging across about half of metazoan phyla [START_REF] Vavra | Microsporidia and "The Art of Living Together[END_REF]. Microsporidia infections can cause serious damages in veterinary and agricultural settings, as well as in immunocompromised humans. Furthermore, as one of the most frequently observed pathogens of animals, microsporidia might play significant roles in evolutionary processes [START_REF] Altizer | Rapid evolutionary dynamics and disease threats to biodiversity[END_REF][START_REF] Vijendravarma | Experimental evolution shows Drosophila melanogaster resistance to a microsporidian pathogen has fitness costs[END_REF].

I.1. Microsporidia are diverse and important parasites

The study of microsporidia began in the 19 th century, when an epidemic disease destroyed the silkworm industry across Europe (Franzen, 2008). Pasteur observed microsporidian spores in the affected silkworms and helped to save the silkworm industry (Franzen, 2008). Today, over 200 genera and over 1,400 species in the microsporidia phylum have been described, and new species are being identified each year [START_REF] Becnel | Checklist of available generic names for microsporidia with type species and type hosts[END_REF]Szumowski and Troemel, 2015). Microsporidia are ubiquitous pathogens infecting a large variety of hosts, including all vertebrate orders, most invertebrates and protists (Keeling and Fast, 2002).

Many microsporidia are of economic importance. Microsporidia infections in some beneficial insects such as honey bees and silkworms cause significant economic losses [START_REF] Fries | Microsporidia, honeybees, and colony collapse disorder[END_REF]. Fish is another type of microsporidia host with economic importance: to date, 120 microsporidia species have been isolated from fish [START_REF] Kent | Microsporidia in fish[END_REF]. Livestock such as cattle, pigs and goats, pets such as dogs, cats and birds have all been reported to carry microsporidia infections [START_REF] Fayer | Epidemiology of microsporidia in human infections[END_REF]. More importantly, microsporidia are also opportunistic pathogens to human.

To date, 17 microsporidia species are known to infect humans [START_REF] Fayer | Epidemiology of microsporidia in human infections[END_REF]. The most clinically significant microsporidia species is Enterocytozoon bieneusi, which infects the epithelium of the gastrointestinal tract and causes chronic diarrhea and wasting in HIV patients [START_REF] Akiyoshi | Genomic survey of the noncultivatable opportunistic human pathogen, Enterocytozoon bieneusi[END_REF]. Human microsporidiosis cases were also known to cause ocular infection, nervous system infection, musculoskeletal infection, etc. [START_REF] Weiss | Clinical syndromes associated with microsporidiosis[END_REF]. Taken together, microsporidia are pathogens with considerable economic and clinical importance.

I.2. Origin of microsporidia

It has long been challenging to place microsporidia in the tree of eukaryotes: microsporidia are highly reduced at many levels compared to other eukaryotes, but they possess a unique complex polar tube as the infection apparatus [START_REF] Keeling | Phylogenetic place of Microsporidia in the tree of Eukaryotes[END_REF].

The debate about the phylogenetic placement of microsporidia has been going on for decades. Firstly, microsporidia have been proposed to be members of Cnidosporidia, based on their similarities of infection machineries with Helicosporidia and Myxosporidia. However, the similarities previously known as "homologous" features were later found to be the result from convergent evolution. Secondly, based on their lack of some fundamental eukaryotic characters (including mitochondria, 9 + 2 microtubule structures) and the phylogeny of eukaryotic microbes using small subunit ribosomal DNA (SSU rDNA), microsporidia were suggested to be very ancient eukaryotes, a lineage of Archezoa. However, several mitochondrial-derived genes and mitochondrial remnant "mitosomes" were later found in microsporidia, suggesting that mitochondria went through reduction in the microsporidia lineage [START_REF] Germot | Presence of a mitochondrial-type 70-kDa heat shock protein in Trichomonas vaginalis suggests a very early mitochondrial endosymbiosis in eukaryotes[END_REF][START_REF] Katinka | Genome sequence and gene compaction of the eukaryote parasite Encephalitozoon cuniculi[END_REF][START_REF] Keeling | Phylogenetic place of Microsporidia in the tree of Eukaryotes[END_REF][START_REF] Corradi | Microsporidia: eukaryotic intracellular parasites shaped by gene loss and horizontal gene transfers[END_REF]. In addition, new phylogenetic analysis based on several protein-coding genes revealed that microsporidia are linked to fungi, which was later supported by phylogenomic analysis [START_REF] Keeling | Alpha-tubulin from early-diverging eukaryotic lineages and the evolution of the tubulin family[END_REF][START_REF] Fast | Phylogenetic analysis of the TATA box binding protein (TBP) gene from Nosema locustae: evidence for a microsporidia-fungi relationship and spliceosomal intron loss[END_REF][START_REF] Capella-Gutiérrez | Phylogenomics supports microsporidia as the earliest diverging clade of sequenced fungi[END_REF]. Now, it is generally accepted that microsporidia are related to fungi [START_REF] Keeling | Phylogenetic place of Microsporidia in the tree of Eukaryotes[END_REF].

However, the problem about the phylogenetic placement of microsporidia has not been solved completely: what is the exact nature of the relationship between fungi and microsporidia? Are microsporidia a subclade of fungi or a sister clade to fungi? Within the fungal phylogeny, Rozella, a monospecific fungal genus, was placed together with microsporidia as a group in a deep-branching position [START_REF] Jones | Discovery of novel intermediate forms redefines the fungal tree of life[END_REF][START_REF] James | Shared signatures of parasitism and phylogenomics unite Cryptomycota and microsporidia[END_REF]. This Rozella-Microsporidia group might be considered "sister" to other fungi species. Currently, microsporidia are suggested to be members of Cryptomycota, a hyperdiverse phylum encompassing Rozella, Aphelids and probably a large number of poorly described species (Figure 1) [START_REF] James | Shared signatures of parasitism and phylogenomics unite Cryptomycota and microsporidia[END_REF][START_REF] Corsaro | Microsporidia-like parasites of amoebae belong to the early fungal lineage Rozellomycota[END_REF][START_REF] Keeling | Phylogenetic place of Microsporidia in the tree of Eukaryotes[END_REF]. The phylogeny is based on the analysis of multiple protein-coding genes. Adapted from [START_REF] Keeling | Phylogenetic place of Microsporidia in the tree of Eukaryotes[END_REF].

I.3. Structure and life cycles of microsporidia

I.3.1. Spore structure

Spore characters formed the basis of microsporidia species recognition and classification. Most microsporidia spores are only a few micrometers long, and can be oval, pyriform, rodlike or spherical in shape [START_REF] Vavra | Structure of Microsporidia[END_REF]. The spore has a thick resistant wall with two layers, called exospore and endospore (Figure 2A,B). Besides the cytoplasmic content and an individual nucleus (or two apposed nuclei) of the spore, the other important part within a spore is the infection apparatus, which is unique to microsporidia and has no obvious homolog in other organisms [START_REF] Keeling | Phylogenetic place of Microsporidia in the tree of Eukaryotes[END_REF]. The infection apparatus system consists in a polar tube, a polaroplast and a posterior vacuole [START_REF] Weiss | Microsporidia-Pathogens of opportunity[END_REF]. The polar tube is threadlike and usually much longer than the length of the spore (Figure 2C). One end of the polar tube is connected to a mushroom-shaped anchoring disk (AD) at the anterior end of the spore. The rest of the polar tube coils around the polaroplast. The number of spore coils has been an important character in species description (Figure 2A) [START_REF] Weiss | Microsporidia-Pathogens of opportunity[END_REF]. With this unique apparatus, microsporidia are able to inoculate their hosts efficiently in only a few seconds [START_REF] Frixione | Dynamics of polar filament discharge and sporoplasm expulsion by microsporidian spores[END_REF]. b,c). Abbreviations are the same as in A., with, in addition: f, polar tube; Pa, anterior polaroplast region; Pp, posterior polaroplast region; Ps, polar sac; R, polyribosomes. C. Scanning electron micrograph of extruded polar tube of a spore of Encephalitozoon intestinalis piercing and infecting Vero E6 green monkey kidney cells in tissue culture. Adapted from [START_REF] Vavra | Structure of Microsporidia[END_REF]; [START_REF] Weiss | Microsporidia-Pathogens of opportunity[END_REF].

I.3.2. Life cycles

The life cycle of microsporidia includes extracellular and intracellular phases. In the extracellular phase, microsporidia spores are typically transmitted horizontally through host ingestion and normally infect the hosts through the gut [START_REF] Vavra | Structure of Microsporidia[END_REF]. With chemical and/or physical stimulations, spores germinate.

The polaroplast is activated to initiate the evagination of the polar tube. The polar tube forms a channel by extruding from the spore and penetrating into the host cell membrane. At the same time, typically, the posterior vacuole expands to push the infective sporoplasm through the polar tube, into the host cytoplasm [START_REF] Franzen | Microsporidia: how can they invade other cells?[END_REF][START_REF] Cali | Developmental morphology and life cycles of the microsporidia[END_REF][START_REF] Vavra | Structure of Microsporidia[END_REF]. Then microsporidia start the intracellular phase of their life cycle.

The intracellular microsporidia develop through the meront stage and the sporont stage, until the formation of mature spores (Figure 3). The sporoplasm injected into the host cell first develops into a meront. Meronts may go through nuclear division and immediate cell division by binary fission. Without immediate cytokinesis, multinucleate meronts will form by multiple rounds of nuclear division in one cell, and eventually divide by plasmotomy, to cells containing one nucleus or double paired nuclei (called diplokaryon). Once meronts have secreted electron-dense material on the external face of the plasma membrane, they enter the sporont stage.

The electron-dense material keeps accumulating on the outer face of the sporont, and forms a thickened wall. Depending on the microsporidia species, sporonts may divide once or many times. Multinucleate cells may also form and divide by plasmotomy at this stage. The final division product of the sporont is called sporoblast. The sporoblast continues to grow and becomes a mature spore [START_REF] Cali | Developmental morphology and life cycles of the microsporidia[END_REF][START_REF] Vavra | Structure of Microsporidia[END_REF].

I.4. The five clades in microsporidia

With a phylogeny based on SSU rDNA, five major clades within microsporidia have been proposed (Figure 4) [START_REF] Vossbrinck | Phylogeny of the Microsporidia[END_REF]. Though exceptions exist, the groupings of species in different clades largely correspond to their hosts and habitats (Figure 4) [START_REF] Vossbrinck | Phylogeny of the Microsporidia[END_REF]. Clade I and Clade III mainly contain microsporidia with hosts from freshwater; Clade II and Clade IV microsporidia have terrestrial animals as their hosts; Clade V microsporidia were primarily isolated from organisms living in marine and freshwater habitats (Figure 4) [START_REF] Vossbrinck | Phylogeny of the Microsporidia[END_REF]. The phylogeny is based on SSU rDNA. Font color indicates the host's habitat: blue for freshwater, brown for terrestrial, green for marine and blue green for hosts that spend part of their life cycle in both marine and freshwater habitats. Nematocida parisii, the natural pathogen of C. elegans, is marked by a red frame. Adapted from [START_REF] Vossbrinck | Phylogeny of the Microsporidia[END_REF].

II. Caenorhabditis elegans and other related nematodes

The nematode Caenorhabditis elegans is a small, free-living nematode that lives across many of the humid temperate areas of the world (Kiontke et al., 2011;[START_REF] Andersen | Chromosome-scale selective sweeps shape Caenorhabditis elegans genomic diversity[END_REF]Frézal and Félix, 2015). In the 1960s, Sydney Brenner first adopted C. elegans as a model organism for investigation of a great variety of biological phenomena [START_REF] Brenner | The genetics of Caenorhabditis elegans[END_REF]. Brenner took the Bristol strain N2 as the standard C. elegans reference strain and cultured it on agar plates seeded with Escherichia coli in the lab [START_REF] Brenner | The genetics of Caenorhabditis elegans[END_REF][START_REF] Ankeny | The natural history of Caenorhabditis elegans research[END_REF]. Ever since then, C. elegans has proven to be a powerful tool to study basic biological processes such as apoptosis and gene silencing, as well as biomedical issues such as Alzheimer's disease [START_REF] Drake | Oxidative stress precedes fibrillar deposition of Alzheimer's disease amyloid β-peptide (1-42) in a transgenic Caenorhabditis elegans model[END_REF][START_REF] Kaletta | Finding function in novel targets: C. elegans as a model organism[END_REF][START_REF] Seth | The C. elegans CSR-1 argonaute pathway counteracts epigenetic silencing to promote germline gene expression[END_REF][START_REF] Yee | The intrinsic apoptosis pathway mediates the prolongevity response to mitochondrial ROS in C. elegans[END_REF]Frézal and Félix, 2015).

The natural ecology of C. elegans has also become an emerging research area [START_REF] Félix | The natural history of Caenorhabditis elegans[END_REF]Frézal and Félix, 2015;Petersen et al., 2015). In the natural habitat of C. elegans and other related nematodes, biotic factors, in particular the pathogens, may significantly shape the evolutionary process of the nematodes. In this part, I will first present the natural history of C. elegans and other related nematodes; then I will describe the interactions of C. elegans with its natural pathogens.

II.1. Natural history of C. elegans II.1.1. Natural habitat

The natural history of C. elegans was a mystery for a long time. During the first phase, from the 20 th century to the beginning of the 21 st century, C. elegans had been regarded and presented as "a soil nematode". In fact, only a few C. elegans isolates Probably due to the fact that the food is ephemeral and patchily distributed in space, the C. elegans populations have a boom-and-bust life style in the wild [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]Frézal and Félix, 2015). With a successful migration to a food source, a single hermaphrodite dauer may bloom into a huge population.

II.1.2. Sex and reproduction

C. elegans has two sexes: the self-fertilizing hermaphrodites (XX) and the facultative males (XO) (Figure 6). Adult males are slender and slightly shorter than adult hermaphrodites (Figure 6A,C). A selfing hermaphrodite can produce about 300 progenies, of which in average about 0.1% are males and all the other are hermaphrodites. Through mating, a hermaphrodite and a male can produce more than 1000 offsprings, with half male progeny and half hermaphrodite progeny [START_REF] Zarkower | Somatic sex determination[END_REF]. Selfing of hermaphrodites allows the maintenance of homozygous C. elegans through generations, whereas mating of the two sexes facilitates genetic variation by recombination. Homozygosity can resume after some rounds of selfing. capable of laying eggs, or becoming an adult male capable of fertilizing hermaphrodite adults (Figure 7). Then the organogenesis and morphogenesis stage will last for 6 to 8 hours, after which the embryo develops into an animal with fully differentiated tissues and organs.

Here, the new animal is able to move and pump inside the eggshell. At about 800 minutes, the embryo hatches [START_REF] Hall | Introduction[END_REF].

II.1.3.2. Post-embryonic stage

Food is vital for all the processes in all the stages in the post-embryonic development of C. elegans. In the presence of enough food, the cell division resumes in 3 hours post-hatching [START_REF]Control of developmental timing in Caenorhabditis elegans[END_REF][START_REF] Hall | Introduction[END_REF]. The animal passes four larval stages (L1-L4) and reaches the adult stage in 40-50 hours at 20-25°C [START_REF] Hall | Introduction[END_REF]. During the larval stages, additional cell divisions of blast cells occur and different systems develop. The larva molts at the end of each stage, with a stagespecific cuticle being shed and resynthesized.

Without food, newly hatched larvae can survive for 6-10 days, but their development is arrested. After food becomes available, these arrested L1 larvae can return to normal development. The dauer larva stage, the other arrested state and an alternative L3 developmental stage of C. elegans, occurs if the environmental conditions are not favorable (Figure 7) [START_REF] Cassada | The dauer larva, a post-embryonic developmental variant of the nematode Caenorhabditis elegans[END_REF][START_REF] Hu | The C. elegans Research Community[END_REF]. Limited food source and high population density are the two most significant factors to induce dauers. The dauer larva is very thin and has a thick cuticle that resists to harsh environments.

Dauers are capable of surviving for several months without food [START_REF] Cassada | The dauer larva, a post-embryonic developmental variant of the nematode Caenorhabditis elegans[END_REF]. The active nictating behavior of dauers facilitates their migration to new food sources. When the environment becomes favorable for further growth, dauers first undergo the L3 development events, then molt to L4 and continue to develop into reproductive adults. If starvation happens in the L4 stage, some C. elegans may enter the adult reproductive diapause (ARD) state. In the reproductive period in adulthood, if food is absent, the adult hermaphrodites stop laying eggs and may undergo a "bagging" process, in which the embryos hatch inside the uterus and the newly hatched larvae consume the hermaphrodite parent [START_REF] Angelo | Starvation protects germline stem cells and extends reproductive longevity in C. elegans[END_REF][START_REF] Seidel | The oogenic germline starvation response in C. elegans[END_REF].

II.1.4. Basic anatomy of C. elegans

C. elegans is a tiny roundworm, only 1 mm as an adult. It has an unsegmented, cylindrical transparent body that tapers at both ends. There are several tissues inside the body of C. elegans, including the epidermis, the muscles, the nervous system, the alimentary system, the reproductive system (Figures 6,8) [START_REF] Altun | Alimentary system, intestine[END_REF]Corsi et al., 2015). Among the different tissues of C. elegans, the epidermis and the intestine are normally used by pathogens as invading routes and tropisms (details in II.3.2.). Here, I will describe the anatomy of the alimentary system in more detail. The alimentary tract of C. elegans consists of the pharynx, the intestine, the rectum and the anus (Figure 6B). C. elegans is a filter-feeder. It pumps food such as bacteria into the pharyngeal lumen. Then the posterior bulb of the pharynx grinds bacteria and transports them into the intestinal lumen (Figure 9A). This grinding behavior may protect the C. elegans from some pathogens, as defective grinder mutant C. elegans seemed more susceptible to pathogen-induced death (Aballay and Ausubel, 2002).

The intestine is a long tube with a lumen inside, which are formed by nine intestinal rings (int I-IX) of twenty large epithelial cells (Figure 9B,C). The anteriormost ring I is comprised of four cells while each of the remaining eight rings consists of two cells (Figure 9B). The intestinal cells are polarized with two plasma membrane domains: the apical domain and the basolateral domain [START_REF] Mcghee | The C. elegans intestine[END_REF][START_REF] Sato | C. elegans as a model for membrane traffic[END_REF].

Similar to the morphological characteristics of mammalian intestinal cells, many microvilli extend into the lumen from the apical side of C. elegans intestinal cells (Figure 9D,E). This brush border of microvilli is anchored in a conserved structure called the terminal web of actin (with ACT-5 protein) and intermediate filaments (with IFB-2 protein) [START_REF] Macqueen | ACT-5 is an essential Caenorhabditis elegans actin required for intestinal microvilli formation[END_REF][START_REF] Mcghee | The C. elegans intestine[END_REF]).

The primary function of the intestine is food digestion and nutrient absorption.

Nutrients are transferred into intestinal cells through the apical membranes, by endocytosis or special transporters [START_REF] Mcghee | The C. elegans intestine[END_REF]. Numerous granules in intestinal cells store nutrients such as lipids, proteins and carbohydrates. As the intestine is a major route for pathogens to invade into the body of C. elegans, the intestinal cells can also act in the defense against pathogens by shedding microvilli and activating the innate immunity pathways (Schulenburg et al., 2004;[START_REF] Mcghee | The C. elegans intestine[END_REF]. The rectum and the anus are at the end of the digestive tract, where the final intestinal content wastes are expelled to the outside (Figure 6).

Figure 9. The anatomy of pharynx and intestine in C. elegans.

A. Graphic rendition of the pharynx. The posterior bulb, also called the terminal bulb, is connected to the intestine through the pharyngeal-intestinal valve. B. Intestinal rings in adult. The intestine has a left-handed twist of 180° around the longitudinal body axis. Cells in the nine rings are indicated depending on positions in the body plan: Left (L), Right (R), Dorsal (D), Ventral (V). C. Epifluorescent image of an adult transgenic animal expressing the reporter gene emr-1::GFP, where the emr-1 gene encodes a homolog of the human integral nuclear membrane protein emerin [START_REF] Howe | WormBase 2016: expanding to enable helminth genomic research[END_REF]. D. Ultrastructure of the third intestinal cell ring. A complex junction (arrow) next to an adherens junction (arrowhead) seals the two intestinal cells to each other (inset). E. Diagram of a cross section through an intestinal cell. Adapted from Altun andHall (2009a, 2009b).

II.1.5. A model organism in the wild

Researchers have made fruitful discoveries with the C. elegans reference strain N2 or mutants in the N2 genetic background. However, as N2 has been maintained under similar laboratory conditions for several decades, this strain has evolved and become distinct from its wild progenitor [START_REF] Sterken | The laboratory domestication of Caenorhabditis elegans[END_REF]. For instance, in the standard culture plates, wild C. elegans display a clumping behavior: wild isolates tend to aggregate with each other, especially on the border of the E. coli lawn (Figure 10, right) [START_REF] Gray | Oxygen sensation and social feeding mediated by a C. elegans guanylate cyclase homologue[END_REF]Barrière and Félix, 2005a). Wild C. elegans clump together, because they prefer a 7-14% oxygen concentration in the microenvironment. By clumping at the border of the bacterial lawn, they can reduce the local O 2 concentration [START_REF] Cheung | Experiencedependent modulation of C. elegans behavior by ambient oxygen[END_REF][START_REF] Rogers | Behavioral motifs and neural pathways coordinating O2 responses and aggregation in C. elegans[END_REF]. However, N2 does not show this clumping behavior (Figure 10,left). This phenotypic polymorphism was further attributed to a single nucleotide polymorphism on the npr-1 (gene npr-1 encodes a neuropeptide Y-like G-protein-coupled receptor) (de Bono and Bargmann, 1998; Barrière and Félix, 2005a;[START_REF] Mcgrath | Quantitative mapping of a digenic behavioral trait implicates globin variation in C. elegans sensory behaviors[END_REF].

To have a more comprehensive understanding of the C. elegans species, other wild C. elegans strains have been used. The most widely studied natural C. elegans strain is CB4856, which was collected from Hawaii. This strain is genetically distant to the N2 strain. CB4856 also shows a great variety of polymorphisms compared to other strains [START_REF] Koch | Single nucleotide polymorphisms in wild isolates of Caenorhabditis elegans[END_REF][START_REF] Andersen | Chromosome-scale selective sweeps shape Caenorhabditis elegans genomic diversity[END_REF][START_REF] Vergara | Genome-wide variations in a natural isolate of the nematode Caenorhabditis elegans[END_REF]. By comparing different strains, the genetic basis of several traits has been found. For instance, after mating, males of CB4856 and some other natural C. elegans isolates deposit a copulatory plug that covers the hermaphrodite vulva, which is known as the plugging phenotype. The plug can decrease the mating success of subsequent males. However, some other wild C. elegans strains and N2 display the non-plugging phenotype.

Mapping approaches between CB4856 and N2 revealed that the plugging phenotype variation results from the polymorphism of the gene plg-1 [START_REF] Hodgkin | Natural variation and copulatory plug formation in Caenorhabditis elegans[END_REF][START_REF] Palopoli | Molecular basis of the copulatory plug polymorphism in Caenorhabditis elegans[END_REF]. Barrière and Félix (2005a).

II.2. Other nematodes

C. elegans is the best-known nematode species, but the phylum Nematoda is an extremely diverse group of animals (Figure 11). About 30,000 species have been described in Nematoda, the total number of which is estimated up to a million [START_REF] Kiontke | Nematodes[END_REF]. Nematodes are probably the most highly ecologically and physiologically adaptable metazoans, since they were found everywhere, including desert soil, polar ice, tropical rainforest and deep sea [START_REF] Félix | Species richness, distribution and genetic diversity of Caenorhabditis nematodes in a remote tropical rainforest[END_REF][START_REF] Kiontke | Nematodes[END_REF]. Nematodes are not just free-living species like C. elegans. Pathogenic nematodes can infect almost all other multicellular organisms, including most plants, arthropods and vertebrates [START_REF] Blaxter | Nematodes: the worm and its relatives[END_REF][START_REF] Kiontke | Nematodes[END_REF]. I will further describe some free living nematodes that are closely related to C. elegans. The position of nematodes in the animal tree is indicated in the inset. The main panel is the phylogeny of Nematoda, which contain three large monophyletic groups: Enoplia, Dorylaimia and Chromadoria. C. elegans belongs to the infraorder Rhabditomorpha (suborder Rhabditina, order Rhabditida, family Rhabditidae), which is indicated by an arrow. The phylogeny of Nematoda is based on SSU rDNA. Adapted from [START_REF] Kiontke | Nematodes[END_REF].

II.2.1. Free-living rhabditid nematodes

C. elegans and its close relatives are in the family Rhabditidae (Figures 11,[START_REF] Ma | C. elegans outside the Petri dish[END_REF]. Most rhabditids are bacteriophagous nematodes, but this group also includes a group of vertebrate parasites such as the human hook worm Necator americanus [START_REF] Kiontke | The phylogenetic relationships of Caenorhabditis and other rhabditids[END_REF][START_REF] Blaxter | Nematodes: the worm and its relatives[END_REF]. In addition to C. elegans, this group also includes several "satellite models", such as various other Caenorhabditis species, Oscheius tipulae and Pristionchus pacificus, which facilitate comparative biological studies [START_REF] Kiontke | The phylogenetic relationships of Caenorhabditis and other rhabditids[END_REF]De Ley, 2006). There are two major clades in rhabditids: "Pleiorhabditis" and "Eurhabditis", and the latter clade contains Caenorhabditis and Oscheius (Figure 12) [START_REF] Kiontke | The phylogenetic relationships of Caenorhabditis and other rhabditids[END_REF].

Figure 12. Phylogeny of rhabditid nematodes.

There are two major clades in rhabditids: Pleiorhabditis and Eurhabditis. The latter clade includes the Oscheius genus and the Caenorhabditis genus. The phylogeny is based on SSU rDNA and large subunit ribosomal DNA (LSU rDNA). From [START_REF] Kiontke | The phylogenetic relationships of Caenorhabditis and other rhabditids[END_REF].

II.2.2. The Caenorhabditis genus

Due to the improved knowledge on the wild micro-habitats of Caenorhabditis and intensive worldwide sampling, many new C. elegans isolates and new Caenorhabditis species have been discovered. To date, 27 Caenorhabditis species have been isolated from different areas around the world (Figure 13) (Kiontke et al., 2011;[START_REF] Félix | A streamlined system for species diagnosis in Caenorhabditis (Nematoda: Rhabditidae) with name designations for 15 distinct biological species[END_REF]Cutter, 2015;Frézal and Félix, 2015). Some Caenorhabditis are cosmopolitan:

Caenorhabditis briggsae is the most widespread, found abundantly in both tropical and temperate areas; most C. elegans and all Caenorhabditis remanei have been found in temperate regions (Figure 13). Caenorhabditis brenneri and Caenorhabditis tropicalis have been found exclusively in tropical areas (Kiontke et al., 2011;Cutter, 2015). Among these 27 species, some species have been found in the same location, for instance, C. elegans and C. briggsae have been found coexisting, even in the same apple in Orsay, France [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]; C. elegans and C. remanei have been found coexisting in North Germany [START_REF] Petersen | The prevalence of Caenorhabditis elegans across 1.5 years in selected North German locations: the importance of substrate type, abiotic parameters, and Caenorhabditis competitors[END_REF].

The species in the genus Caenorhabditis can be divided into three groups: Elegans group, Drosophilae group and Japonica group (Figure 13). Most collected strains belong to the Elegans group. C. elegans and C. briggsae are the two most highly sampled species (Figure 13) (Cutter, 2015). These two species are quite similar in morphology, life style, behavior and genome (about 104Mb) [START_REF] Stein | The genome sequence of Caenorhabditis briggsae: a platform for comparative genomics[END_REF][START_REF] Baird | Caenorhabditis briggsae methods[END_REF][START_REF] Gupta | Genomics and biology of the nematode Caenorhabditis briggsae[END_REF]. Nevertheless, there are many species-specific behaviors and genes between C. elegans and C. briggsae. For instance, similar to N2, C. briggsae has a non-clumping behavior on standard culture plates [START_REF] Barrière | Isolation of C. elegans and related nematodes[END_REF].

Relatively, C. briggsae prefers higher temperature, while C. elegans prefers lower temperature [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]. This temperature preference of C. briggsae may due to a recent expansion of C. briggsae from tropical to temperate areas, which provides a good example to study species distribution, gene flow and genome evolution [START_REF] Baird | Caenorhabditis briggsae methods[END_REF][START_REF] Cutter | Patterns of nucleotide polymorphism distinguish temperate and tropical wild isolates of Caenorhabditis briggsae[END_REF]Cutter et al., , 2010aCutter et al., , 2010b)). The main panel shows the phylogeny of the major Caenorhabditis groups with 27 species (former numerical identifiers in parentheses). The three species with selfing mode of reproduction are in red, while the remaining known Caenorhabditis species are male and female. Geographic sampling locations and number of isolated strains are shown on the right of the phylogeny. Adapted from Cutter (2015); Frézal and Félix (2015).

II.2.3. Oscheius tipulae

Oscheius tipulae is another model organism. Morphologically, the animals have a darker color, smaller size and longer rectum, compared to C. elegans [START_REF] Félix | The C. elegans Research Community[END_REF][START_REF] Barrière | Isolation of C. elegans and related nematodes[END_REF]. O. tipulae shares many similarities with C. elegans, such as the reproduction mode of self-fertilizing hermaphrodites and facultative males, and the culture condition in the laboratory. It has been used in comparative studies with C. elegans, especially on vulva development [START_REF] Félix | Comparative developmental studies using Oscheius/Dolichorhabditis sp. CEW1 (Rhabditidae)[END_REF][START_REF] Dichtel-Danjoy | The two steps of vulval induction in Oscheius tipulae CEW1 recruit common regulators including a MEK kinase[END_REF].

Except for very dry or very cold areas, O. tipulae is widely distributed at different spatial scales all over the world. Dauer stage O. tipulae was found abundant in soil, composts and invertebrates (Baïlle et al., 2008); O. tipulae in proliferative stages was also found in rotting fruits and stems [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]. O. tipulae has two morphologically indistinguishable close relatives: Oscheius sp. 2 and Oscheius sp. 3 (Figure 14) [START_REF] Félix | Three biological species closely related to Rhabditis (Oscheius) pseudodolichura Körner in Osche[END_REF]. However, these two Oscheius species were only found in limited locations [START_REF] Félix | Three biological species closely related to Rhabditis (Oscheius) pseudodolichura Körner in Osche[END_REF]Baïlle et al., 2008). Interestingly, a study on the natural populations of O. tipulae showed a relatively high genetic diversity, which may be better suited for further studies on molecular evolution (Baïlle et al., 2008). From Félix (2006).

II.3. Interactions of C. elegans with diverse microorganisms

In the wild, C. elegans has a great variety of interactions with several other rhabditid nematodes (such as C. briggsae, C. remanei and O. tipulae), diverse animals (arthropods in particular) and many microorganisms (such as bacteria, fungi). For instance, since the food sources are transient, intraspecific competition for food among bacterivore nematodes may be substantial [START_REF] Félix | The natural history of Caenorhabditis elegans[END_REF].

Diplogastrid nematodes, such as Pristionchus species, may be predators of C. elegans and other bacterivore nematodes [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF][START_REF] Barrière | Isolation of C. elegans and related nematodes[END_REF]. Some mollusks and arthropods may serve as dispersal vectors for C. elegans and related nematodes, while other arthropods such as mites may act as vector as well as predator [START_REF] Félix | The natural history of Caenorhabditis elegans[END_REF][START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]Cutter, 2015;Frézal and Félix, 2015). The most notable interactions that C. elegans has are with microorganisms. The various microbes may act as food, commensal microorganisms, pathogens or predators, to C. elegans and other nematodes (Petersen et al., 2015). In the following parts, I will describe the interactions of C. elegans with the nonpathogenic and pathogenic microbes.

II.3.1. The interaction of C. elegans with non-pathogenic microbes

In the laboratory, N2 is usually fed with E. coli strain OP50. N2 may not carry any live microbes in its gut due to E. coli being disrupted by the animal's grinder. In the gut of wild C. elegans, however, a live bacterial flora could often be observed [START_REF] Félix | The natural history of Caenorhabditis elegans[END_REF][START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]. This is not a surprise, as wild C. elegans live in the environments that are full of decaying organic material, which are food resources for microbes. However, this also means that some microbes may be able to pass the grinder of C. elegans. Using high-power light microscope, assemblages of bacteria and fungi can be seen colonizing the rotting fruits and stems [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]. Wild C. elegans may encounter a great variety of microbes through its whole life history.

A very recent study revealed that the most abundant bacteria in the natural environment of C. elegans belong to phyla Proteobacteria, Bacteroidetes, Firmicutes and Actinobacteria (Samuel et al., 2016). Several studies also characterized the microbial community that is directly associated with C. elegans, which was dominated by Proteobacteria such as unclassified Enterobacteriaceae and members of Pseudomonas, Stenotrophomonas, Ochrobactrum, Sphingomonas and Xanthomonadaceae [START_REF] Montalvo-Katz | Association with soil bacteria enhances p38-dependent infection resistance in Caenorhabditis elegans[END_REF][START_REF] Berg | Assembly of the Caenorhabditis elegans gut microbiota from diverse soil microbial environments[END_REF]Dirksen et al., 2016).

C. elegans displays different responses to different microbial communities: C. elegans populations proliferate better in an alpha-Proteobacteria rich environment, but generate more dauers in a detrimental Bacteroidetes rich community (Samuel et al., 2016). Some associated microbes may also be able to increase the defensive ability of C. elegans in the presence of pathogens. For example, Bacillus megaterium and Pseudomonas mendocina can enhance the resistance of C. elegans to the pathogen Pseudomonas aeruginosa [START_REF] Montalvo-Katz | Association with soil bacteria enhances p38-dependent infection resistance in Caenorhabditis elegans[END_REF]Dirksen et al., 2016).

More importantly, C. elegans uses some bacteria and yeast in its natural habitat as food resources [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]Frézal and Félix, 2015). The exact food source of bacteria or yeast species of C. elegans in the wild is unclear, but different diets can affect C. elegans life history traits such as longevity and growth rate [START_REF] Macneil | Food, pathogen, signal: the multifaceted nature of a bacterial diet[END_REF]. Some microbes may decrease the growth rate or longevity of C. elegans, while other microbes may show directly pathogenic effects. It may be difficult to disentangle pathogenic effects from bad dietary effects, but it is clear that overlaps exist between dietary, commensal and pathogenic microbes [START_REF] Macneil | Food, pathogen, signal: the multifaceted nature of a bacterial diet[END_REF][START_REF] Clark | Commensals, probiotics and pathogens in the Caenorhabditis elegans model[END_REF]. The non-pathogenic microbes colonizing the gut may improve the resistance of C. elegans to its true pathogens.

II.3.2. The interaction of C. elegans with its pathogens

Diverse pathogens are able to infect C. elegans. In the wild environment, several microbes have been characterized as natural pathogens to C. elegans and other nematodes (Figure 15). These pathogens infect C. elegans mainly via two different routes: the external surfaces or the oral route. For instance, the fungus Drechmeria coniosporia and the bacterium Leucobacter species adhere to the cuticle [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]Hodgkin et al., 2013;Frézal and Félix, 2015). The fungus Harposporium sp., the bacteria Serratia marcescens and Microbacterium nematophilum, the Orsay virus and the microsporidia Nematocida parisii infect C. elegans via the oral route [START_REF] Hodgkin | A novel bacterial pathogen, Microbacterium nematophilum, induces morphological change in the nematode C. elegans[END_REF]Troemel et al., 2008;[START_REF] Félix | Natural and experimental infection of Caenorhabditis nematodes by novel viruses related to Nodaviruses[END_REF][START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF][START_REF] Parsons | Oral ingestion of Microbacterium nematophilum leads to anal-region infection in Caenorhabditis elegans[END_REF]. The latter two pathogens further induce intracellular infections in intestinal cells. In addition to its natural pathogens, C. elegans has been found susceptible to a large number of "universal" pathogens, including some human pathogens [START_REF] Kurz | Caenorhabditis elegans for the study of host-pathogen interactions[END_REF][START_REF] Sifri | The worm has turned -microbial virulence modeled in Caenorhabditis elegans[END_REF] C. elegans with its pathogens, especially its natural pathogens in contact with which its immune system evolved, makes a powerful model for host-pathogen studies. In the following part of this chapter, I will first present briefly the different defensive responses of C. elegans. Then I will describe in more detail the interactions of C. elegans with its natural pathogens: the newly found Leucobacter species and the newly discovered intracellular viral infections in Caenorhabditis. The studies on the interaction between microsporidia and C. elegans will be presented in detail in Chapter III.

II.3.2.1. Defense of C. elegans against pathogens

To defend itself, C. elegans mainly uses three strategies: the avoidance behavior response, physical barriers and the innate immunity (Schulenburg et al., 2004;[START_REF] Engelmann | Innate immunity in C. elegans[END_REF].

A. Behavioral avoidance

C. elegans is a bacterivore with a foraging behavior. In a natural microbe-rich habitat, it is vital for C. elegans to recognize non-pathogenic food from harmful pathogens. C. elegans has 60 ciliated sensory neurons, functioning in chemosensation and mechanosensation, which may enable the recognition of C. elegans to different microbes [START_REF] Allen | Sensory systems: their impact on C. elegans survival[END_REF]. For instance, while some S. marcescens attract C. elegans, the cyclic pentapeptide biosurfactant serrawettin W2 expressed by S. marcescens strain Db10 leads to an avoidance behavior of C. elegans [START_REF] Pujol | A reverse genetic analysis of components of the Toll signaling pathway in Caenorhabditis elegans[END_REF][START_REF] Pradel | Detection and avoidance of a natural product from the pathogenic bacterium Serratia marcescens by Caenorhabditis elegans[END_REF]. This innate avoidance of C. elegans was also observed in the presence of M. nematophilum [START_REF] Yook | Mos1 mutagenesis reveals a diversity of mechanisms affecting response of Caenorhabditis elegans to the bacterial pathogen Microbacterium nematophilum[END_REF]. Despite these innate repulsive responses, C. elegans also showed aversive learning avoidance to P. aeruginosa and S. marcescens. Though initially C. elegans was attracted to the odors produced by P. aeruginosa and S. marcescens, it could modify its olfactory preference and learn to avoid the pathogenic lawns [START_REF] Zhang | Pathogenic bacteria induce aversive olfactory learning in Caenorhabditis elegans[END_REF][START_REF] Beale | Caenorhabditis elegans senses bacterial autoinducers[END_REF]. The avoidance behavior of C. elegans to P. aeruginosa was also reported to be affected by the mechanosensation of C. elegans [START_REF] Chang | Natural polymorphisms in C. elegans HECW-1 E3 ligase affect pathogen avoidance behaviour[END_REF]). The sensory system mediates both the food seeking and pathogen avoidance behaviors [START_REF] Meisel | Behavioral avoidance of pathogenic bacteria by Caenorhabditis elegans[END_REF].

B. Physical barriers

C. elegans has a strong collagenous cuticle, which protects itself from environmental threats, especially from the invasion of some pathogens. In addition, the pharyngeal grinder could destroy live pathogens before infection occurs. However, some pathogens could break the physical barrier of C. elegans. For instance, spores of the fungal pathogen D. coniospora mainly adhere to the cuticle in the head or around the vulva of C. elegans, and then penetrate the cuticle. S. marcescens can disrupt the grinder, enter and colonize the intestine of C. elegans; spores of microsporidia and some other pathogens resist to the pharyngeal grinding [START_REF] Darby | Interactions with microbial pathogens[END_REF][START_REF] Engelmann | Innate immunity in C. elegans[END_REF][START_REF] Taffoni | Mechanisms of innate immunity in C. elegans epidermis[END_REF].

C. Innate immunity

C. elegans monitors physiological perturbations such as epidermal injury, DNA damage, inhibition of transcription or translation, mitochondrial dysfunction. The surveillance on homeostasis could act as a trigger for the initiation of immune defense [START_REF] Taffoni | Mechanisms of innate immunity in C. elegans epidermis[END_REF][START_REF] Ewbank | Local and long-range activation of innate immunity by infection and damage in C. elegans[END_REF][START_REF] Pukkila-Worley | Surveillance immunity: an emerging paradigm of innate defense activation in Caenorhabditis elegans[END_REF].

Several signaling pathways have been shown to be involved in innate immunity in C. elegans, such as the mitogen-activated protein kinase (MAPK) pathways, the DAF-2/DAF-16 insulin-like signaling pathway, the transforming growth factor ß (TGF-ß) pathway. [START_REF] Ewbank | Signaling in the immune response[END_REF][START_REF] Engelmann | Innate immunity in C. elegans[END_REF][START_REF] Kim | Signaling in the innate immune response[END_REF]. The diverse signaling pathways control the expression of many immune effectors, including the antimicrobial peptides (AMP), the caenacins (CNC), lysozymes, lectins. [START_REF] Engelmann | Innate immunity in C. elegans[END_REF][START_REF] Kim | Signaling in the innate immune response[END_REF].

II.3.2.2. Leucobacter species

Leucobacter are Gram-positive, non-motile bacteria that have been isolated from diverse environments, including chromium-rich soil and water, foodstuffs, nematodes [START_REF] Morais | Leucobacter luti sp. nov., and Leucobacter alluvii sp. nov., two new species of the genus Leucobacter isolated under chromium stress[END_REF][START_REF] Muir | Leucobacter chromiireducens subsp. solipictus subsp. nov., a pigmented bacterium isolated from the nematode Caenorhabditis elegans, and emended description of L. chromiireducens[END_REF][START_REF] Shin | Leucobacter celer sp. nov., isolated from Korean fermented seafood[END_REF][START_REF] Ge | Distinct and effective biotransformation of hexavalent chromium by a novel isolate under aerobic growth followed by facultative anaerobic incubation[END_REF]. Recently, a new subspecies of Leucobacter celer and a new species named Leucobacter musarum with its two subspecies were isolated from wild Caenorhabditis (Hodgkin et al., 2013). The Leucobacter celer subsp. astrifaciens CBX151 and the Leucobacter musarum subsp. musarum CBX152 were isolated co-infecting a C. tropicalis strain (JU1635) from Cape Verde. This infected C. tropicalis strain showed a swollen tail response that referred to as the Dar (Deformed Anal Region) phenotype, and bacterial coating over the cuticle (Figure 16A). The Leucobacter musarum subsp. japonicus was isolated from a wild C. elegans strain (JU1088) in Kakegawa, Japan. This infected C. elegans strain exhibited the Dar phenotype (Hodgkin et al., 2013;[START_REF] Clark | Leucobacter musarum subsp. musarum sp. nov., subsp. nov., Leucobacter musarum subsp. japonicus subsp. nov., and Leucobacter celer subsp. astrifaciens subsp. nov., three nematopathogenic bacteria isolated from Caenorhabditis, with an emended description of Leucobacter celer[END_REF].

The Leucobacter species displayed diverse interactions with C. elegans. On the surface of an agar plate, the L. celer astrifaciens CBX151 infection can inhibit the movement and growth of C. elegans, but is not lethal. In liquid culture, L. celer astrifaciens CBX151 infection induces formation of "worm-stars", where C. elegans individuals aggregate by their tails, and then leads to death of most trapped animals (Figure 16B). However, within the first hour after the star formation, some trapped C. elegans individuals are able to escape the worm-stars, albeit with broken tail spikes.

These wounded C. elegans will be healed later and be able to survive. Additionally, a small proportion (5-10%) of L4 stage C. elegans individuals could escape the star even later (24 hours after star formation) through a form of autotomy. By autotomy, an anterior "half-worm" could escape and remain viable for several days, with the possibility to have a few selfing progenies (Hodgkin et al., 2013[START_REF] Hodgkin | Worm-stars and half-worms: novel dangers and novel defense[END_REF]. On the other hand, L. musarum musarum CBX152 is lethal to C. elegans on the agar plate only at higher temperature. At 25° C, L. musarum musarum CBX152 infection causes large vacuoles that distort the internal organs and distend the whole body of C. elegans, resulting in death of most animals within 24-48 h (Figure 16C). At lower temperature, L. musarum musarum CBX152 causes the Dar phenotype in C. elegans (Figure 16D).

The Dar phenotype was also observed in C. elegans infected with Microbacterium nematophilum, a Gram-positive bacterium that specifically infects C. elegans and some other nematodes [START_REF] Hodgkin | A novel bacterial pathogen, Microbacterium nematophilum, induces morphological change in the nematode C. elegans[END_REF]. Previously, through chemical and transposon mutagenesis, several C. elegans mutants that are resistant to M. nematophilum were selected [START_REF] Gravato-Nobre | Multiple genes affect sensitivity of Caenorhabditis elegans to the bacterial pathogen Microbacterium nematophilum[END_REF][START_REF] Yook | Mos1 mutagenesis reveals a diversity of mechanisms affecting response of Caenorhabditis elegans to the bacterial pathogen Microbacterium nematophilum[END_REF]. Most of these mutants were also found to resist to L. musarum musarum CBX152 (Hodgkin et al., 2013). Mutants such as bus-2, bus-4 were shown to have an altered surface glycosylation, which may affect the adhesion of L. musarum musarum CBX152 to the cuticle of C. elegans [START_REF] Gravato-Nobre | Glycosylation genes expressed in seam cells determine complex surface properties and bacterial adhesion to the cuticle of Caenorhabditis elegans[END_REF]. However, on the agar surface, L. celer astrifaciens CBX151 can efficiently kill the C. elegans mutants that resist to L. musarum musarum CBX152. Furthermore, C. elegans isolates primarily infected with L. celer astrifaciens CBX151 seemed more resistant to L. musarum musarum CBX152 infection. Altogether, these observations revealed the complementary virulence between the two Leucobacter pathogens, which may generate opposing selective forces on C. elegans (Hodgkin et al., 2013[START_REF] Hodgkin | Worm-stars and half-worms: novel dangers and novel defense[END_REF].

II.3.2.3. Orsay virus

In 2011, the Orsay virus and the Santeuil virus were identified as the first known viruses that naturally infect C. elegans and C. briggsae, respectively [START_REF] Félix | Natural and experimental infection of Caenorhabditis nematodes by novel viruses related to Nodaviruses[END_REF]. One year later, a third Caenorhabditis virus, Le Blanc virus, was characterized, also infecting C. briggsae, like the Santeuil virus [START_REF] Franz | Complete genome sequence of Le Blanc virus, a third Caenorhabditis nematode-infecting virus[END_REF]. All of these three novel viruses are positive strand RNA viruses that are distantly but most closely related to Nodaviruses [START_REF] Félix | Natural and experimental infection of Caenorhabditis nematodes by novel viruses related to Nodaviruses[END_REF][START_REF] Franz | Complete genome sequence of Le Blanc virus, a third Caenorhabditis nematode-infecting virus[END_REF][START_REF] Jiang | Orsay virus utilizes ribosomal frameshifting to express a novel protein that is incorporated into virions[END_REF].

Fluorescent in situ hybridization (FISH) and immunofluorescence assays (IFA) both determined that all the three viruses have the worm's intestinal cells as their primary tropism [START_REF] Félix | Natural and experimental infection of Caenorhabditis nematodes by novel viruses related to Nodaviruses[END_REF][START_REF] Franz | Orsay, Santeuil and Le Blanc viruses primarily infect intestinal cells in Caenorhabditis nematodes[END_REF]. C. elegans and C. briggsae with these viral infections display abnormal morphologies in their intestinal cells: the intestinal granules and nuclei may disappear; the intestinal cytoplasm may become fluid; intestinal cells may fuse (Figure 17). In the original study on the Orsay virus and C. elegans, comparisons between sequenced small RNA libraries of C. elegans populations with or without the infection of Orsay virus suggested that C. elegans produces small RNAs in response to the viral infection. The antiviral role of RNA interference (RNAi) in the immunity of C. elegans was further revealed: compared to N2, the RNAi-deficient mutants showed increased levels of viral RNA abundance [START_REF] Félix | Natural and experimental infection of Caenorhabditis nematodes by novel viruses related to Nodaviruses[END_REF]. A more recent study determined the roles of the gene drh-1 in the recognition of the RNA virus and in the sensitivity variation of C. elegans to the viral infection [START_REF] Ashe | A deletion polymorphism in the Caenorhabditis elegans RIG-I homolog disables viral RNA dicing and antiviral immunity[END_REF]. The drh-1 gene encodes a RIG-I-like helicase, which is a homologue of the mammalian viral recognition protein RIG-I. Using a genome-wide association mapping in 97 wild C. elegans isolates of the viral loads upon Orsay virus infection, a 159 base-pair deletion in the drh-1 gene of C. elegans JU1580 was identified as a major determinant of the viral sensitivity in C. elegans. This deletion was further found to impair the fitness of JU1580 and drh-1 mutants in the presence of viral infection: upon the infection of Orsay virus, JU1580 and drh-1 mutants displayed delayed and decreased total progeny compared to uninfected isolates. However, the detrimental deletion of drh-1 was found in many wild isolates of C. elegans, which may result from the low prevalence of Orsay virus in the wild, or a linkage between the drh-1 deletion and a beneficial mutation, or a conditional deleterious effect of carrying the intact drh-1 gene. Further experiments with double mutants determined that DRH-1 is an upstream factor of the antiviral siRNA pathway. DRH-1 might use a similar mechanism to RIG-I to recognize the RNA of the Orsay virus [START_REF] Ashe | A deletion polymorphism in the Caenorhabditis elegans RIG-I homolog disables viral RNA dicing and antiviral immunity[END_REF].

III. Microsporidia infections in C. elegans and related nematodes

Historically, nematode-infecting microsporidia were primarily reported in the parasitic nematodes. Protospirura muris, a parasitic nematode of mouse, was reported to carry an infection by Thelohania reniformis [START_REF] Kudo | Notes on a Microsporidian parasite of a nematode[END_REF].

Nosema mesnili and Plistophora schubergi are able to infect an entomopathogenic nematode, Neoaplectana carpocapsae, as well as insect hosts (Veremtchuk and Issi, 1970;Poinar and Hess, 1988;Bakowski et al., 2014a). The first restricted nematodeinfecting microsporidia was called "Microsporidium" rhabdophilum (undefined genus), which was isolated from a nonparasitic nematode Oscheius (Rhabditis) myriophila [START_REF] Poinar | Microsporidium rhabdophilum sp. n. (Microsporida: Pansporoblastina), a parasite of the nematode, Rhabditis myriophila (Rhabditina: Rhabditidae)[END_REF]Bakowski et al., 2014a).

In 2008, three wild-caught C. elegans strains from France and one wild C. briggsae strain from India were reported to carry microsporidia infections (Troemel et al., 2008). A new microsporidian genus, Nematocida, was established with two species:

Nematocida parisii and the undescribed Nematocida sp. 1. The genus Nematocida was placed in the Clade II of the microsporidia phylum phylogeny (Figure 4). In the phylogenetic analysis, the closest genus to Nematocida is Ovavesicula, with the type species Ovavesicula popilliae (Figure 4) [START_REF] Vossbrinck | The phylogenetic position of Ovavesicula popilliae (Microsporidia) and its relationship to Antonospora and Paranosema based on small subunit rDNA analysis[END_REF]Troemel et al., 2008). O. popilliae infects the Japanese beetle, Popillia japonica, and forms oval-shaped spores [START_REF] Andreadis | Ultrastructural Study and Description of Ovavesicula popilliae n. g., n. sp. (Microsporida: Pleistophoridae) from the Japanese beetle, Popillia japonica (Coleoptera: Scarabaeidae)[END_REF]. Nematocida, on the other hand, literally means nematode killer (Troemel et al., 2008). Both N. parisii and N. sp. 1 are transmitted horizontally and infect the intestinal cells of C. elegans or C. briggsae, with rod-shaped spores of two distinct sizes (Troemel et al., 2008). Further sampling revealed that microsporidian infections are quite prevalent among wild Caenorhabditis and related nematodes [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]. These Nematocida pathogens are lethal threats to C. elegans individuals and other related nematode species.

In addition to Caenorhabditis, a free-living marine nematode, Odontophora rectangular was found infected with a novel microsporidia species, named Sporanauta perivermis [START_REF] Ardila-Garcia | Microsporidian infection in a free-living marine nematode[END_REF]. Later, a deep sea nematode Desmodora marci was found infected with a microsporidium, which was named Nematocenator marisprofundi (Sapir et al., 2014). Neither of these two nematodeinfecting microsporidia are closely related to Nematocida species. S. perivermis was placed in the Clade IV of the microsporidia, while N. marisprofundi was in none of the described clade, but in the basal position of the microsporidia phylum. S. perivermis infects host's muscle and hypodermis, and a vertical transmission mode was suggested [START_REF] Ardila-Garcia | Microsporidian infection in a free-living marine nematode[END_REF]. N. marisprofundi was shown to cause infection in the host's body wall muscles. N. marisprofundi does not seem to be transmitted through the fecal-oral route or the vertical transmission mode, as their spores have never been observed in the nematode intestines or nematode eggs. However, N. marisprofundi spores were observed in the reproductive organs of D. marci males and females, which suggested that the transmission is through the mating behavior (Sapir et al., 2014).

In the following parts, I will further present the interactions of C. elegans with Nematocida species.

III.1. Nematocida parisii

The very original N. parisii strain ERTm1 was isolated from the wild C. elegans strain CPA24 near Paris, France (Troemel et al., 2008). Wild C. elegans strains JU1247 and JU1395, C. briggsae strain JU2055 were also naturally infected with N. parisii (N. parisii strains named ERTm3, ERTm4, ERTm5 respectively) (Troemel et al., 2008;Balla et al., 2015). In both host species, N. parisii is transmitted horizontally through the fecal-oral route, with meronts and two distinct sizes of spores appearing in the host intestinal cells (Figure 18). The small spore is 2.18 ± 0.15 µm long and 0.8 ± 0.08 µm wide, with one observed polar tube coil; while the large spore is 3.17 ± 0.22 µm long and 1.31 ± 0.15 µm wide, with up to five visible polar tube coils (Figure 19 F,G). Transmission electron microscopy (TEM) suggests that N. parisii is monokaryotic, as only one nucleus or multiple unpaired nuclei have been observed in its meront. Molecular identification and phylogenetic analysis using small subunit rDNA characterized N. parisii and placed it in the Clade II of the microsporidia phylum (Figure 4) (Troemel et al., 2008). A. An irregular shaped multinucleate meront indicated with an arrow. A host vesicle is marked with an asterisk. Scale bar is 1µm. B. A later stage meront with two nuclei and more regularly shaped plasma membrane. C. and D. are sporonts with electrondense membranes. E. Developing N. parisii spores. F. A small sized spore. G. A larger sized spore. Arrowheads indicate cross-sections of polar tube coils in F and G. H. Vesicles of spores. B-H, scale bar is 500 nm. PP refers to the polaroplast membrane; PV refers to the posterior vacuole. Adapted from Troemel et al. (2008).

III.1.1. Life cycles of N. parisii and symptoms in the host

Ingestion of N. parisii spores by C. elegans begins the infection (Figure 20). N. parisii spores can survive the pharyngeal grinding and reach the intestinal lumen of C. elegans. Though the injecting process has not been observed directly yet, N. parisii spore, like other microsporidia species, probably uses the polar tube to inject its sporoplasm into the host intestinal cells.

Before becoming newly mature spores, two different developmental stages of N. parisii could be observed under transmission electron microscopy: the meront stage and the sporont stage. At 25 °C, within the first day after inoculation, the injected N. parisii sporoplasm replicates and develops into a meront, which causes displacement of gut granules and "grooves" in the C. elegans intestinal cells (Figure 18B). Early stage meronts are irregularly shaped and later stage meronts are more regularly shaped. Both may contain one or several nuclei (Figure 19A,B). The multinucleate meronts of N. parisii may divide by plasmotomy to cells containing one nucleus (Figure 20). Meronts develop into sporonts, which possess electron-dense membranes (Figure 19C,D). Multinucleate sporonts may also form and eventually divide by plasmotomy. Small spores appear around 40 hours after inoculation (Figure 19E,F). Large spores are observed later than small spores (Figure 18C,19G); discrete vesicles filled with spores appear at the end (Figures 18D,19H) (Troemel et al., 2008).

Newly formed mature spores of N. parisii exit from the host cell into the intestinal lumen soon after their formation. The spores will be released to the outside environment presumably through the defecation process of C. elegans (Figure 20) (Troemel et al., 2008;Estes et al., 2011).

Figure 20. Diagram of life cycle of N. parisii infection in C. elegans.

N. parisii infects C. elegans intestinal cells (yellow), where it grows intracellularly through the meront and sporont stages. Newly formed spores exit apically into the lumen and then are defecated by the animal. The actin-rich microvilli are anchored into a terminal web (actin is labeled orange). Adapted from Szumowski et al. (2014).

N. parisii proliferates rapidly in C. elegans, with a doubling time about 2.9h to 3.3h during the meront stage (Cuomo et al., 2012). A single infected C. elegans can shed thousands of spores per hour to the environment (Estes et al., 2011). N. parisii can be rapidly transmitted across an entire C. elegans population. Infected C. elegans individuals with only small sized spores can infect other individuals, forming spores of both sizes in recipient C. elegans (Troemel et al., 2008). Infected C. elegans with only a small number of spores inside its intestinal cells is able to infect other uninfected C. elegans, whereas infected C. elegans with only meronts is not infectious (Troemel et al., 2008).

On the host side, compared to healthy C. elegans, infected C. elegans has a paler color appearance, a thinner and smaller body size. N. parisii infection could also lead to a high rate of early death, but infected C. elegans could show grossly normal behaviors even when their intestinal cells are fully filled with N. parisii spores, reflecting the tolerance of C. elegans to N. parisii infection (Troemel et al., 2008). A recent study showed that in the presence of very high doses of N. parisii spores, C. elegans N2 in the L1 stage may enter a larval developmental arrest stage [START_REF] Luallen | Characterization of microsporidiainduced developmental arrest and a transmembrane leucine-rich repeat protein in Caenorhabditis elegans[END_REF].

III.1.2. Exit mechanism from C. elegans intestinal cells

N. parisii employs a specific non-lytic mechanism to exit from C. elegans intestinal cells to the lumenal space, while minimizing damages to the host (Estes et al., 2011;[START_REF] Troemel | Host-microsporidia interactions in Caenorhabditis elegans, a model nematode host[END_REF]. The N. parisii spore exit strategy can be divided into two successive phases: 1) N. parisii infections restructure the terminal web of the host intestinal cell, by first altering the localization of apically-restricted actin. ACT-5, a specialized actin isoform that is localized to the microvilli and the terminal web in uninfected C. elegans, is localized on the basolateral side of intestinal cells of infected C. elegans (Figure 20,21A), while the levels of the apical actin often appeared to decrease. The rearrangement of the actin may serve as a trigger to induce IFB-2, a component of the intermediate filament in the terminal web, to reorganize and cause gaps in the terminal web (Figure 21) (Troemel et al., 2008;Estes et al., 2011). The actin rearrangement and the formation of terminal web gaps were determined to be a 60 discrete, highly regulated and necessary event, likely facilitating spore exiting by removing a barrier (Estes et al., 2011). 2) N. parisii spores exit from C. elegans intestinal cells through a RAB-11-mediated directional exocytosis mechanism: in the meront stage, N. parisii is in direct contact with the host intestinal cytoplasm, but transmission electron microscopy showed that a newly formed N. parisii spore is surrounded by an additional membrane (AM), defining a spore-containing compartment (SCC) (Figure 22B) (Szumowski et al., 2014). The localization of the host small GTPase protein RAB-11 to this compartment is required for SCCs to traffic to the apical side of the host cell and then fuse to the host membrane (Figure 22A). Other host GTPase proteins such as RAB-5 and RAB-10 are also important to the fusion process (Szumowski et al., 2014). After the SCCs fuse with the apical membrane (Figure 22C,D), spores gain access to the intestinal lumen, and acquire an apical membrane transporter protein PGP-1 marker (Szumowski et al., 2014). Then the spores are able to exit from the host cell into the lumen (Figure 22A). The newly shed spores in the lumen do not possess the additional membrane (Estes et al., 2011). In summary, N. parisii reorganizes the C. elegans intestinal cytoskeleton, then fuses to the host apical membrane in a GTPase-dependent mode, and finally exits host cells without causing cell lysis or any other lethal damages to the host.

III.1.3. Defensive response of C. elegans to N. parisii

All postembryonic stages of C. elegans are susceptible to N. parisii infection, except for the non-feeding dauer stage, at which the animals still can be a carrier of N. parisii (Troemel et al., 2008). However, neither the PMK-1 p38 MAPK immune pathway nor the DAF-2/DAF-16 insulin-like immune pathway of C. elegans showed any substantial roles in defense against N. parisii infection. In C. elegans, many genes upregulated upon P. aeruginosa and D. coniospora infections do not present clear expression changes when C. elegans is infected with N. parisii (Troemel et al., 2008).

RNA-seq analysis showed that the transcriptional response of C. elegans to N. parisii infection is similar to that upon intracellular infection by the Orsay virus, but very different from some extracellular infections such as P. aeruginosa and D. coniospora (Bakowski et al., 2014b). Taken together, C. elegans may have a distinct defensive response to N. parisii infection compared to its immune responses to other fungal or bacterial pathogens.

C. elegans appears to be able to monitor the functioning and homeostasis of core cellular processes, and then trigger the expression of defensive genes upon any perturbation or disruption of these processes. Core process surveillance pathways could help C. elegans to recognize pathogen invasion. For instance, the C. elegans ubiquitin-proteasome system (UPS) can trigger defensive responses, such as the ubiquitin-mediated responses, upon N. parisii and Orsay virus infections.

Upregulation of Skp1-Cull-F-box protein (SCF) ligase components could help to restrict microsporidia growth. However, this ubiquitin-mediated defense is only able to target a small percentage of N. parisii cells in their early developmental stage, which may partially be due to the fact that N. parisii is able to suppress the host UPS defensive machinery. The proteasome and autophagy pathways also promote host defense against N. parisii (Bakowski et al., 2014b).

A recent study showed that natural variation exists among six C. elegans wild isolates in their sensitivity upon N. parisii infection, as the different mortality rates among these strains are positively correlated with the pathogen loads (Figure 23A,B). This result indicates variation in host resistance rather than tolerance (Balla et al., 2015).

Fecundity was further compared between N2 and CB4856. The fecundity of infected C. elegans seemed to depend on the developmental stage of the C. elegans individuals when they were inoculated. Indeed, when the inoculation of N. parisii occurred at the L4 stage, neither N2 nor CB4856 showed significant fecundity reduction, compared to uninfected individuals. However, when the inoculation was initiated at the L1 stage, the fecundity of N2 could drop drastically compared to CB4856. The L1 stage animals of CB4856 showed an enhanced defensive ability to control and clear N. parisii infection. This clearing ability of CB4856 L1 larvae helps to maintain a relatively normal brood size and gain a reduced mortality rate (Balla et al., 2015) (Figure 23C,D). Though this increased immune ability of the strain CB4856 is restricted to the L1 stage, it provides a selective advantage over N2 (Balla et al., 2015). The npr-1 gene, though known to be responsible for several phenotypic variations in growth, physiology and response to some other pathogens between CB4856 and N2, does not appear to have a role here for the L1-stage specific enhanced resistance in CB4856 [START_REF] Andersen | A variant in the neuropeptide receptor npr-1 is a major determinant of Caenorhabditis elegans growth and physiology[END_REF]Balla et al., 2015;[START_REF] Nakad | Contrasting invertebrate immune defense behaviors caused by a single gene, the Caenorhabditis elegans neuropeptide receptor gene npr-1[END_REF]. Again, this indicated that C. elegans may employ a distinct immune response to defend itself against N. parisii.

III.2. Nematocida sp. 1

Nematocida sp. 1 was reported in the initial article on N. parisii (Troemel et al., 2008). Its SSU rDNA sequence is approximately 95% identical to the same region of N. parisii. N. sp. 1 and N. parisii were placed together as sister species in the phylogeny of microsporidia (Troemel et al., 2008). The first N. sp. 1 strain (ERTm2) was found from a wild C. briggsae strain JU1348 collected in Kerala, India (Troemel et al., 2008). The second N. sp. 1 strain ERTm6 was isolated from another wild-caught C. briggsae strain JU1638 in Cape Verde Islands. Though both N. sp. 1 strains were found in C. briggsae, N. sp. 1 is also capable of infecting C. elegans (Bakowski et al., 2014c). N. sp. 1 has a similar life cycle and developmental stages to N. parisii (Troemel et al., 2008).

III.3. Nematocida genomics

Similar to other microsporidia, genomes of Nematocida species are quite small and compact.

The N. parisii strains ERTm1 and ERTm3 share 99.8% identical genomes, that have 4.1 Mb in size. The N. sp. 1 strain ERTm2 has a larger genome of 4.6 Mb, with only 68.3% average identity to the N. parisii ERTm1 genome (Cuomo et al., 2012). The other N. sp.1 strain ERTm6, however, possesses a genome of about 4.28 Mb (Bakowski et al., 2014c). The genomes of both species are compact, with 72.8% of ERTm1 and 67.48% of ERTm6 genome sequences predicted to be coding, and a mean distance between coding sequences of 418 and 579 bp, respectively (Cuomo et al., 2012;Bakowski et al., 2014c). In the N. parisii ERTm1 genome, no introns and no untranslated regions (UTRs) were predicted in protein-coding genes, or unusually short UTRs (Cuomo et al., 2012).

Genome sequence analyses further indicated that Nematocida may have experienced a rare or recent recombination event: all four Nematocida strains of N. parisii and N. sp. 1 were found likely diploid and heterozygous, but in the N. sp. 1 ERTm2 genome, loss of heterozygosity (LOH) regions were also found (Cuomo et al., 2012;Bakowski et al., 2014c;[START_REF] Troemel | Host-microsporidia interactions in Caenorhabditis elegans, a model nematode host[END_REF].

N. sp. 1 has a higher rate of heterozygosity and polymorphism than N. parisii. N. sp. 1

ERTm2 has an average of one single nucleotide polymorphism (SNP) every 82 bases.

However, a single large homozygous region in each of the scaffolds 1, 2 and 4 was found in the genome of ERTm2, which suggested that LOH has occurred in the three scaffolds. Further experiments showed that the LOH event did not occur in ERTm2 in the lab, as the ERTm2 isolate serially passaged for 14 weeks showed a nearly identical pattern of polymorphism to the original isolate. These findings together suggested that Nematocida may have a rare sexual cycle or mitotic recombination events during its life history in the wild.

Phylogenomic analysis with fungi showed that all microsporidia shared 882 highly conserved core genes, 137 of which were identified to be microsporidia-specific genes (Cuomo et al., 2012). Nematocida are found to be quite divergent from other microsporidian species, with a very large Nematocida-specific gene family in both N. parisii (ERTm1 and ERTm3) and N.

sp. 1 (ERTm2). In the genome of N. parisii ERTm1, a total of 2,661 genes were predicted, while the predicted number is 2,443 for N. sp. 1 ERTm6 (Cuomo et al., 2012;Bakowski et al., 2014c). Pfam protein domain analysis showed that, similar to other microsporidia, the Nematocida genome encodes the nucleoside phosphate transporter (Npt) proteins and the nucleoside H + symporter, but lost the retinoblastoma (RB) tumor suppressor gene. Two Npt transporters were suggested to be encoded in the Nematocida genome. The Npt transporters may help import host nucleotides into the pathogen. Nematocida hexokinase genes were also found to have a predicted secretion signal to the host cell cytoplasm, which may boost host synthesis of building blocks that later promote the rapid growth of Nematocida (Cuomo et al., 2012).

IV. The background of my PhD thesis

Hosts and parasites are tightly coupled during their life time and may coevolve antagonistically. Hosts evolve better remote detection and avoidance ability, while parasites evolve better seeking skills; hosts evolve resistance to control or clear infection, while parasites evolve higher virulence to proliferate on/in the host. These reciprocal interactions could exert strong selection pressures on both partners and thus drive antagonistic coevolution. Host-parasite coevolution contributes to the diversity in both hosts and parasites [START_REF] Schulte | Host-parasite coevolution favours parasite genetic diversity and horizontal gene transfer[END_REF][START_REF] Bever | Maintenance of plant species diversity by pathogens[END_REF]. Meanwhile, host-parasite co-evolution and adaptation may also lead to specific interactions between hosts and parasites [START_REF] Barrett | Continua of specificity and virulence in plant host-pathogen interactions: causes and consequences[END_REF]Antonovics et al. 2013). Patterns of specificity may be found at either the interspecific or the intraspecific level for either the host or the parasite. To study the specificity and the molecular mechanisms underlying host-parasite interaction and coevolution is crucial to understand the defense and infection mechanisms themselves as well as their evolutionary dynamics.

C. elegans and its natural intracellular parasite Nematocida parisii provide an excellent model system to study host-parasite interactions and coevolution (Troemel et al., 2008;[START_REF] Hodgkin | Caenorhabditis elegans meets microsporidia: the nematode killers from Paris[END_REF][START_REF] Balla | Caenorhabditis elegans as a model for intracellular pathogen infection[END_REF]. Studies with C. elegans and N. parisii have already made many progresses. However, it is still unclear how widespread and diverse microsporidia infections are in C. elegans or other related nematodes in the wild. The discovery of the rotting substrates as the natural habitats of C. elegans and other related nematodes provided the opportunities to address more ecological questions in the wild nematode populations [START_REF] Félix | The natural history of Caenorhabditis elegans[END_REF]Kiontke et al., 2011). Among the samples of wild nematode isolates, the prevalence of microsporidia infection was further observed [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]. When I began my PhD studies in October 2013 in the Felix Lab, there were dozens of wild nematode strains, including several species, that displayed the putative microsporidia infection. Furthermore, natural variation in the sensitivity of different wild C. elegans strains to the microsporidian infection was also observed preliminarily.

During the process of my PhD project, I first characterized these microsporidia by their molecular and morphological features. Then, at the interspecific level, I studied the hostparasite specificity between different nematodes and microsporidia; at the intraspecific level, I investigated the variation in sensitivity of different wild C. elegans to the same microsporidia infection, with the further goal of determining the loci involved in C. elegans host sensitivity variation. Overall, my PhD project focused on the host-pathogen interactions of C. elegans and other nematodes with microsporidia.

70

RESULTS

I. A large collection of novel nematode-infecting microsporidia and their diverse interactions with Caenorhabditis elegans and other related nematodes

In the Felix Lab collection of wild nematodes, 47 strains of ten species, mainly including Caenorhabditis and Oscheius species, were noted to display putative microsporidia infections.

In preliminary observations using Nomarski microscopy, spores and meronts were all found in the intestinal cells of these putative infected nematode strains, except for one C. brenneri strain where spores were mainly located in the epidermis. We studied these natural infections in the wild nematode isolates. Here, we describe our discoveries of novel microsporidia species and genera infecting wild nematode isolates. In all cases, infections were found to be horizontally transmitted. We also determined their host specificities. I summarize below the main findings, followed by the article that was recently published in PLOS Pathogens.

To characterize these putative microsporidian infections in the nematodes, we first amplified and sequenced two genes (SSU rDNA and β-tubulin genes). Phylogenetic trees were constructed using these sequences and published sequences from species belonging to the microsporidia phylum. In addition to Nematocida parisii and Nematocida ausubeli (formerly called N. sp. 1), we have found six new Nematocida species. Furthermore, we have discovered two new genera, named Enteropsectra and Pancytospora.

The morphology (spore sizes, spore shapes, spore distribution in the host cells) of all the new microsporidia species were examined by Nomarski microscopy. The similarities and differences in the morphological features matched their groupings according to the phylogenetic analysis. Using transmission electron microscopy (TEM), we discovered that N.

ausubeli and E. longa use different mechanisms for the exit of the spores out of their host intestinal cells.

Since each microsporidia species was isolated from a limited number of host species, we further investigated the host specificity of different microsporidia. Cross-inoculation tests were performed using eight microsporidia species and four nematode species. The host range differs depending on the microsporidia species. For example, Nematocida homosporus could infect the two Caenorhabditis species and the two Oscheius species. However, Enteropsectra longa only infected Oscheius sp. 3. The host specificity of different microsporidia further revealed the diverse interactions between microsporidia and nematodes. Inter-and intraspecific host specificity exists among different nematode-infecting microsporidia.

Among the different nematode species, C. elegans has been the best studied. 19 C. elegans wild strains were naturally infected with N. parisii or N. ausubeli. C. elegans was also found susceptible to N. major, N. homosporus and P. epiphaga in the infection tests. We further used two C. elegans strains carrying transgenic reporter genes upregulated upon N. parisii infection, to compare the host response to the infections by five microsporidia species.

Interestingly, the results suggested that C. elegans may have a different transcriptional response to the infection by N. ausubeli than to the infections by the other tested Nematocida species.

Overall, our study considerably enriched the knowledge on nematode-infecting microsporidia and their diverse interactions. We found novel microsporidia species and genera in wild nematode isolates. These nematode-infecting microsporidia display different host specificities. As C. elegans has proven to be a successful model organism, and microsporidia are obligate intracellular pathogens of C. elegans and other nematodes, different microsporidia and nematodes provide an ideal system to study host-pathogen interactions.

Our study enlarged the available resources and provided a wide variety of natural nematodemicrosporidia host-pathogen pairs. My contributions in this study: I contributed the majority of assays in this study, except the TEM part (Figure 5,8; Supplemental Figure 1,6), for which we collaborated with Martin Sachse and Marie-Christine Prevost from Pasteur Institute in Paris, and I contributed to the fixations and observations. The assays with transgenic C. elegans strains (Figure 9) were performed with Robert J.

Abstract

Microsporidia are fungi-related intracellular pathogens that may infect virtually all animals, but are poorly understood. The nematode Caenorhabditis elegans has recently become a model host for studying microsporidia through the identification of its natural microsporidian pathogen Nematocida parisii. However, it was unclear how widespread and diverse microsporidia infections are in C. elegans or other related nematodes in the wild. Here we describe the isolation and culture of 47 nematodes with microsporidian infections. N. parisii is found to be the most common microsporidia infecting C. elegans in the wild. In addition, we further describe and name six new species in the Nematocida genus. Our sampling and phylogenetic analysis further identify two subclades that are genetically distinct from Nematocida, and we name them Enteropsectra and Pancytospora. Interestingly, unlike Nematocida, these two genera belong to the main clade of microsporidia that includes human pathogens. All of these microsporidia are horizontally transmitted and most specifically infect intestinal cells, except Pancytospora epiphaga that replicates mostly in the epidermis of its Caenorhabditis host. At the subcellular level in the infected host cell, spores of the novel genus Enteropsectra show a characteristic apical distribution and exit via budding off of the plasma membrane, instead of exiting via exocytosis as spores of Nematocida. Host specificity is broad for some microsporidia, narrow for others: indeed, some microsporidia can infect Oscheius tipulae but not its sister species Oscheius sp. 3, and conversely some microsporidia found infecting Oscheius sp. 3 do not infect O. tipulae. We also show that N. ausubeli fails to strongly induce in C. elegans the transcription of genes that are induced by other Nematocida species, suggesting it has evolved mechanisms to prevent induction of this host response. Altogether, these newly isolated species illustrate the diversity and ubiquity of microsporidian infections in nematodes, and provide a rich resource to investigate host-parasite coevolution in tractable nematode hosts.

Introduction

Microsporidia are fungi-related obligate intracellular pathogens, with over 1400 described species [1,[START_REF] Szumowski | Microsporidia-host interactions[END_REF]. Interest in these organisms started 150 years ago when researchers, especially Louis Pasteur, studied silkworm disease that was caused by a microsporidian species later named Nosema bombycis [START_REF] Franzen | Microsporidia: a review of 150 years of research[END_REF]. In the past decades, microsporidia have attracted more attention when they were revealed to be a cause of diarrhea in immunocompromised patients and were further demonstrated to have a high prevalence in some areas in immunocompetent patients and healthy individuals [START_REF] Nkinin | Microsporidian infection is prevalent in healthy people in Cameroon[END_REF][START_REF] Mumcuoglu | Prevalence of microsporidia in healthy individuals and immunocompetent patients with acute and chronic diarrhea[END_REF][START_REF] Stentiford | Microsporidia-Emergent pathogens in the global food chain[END_REF].

Microsporidia are transmitted between hosts through a spore stage. Inside the microsporidian spore is found a characteristic structure called the polar tube, which at the time of infection can pierce through host cell membranes and introduce the sporoplasm (the spore cytoplasm and nucleus) into host cells [1,[START_REF] Keeling | Microsporidia: biology and evolution of highly reduced intracellular parasites[END_REF]. These obligate intracellular pathogens are known to infect a wide range of hosts among protists and animals, especially insects, fish and mammals [1]. Even though nematodes constitute a huge phylum with over 25,000 described species, very few studies on microsporidian infections in nematodes have been reported so far [1].

The free-living nematode Caenorhabditis elegans has been used as a major biological model species over the last 50 years [START_REF] Corsi | A transparent window into biology: A primer on Caenorhabditis elegans[END_REF]. However, until the past decade, little was known about its biology and ecology in its natural environment and no natural pathogens were isolated until C. elegans could be readily isolated from natural environments. C. elegans is now known to be found in compost heaps, rotting fruits (apples, figs, etc.) and herbaceous stems, as well as with diverse carrier invertebrates (snails, isopods, etc.) [START_REF] Barrière | High local genetic diversity and low outcrossing rate in Caenorhabditis elegans natural populations[END_REF][START_REF] Fe ´lix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF][START_REF] Barrière | Isolation of C. elegans and related nematodes[END_REF]. C. elegans coexists with a variety of prokaryotic and eukaryotic microbes, including both its food and pathogens, which likely have a large impact on its physiology and evolution [START_REF] Ma | C. elegans outside the Petri dish[END_REF][START_REF] Petersen | Why we need more ecology for genetic models such as C. elegans[END_REF][START_REF] Dirksen | The native microbiome of the nematode Caenorhabditis elegans: gateway to a new host-microbiome model[END_REF][START_REF] Samuel | Caenorhabditis elegans responses to bacteria from its natural habitats[END_REF].

With an improved understanding of the natural history of Caenorhabditis [START_REF] ´lix | The natural history of Caenorhabditis elegans[END_REF][START_REF] Cutter | Caenorhabditis evolution in the wild[END_REF], dramatically increased number of various wild rhabditid nematode strains and species have been isolated and identified. C. elegans' close relatives such as Caenorhabditis briggsae or Caenorhabditis remanei are isolated from similar environments [START_REF] Kiontke | A phylogeny and molecular barcodes for Caenorhabditis, with numerous new species from rotting fruits[END_REF]. Oscheius tipulae, a very common bacteriovorous nematode species, also in family Rhabditidae, can be readily isolated from soil and rotting vegetal matter [START_REF] Fe ´lix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF][START_REF]Oscheius tipulae[END_REF]20], as well as its closest known relative Oscheius sp. 3, with which it cannot interbreed [START_REF] ´lix | Three biological species related to Rhabditis (Oscheius) pseudodolichura Ko ¨rner in Osche[END_REF]. Interest in these rhabditid nematodes concerns not only the evolution of genomic and phenotypic characters, but also their inter-and intraspecific interactions and co-evolution with other organisms, especially with various microbes in their natural habitats. While nematodes feed on bacteria and small eukaryotes, some microbes take nematodes as their food source [START_REF] Petersen | Why we need more ecology for genetic models such as C. elegans[END_REF][START_REF] Dirksen | The native microbiome of the nematode Caenorhabditis elegans: gateway to a new host-microbiome model[END_REF][START_REF] ´lix | The natural history of Caenorhabditis elegans[END_REF]. Among them, microsporidia are obligate intracellular parasites and thus are in particularly tight association with their hosts.

The microsporidian Nematocida parisii was the first found natural intracellular pathogen of C. elegans, which we isolated from a wild C. elegans sampled near Paris, France [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF]. Nematocida sp. 1 (described here as Nematocida ausubeli) was further isolated from a wild Caenorhabditis briggsae strain in India [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF]. A microsporidian species isolated in C. elegans was found to infect the epidermis and muscles and was named Nematocida displodere [START_REF] Luallen | Discovery of a natural microsporidian pathogen with a broad tissue tropism in Caenorhabditis elegans[END_REF]. Two microsporidia infecting marine nematodes have also been described, namely Sporanauta perivermis [START_REF]Microsporidian infection in a free-living marine nematode[END_REF] and Nematocenator marisprofundi [START_REF] Sapir | Microsporidia-nematode associations in methane seeps reveal basal fungal parasitism in the deep sea[END_REF][START_REF] Bakowski | Microsporidia infections in Caenorhabditis elegans and other nematodes[END_REF]. However, the extent and diversity of microsporidia infections in nematodes remained sparsely described.

Here we describe a collection of 47 terrestrial nematode strains that we isolated from the wild with a microsporidian infection. The microsporidia can be grown in the laboratory in their host using C. elegans culture conditions and stored frozen with their nematode host. They are all transmitted horizontally. In this set, we found that N. parisii and N. ausubeli (formerly called N. sp. 1) are in association with further host species and display a wider geographical distribution than so far reported [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF]. N. parisii is the most common C. elegans-infecting species we found in the wild. We further discovered new nematode-infecting microsporidian species. From our phylogenetic analysis using small subunit (SSU) ribosomal DNA and βtubulin sequences, five new microsporidia species were placed in the Nematocida genus, while the others defined two new genera in the microsporidian clade often designated as Clade IV, which includes human pathogenic microsporidia such as Enterocytozoon bieneusi and Vittaforma corneae. The similarities and differences in the morphological features of these microsporidia matched their groupings by sequence similarity. We therefore describe two new microsporidian genera, Enteropsectra and Pancytospora. and nine new species in these two genera and Nematocida. We further examined Nematocida ausubeli and Enteropsectra longa by electron microscopy, which allowed us to observe different mechanisms for their exit from host intestinal cells, through a vesicular pathway for N. ausubeli (as described for N. parisii; [START_REF] Szumowski | The small GTPase RAB-11 directs polarized exocytosis of the intracellular pathogen N. parisii for fecal-oral transmission from C. elegans[END_REF]), but surprisingly through membrane budding for E. longa. Concerning specificity of infection, we find cases of tight specificity between host and pathogen. We also find that N. ausubeli fails to strongly induce the transcription of genes that are induced in C. elegans by N. parisii infection. Overall, our study points to strong and diverse interactions between wild rhabditid nematodes and microsporidia, and provides a platform for further study of these infections.

Results

A large collection of microsporidian-infected nematode cultures

Our worldwide sampling of bacteriovorous nematodes was primarily aimed at isolating Caenorhabditis species and, to a lesser degree, Oscheius species. From this sampling, we identified a subset of strains with a pale body color (Fig 1A), some of which, upon morphological examination using Nomarski microscopy [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF], appeared infected with microsporidia. In total, we collected 47 nematode strains (S1 Table) displaying putative microsporidian infections, comprising 10 nematode species from different parts of the world (Tables 1 and2; Fig 1B). The microsporidia strain JUm2807 was isolated during these sampling efforts and described elsewhere as Nematocida displodere, and is not considered here [START_REF] Luallen | Discovery of a natural microsporidian pathogen with a broad tissue tropism in Caenorhabditis elegans[END_REF].

The unidentified microsporidian strains were characterized by sequencing of PCR fragments of the SSU rDNA and β-tubulin genes. We were able to amplify 45 SSU rDNA sequences (most 1390 bp long) and 32 β-tubulin sequences (most 1210 bp long) (S1 Table ). We first blasted the sequences in GenBank for initial grouping, then built phylogenetic trees and calculated interspecific genetic distances, based on our sequences and the sequences of related species from GenBank. We present below the grouping and phylogenetic distribution of new microsporidia strains, starting with those closest to N. parisii.

N. parisii and N. ausubeli are commonly found in Caenorhabditis nematodes

Molecular sequences of microsporidia in ten wild C. elegans strains and four C. briggsae strains showed 99% SSU rDNA and 97% β-tubulin sequence identities to N. parisii sequences in [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF] d Note that a recent article describes a new species for this strain based on genome divergence [START_REF] Reinke | Troemel ER Identification of microsporidia host-exposed proteins reveals a repertoire of large paralogous gene families and rapidly evolving proteins[END_REF].

doi:10.1371/journal.ppat.1006093.t001

GenBank. In the global phylogenetic analysis of microsporidia, these 14 sequences form a group with previously reported sequences of N. parisii strains ERTm1, ERTm3 and ERTm5 [START_REF] Bakowski | Microsporidia infections in Caenorhabditis elegans and other nematodes[END_REF] (Fig 2). The N. parisii isolates were all found in Europe (note however that the sampling is highly biased towards Europe, especially France), with the exception of the previously reported ERTm5 (JUm2055), isolated from a C. briggsae strain sampled in Hawaii (Fig 1B; Table 1A) [START_REF] Balla | A wild C. elegans strain has enhanced epithelial immunity to a natural microsporidian parasite[END_REF]. Note that a recent article assigns this strain to a new species based on genome divergence [START_REF] Reinke | Troemel ER Identification of microsporidia host-exposed proteins reveals a repertoire of large paralogous gene families and rapidly evolving proteins[END_REF]. Eight other microsporidian strains showed 99% SSU rDNA and 95% β-tubulin sequence identities to the corresponding genes of the unnamed Nematocida sp. 1 in GenBank (Table 1B), previously reported in C. briggsae [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF][START_REF] Bakowski | Genome sequence of the microsporidian species Nematocida sp.1 strain ERTm6 (ATCC PRA-372)[END_REF]. This N. sp. 1 group is most closely related to N. parisii in the microsporidian phylogeny and the sequences of both SSU and βtubulin genes gave the same grouping (Figs 2 and3; S2 Fig; Table 3). Because of these new samples of N. sp. 1 and their phylogenetic difference and genetic distance to the N. parisii group, here we describe N. sp. 1 as Nematocida ausubeli n. sp. (see Taxonomy section after the Discussion). Whereas N. ausubeli was so far only reported from C. briggsae (India, Cape Verde [START_REF] Bakowski | Genome sequence of the microsporidian species Nematocida sp.1 strain ERTm6 (ATCC PRA-372)[END_REF]), we also found it in C. elegans and C. remanei, in France, Portugal and Germany (Table 1B; Fig 1B), thus broadening its geographic and host range to several species of the Elegans group of Caenorhabditis from Europe.

The remaining 20 microsporidia strains that we identified are distributed among several other species, including some species in another clade (see below). Thus the dominant

Diversity of Nematocida species

Of the remaining 19 microsporidian strains, nine had a Nematocida species as their top blast hit in GenBank, with similarity between 81% ~86% of SSU rDNA and 74% ~84% of β-tubulin genes. In terms of host and geographical ranges, these microsporidia were found in two C. briggsae strains (Thailand and Guadeloupe), one C. tropicalis strain (Guadeloupe), one C. sp. 42 strain (NIC1041 from French Guiana), three Oscheius tipulae strains (France, Czech Republic, and Armenia), one Rhabditella typhae strain (Portugal) and one Procephalobus sp. strain (JU2895 from Spain). In the phylogenetic analysis of SSU rDNA, the corresponding sequences formed a single clade with N. parisii and N. ausubeli, with Ovavesicula popilliae as sister group within Clade II of the microsporidian phylum (see Fig 2) [START_REF] Vossbrinck | Molecular phylogeny of the Microsporidia: ecological, ultrastructural and taxonomic considerations[END_REF]. In addition, the JUm2807 strain that has been recently described as Nematocida displodere [START_REF] Luallen | Discovery of a natural microsporidian pathogen with a broad tissue tropism in Caenorhabditis elegans[END_REF] is distinct from all of them. From phylogenetic analysis and genetic distance of SSU rDNA genes, these Nematocida strains form four groups. These putative new Nematocida species have a mean genetic distance among them of at least 0.06 (Table 3), while their intra-specific genetic distances are all 0.00 (when several strains were isolated). This inter-group distance is also greater than the distance between N. parisii and N. ausubeli. Hence we describe them below as four new species: Nematocida minor, Nematocida major, Nematocida homosporus and Nematocida ciargi n. spp. (see Taxonomy section).

In terms of the phylogenetic relationships within the Nematocida genus in the SSU rDNA tree, the first outgroup clade to N. parisii + N. ausubeli was formed by JUm2751, JUm2747 and JUm2751, corresponding to N. major (Fig 2). The second branch out was formed by JUm1510 and JUm2772, described here as N. minor. N. ciargi JUm2895 was placed in a basal position to the clade formed by N. parisii, N. ausubeli, N. major and N. minor (Fig 2). At the base of the Nematocida genus, the most externally branching sequences appeared to be those of N. displodere JUm2807, and of N. homosporus JUm1504 and NICm516. All topologies were highly supported, except for the node defining the latter clade of N. homosporus and N. displodere (Fig 2). In the phylogenetic tree based on both genes (SSU rDNA and β-tubulin), N. ciargi was placed at the base of Nematocida genus, while N. displodere and N. homosporus still formed one clade (Fig 3). The phylogenetic tree only based on β-tubulin sequences supported the grouping of strains and overall their relative positions (S2 Fig) , except that the relative placement of N. displodere and N. ciargi was exchanged. The β-tubulin phylogeny has one more branch formed by NICm1041, numbered provisionally N. sp. 7, for which we failed to amplify the SSU rDNA fragment. Whole-genome analysis could be performed in the future to refine these placements.

The Nematocida consensus phylogeny is shown in Fig 3 next to the phylogenetic relationships of the nematode hosts in which they were naturally found (see below for further specificity tests). Although the numbers of samples and species are too low for rigorous testing, the data are at least consistent with the intestinal microsporidia species branching through continuous co-evolution with their nematode host. For example, all intestinal Nematocida species found in Caenorhabditis species form one clade, with a first outgroup including Oscheius and Rhabditella pathogens and a distant outgroup infecting the distant outgroup Procephalobus (Fig 3). The exception is N. displodere that was found a single time, in C. elegans, and corresponds to a change in tissue tropism.

Lifecycle of new Nematocida species

As with previously isolated Nematocida, the newly identified microsporidia appeared to be transmitted horizontally, because a bleaching treatment [START_REF] Stiernagle | Maintenance of C. elegans[END_REF] of infected gravid adults eliminated the infection in the culture and reinfection could be obtained by exposure to spores in the environment. All Nematocida microsporidia stages described here were found exclusively in the intestinal cells and were not detected in the germ line.

As previously described for N. parisii [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF], two main stages could be distinguished by Nomarski optics. First, the meront stage appeared as areas of infected intestinal cells devoid of storage granules. These areas were first small circular regions, then extending to longer grooves. Second, rod-shaped sporoblasts and spores appeared in the intestinal cell cytoplasm. In host cells that were heavily infected with N. parisii and some other species, groups of spores inside vesicles could be seen [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF], possibly derived from spore re-endocytosis [START_REF] Szumowski | Small GTPases promote actin coat formation on microsporidian pathogens traversing the apical membrane of Caenorhabditis elegans intestinal cells[END_REF]. In this study, as described before [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF], all N. parisii and N. ausubeli infections displayed two distinct classes of spore size (Table 1; Fig 4A and4B; S3A Fig) .  N. major and N. minor also displayed two spore size classes. N. major formed slightly longer but thinner spores than N. parisii. N. minor showed however much smaller spores, for each class taken separately (Tables 2 and4; Fig 4C and4D). In contrast, N. homosporus and N. ciargi only have a single class of spore size, with N. homosporus spores having an intermediate size (2.00 ± 0.22 μm long, 0.72 ± 0.12 μm wide) and N. ciargi spores having a smaller size (1.39 ± 0.20 μm long, 0.59 ± 0.13 μm wide). Spore vesicles were observed more frequently with N. homosporus or N. ciargi infections than with other Nematocida infections (Fig 4E and4F).

N. ausubeli being the most commonly found parasite of C. elegans besides N. parisii, we further chose to study its lifecycle by electron microscopy. The ultrastructure by electron microscopy and the deduced lifecycle of N. ausubeli overall resembled those of N. parisii, with possible differences outlined below. High-pressure freezing/freezesubstitution allowed better to visualize lipid membranes compared to room temperature preparation methods. We observed meronts, which are separated from the host cell by a single membrane bilayer, likely pathogen-derived (Fig 5A and 5B;S1A and S1B Fig). Their cytoplasm appeared packed with ribosomes. Some meronts displayed an elongated shape and contained several nuclei (Fig 5B and5L). The membrane enclosing the meronts appeared to darken progressively and intracellular membrane compartments developed, likely corresponding to the progressive transition to a sporont stage (Fig 5C). We further observed sporogony, whereby individual sporoblasts with a single nucleus are formed, each surrounded by a membrane (Fig 5D and 5E;S1A and S1D Fig). We did not observe any nuclear division at this stage (unlike in Enteropsectra longa, where they were easily found; see below). We observed progressive stages of sporogenesis, including formation of the anchoring disk, polaroplast membranes, polar tube, posterior vacuole and spore coat S1C,S1D,S1F and S1G Fig).

In the final stages of sporogenesis and in mature spores that corresponded to the small size class observed in light microscopy, two polar tube coil cross-sections could usually be observed (Fig 5H and 5J;S1L Fig) . A single large spore could be found, which displayed three polar tube coil sections on either side of the spore (six sections in total; Fig 5K). Thus, the tube coiled several times in large spores, instead of once in the small spores. In N. parisii, five polar tube sections were reported on one side of the large spores [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF]; it is thus possible that large spores of N. ausubeli harbor fewer polar tube coils than those of N. parisii (because a single large spore was found in each species, it is however difficult to conclude). The anchoring disk defines the anterior pole of the spore. Below the anchoring disk, the polar tube is lined on either side by polaroplast membranes (visible in Fig 5F and5G). A polar tube cross-section with several layers could be seen in Fig 5J and the posterior turn of the polar tube in S1I Fig. The spores in the host cytoplasm appeared either isolated, or clustered within a large vesicle. Some isolated spores were surrounded by an additional membrane outside the spore coat and the inner face of this membrane appeared coated (Fig 5H). Unlike in N. parisii [START_REF] Szumowski | The small GTPase RAB-11 directs polarized exocytosis of the intracellular pathogen N. parisii for fecal-oral transmission from C. elegans[END_REF], we could not see the additional membrane around all spores. Fig 6A shows a spore apparently exiting the host cell through exocytosis (although we cannot rule out that such images correspond to endocytotic events). Spores in the lumen were not surrounded by any additional membrane (Figs 5K,5L and 6A).

When spores were clustered in a vesicle, two membranes could be observed around them (Fig 5J, and other instances).

Nematode-infecting microsporidia in Clade IV

Whereas the Nematocida genus is in Clade II of the microsporidia [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF][START_REF] Vossbrinck | Molecular phylogeny of the Microsporidia: ecological, ultrastructural and taxonomic considerations[END_REF], the remaining nine microsporidia strains in our collection were placed in Clade IV, which, unlike Clade II, contains several human-infecting microsporidia (Fig 2). This clade assignment was based on SSU rDNA sequences, which had closest (88-89%) identities to the insect parasite Orthosomella operophterae (host: moth Operophtera brumata) (Table 2). Only four β-tubulin sequences could be obtained, and these were closest (75% ~76% identity) to Vittaforma corneae, a human-infecting microsporidia species and a close relative of Orthosomella operophterae (whose β-tubulin sequence is not available), consistent with rDNA analysis. We thus isolated nematode-infecting microsporidia that are in a distinct evolutionary branch compared to Nematocida and are closer relatives of the human-infecting microsporidia.

Eight out of the nine strains in this group have Oscheius species as their nematode host and infect their gut: seven of them from different locations in France were found in O. tipulae, while JUm408 was found in Oscheius sp. 3 [START_REF] ´lix | Three biological species related to Rhabditis (Oscheius) pseudodolichura Ko ¨rner in Osche[END_REF] from Iceland. The ninth strain, JUm1396, was isolated from a C. brenneri strain and is the only one in this set to infect non-intestinal tissues.

In the phylogenetic analysis, these nine strains separated into two groups, corresponding to the two new genera described below, Enteropsectra and Pancytospora (see section on Taxonomy) (Fig 2 ; S4 Fig) . The first group included four strains, JUm408, JUm1456, JUm2551 and JUm1483, which were phylogenetically placed as a sister group to Liebermannia species (with hosts such as grasshoppers) (Fig 2). In the β-tubulin phylogeny, Enteropsectra strains also showed a sister relationship with the group of V. corneae and Enterocytozoon bieneusi, a human intestinal parasite (S2 Fig) . However, with β-tubulin, JUm408 and JUm1483 formed a branch, JUm1456 and JUm2551 another branch, which was different from their SSU rDNA phylogenetic position. Based on molecular sequences, spore morphology and host specificity (below), we describe two species in the Enteropsectra genus, E. longa (type strain JUm408) and E. breve (type strain JUm2551), and do not assign the two other strains to a species. E. longa anchoring disk and the membranes of the polaroplast are visible on the anterior side, chromatin and ribosomes on the posterior side. J. Cross-section of a spore vesicle containing four spores, each showing two polar tube sections (arrowheads). The upper inset shows two membranes around the vesicle (indicated by arrows). The lower inset shows an enlarged multilayered polar tube. K. A large size spore, with two insets showing the posterior vacuole and at least three polar tube coils (three cross-sections on either side of the spore, arrowheads). L. Lower magnification view of several N. ausubeli infection stages in host intestinal cells. Large arrow and small arrow indicate large spore and small spore, respectively. The large spore is that shown in panel K in another plane of section. Arrowheads indicate sporonts. Two multinucleate meronts are indicated. Scale bar is 500 nm, unless indicated otherwise. A, anchoring disk; Chr, chromatin; M, meront; Nu, nucleus; Pa, anterior polaroplast; Pp, posterior polaroplast; Pt, polar tube; Pv, posterior vacuole. doi:10.1371/journal.ppat.1006093.g005 and E. breve strains have a small mean SSU genetic distance of 0.005 (Table 3) but differ in spore size and host specificity (see below). While E. longa and E. breve form a sister group to Liebermannia species on the SSU rDNA phylogeny, they have a smaller mean genetic distance to O. operophterae (0.08) than to Liebermannia (0.11).

The second new clade of nematode-infecting microsporidia includes the five remaining strains and showed strong support as sister lineage to the clade formed by Enteropsectra and [START_REF] Szumowski | The small GTPase RAB-11 directs polarized exocytosis of the intracellular pathogen N. parisii for fecal-oral transmission from C. elegans[END_REF][START_REF] Szumowski | Small GTPases promote actin coat formation on microsporidian pathogens traversing the apical membrane of Caenorhabditis elegans intestinal cells[END_REF], spores appear surrounded by a membrane that fuses with the apical membrane of the host intestinal cell, resulting in the release of spores. We also observed apparently mature spores without an additional membrane and do not know whether they will later acquire a membrane or exit in another manner. Earlier stages were omitted here for simplicity. B. Enteropsectra longa. The top panel is an electron micrograph of Enteropsectra longa spores exiting from the intestinal cell into the lumen, with the host intestinal cell membrane folding out around the E. longa spores (arrow). The diagram below illustrates the exit of E. longa spore from the intestinal cell. The host intestinal cell membrane folds out around the spore until the whole spore exits the cell, after which the host membrane around the spore seems to disappear.

Tissue tropism and lifecycle of nematode-infecting Clade IV microsporidia species

As with Nematocida, all of the infections by Clade IV microsporidian strains mentioned above appeared to be transmitted horizontally, as bleaching of the nematode culture eliminated the infection. The Enteropsectra strains and P. philotis were only observed to infect the intestine of Oscheius nematodes. By contrast, P. epiphaga A striking feature of Enteropsectra strains is their cellular localization within the nematode intestinal cells: Enteropsectra were all observed to form their spores on the apical side of the epithelial cell at first, while meront stages could be seen in a more basal position (Figs 7A, 7B, 7E; 8L). This polarization within the host intestinal cell was not observed in infections of P. philotis nor of any Nematocida species (Table 2; Figs 4 and7).

The Enteropsectra and Pancytospora species displayed quite different sizes and shapes of spores from those of Nematocida species and we did not see any spore-containing vesicles in these microsporidian infections. They all show a single class of spore size. Though apart in the phylogenetic analysis, E. longa (JUm408) and P. philotis share similar dimensions of spores, which are particularly long and thin: E. longa (JUm408) spores measure 3.76 ± 0.38 μm by 0.49 ± 0.06 μm, while P. philotis spores measure 3.46 ± 0.48 μm long by 0.42 ± 0.06 μm. These spores are even longer than the largest spores and thinner than the smallest spores in Nematocida. In stark contrast, E. breve (JUm2551) form small rod-shaped and crescent-shaped spores (Fig 7B ;Table 4).

Because of the striking difference in spore distribution, we further analyzed by electron microscopy the type species of the Enteropsectra genus, Enteropsectra longa (JUm408) in Oscheius sp. 3 JU408. The meront stage appeared overall similar to that of Nematocida species: the early stages displayed a cytoplasm packed with ribosomes and very few membranes (Fig 8A ); elongated multinucleated meronts could also be observed (Fig 8B). The parasite membrane then progressively darkened, indicating the transition to the sporont stage (Fig 8C -8F). By electron microscopy, we observed a potential key difference in the exit mode of the spores between Enteropsectra longa (JUm408) on one hand, and N. parisii and N. ausubeli on the other hand. First, the sporoblasts and mature spores of E. longa were never seen to be surrounded by an additional membrane outside the spore wall, precluding exocytosis as an exit route. Second, the spores were seen to protrude on the apical side of the host cell, pushing out the host cell membrane like a finger in a glove ( On the host side, rough endoplasmic reticulum was often seen to wrap around sporoblasts, yet never encircling them fully (Fig 8G). The host cell nuclei presented a characteristic nucleolar structure, which became organized in long tubules (often appearing circular in cross-sections;. S6J and S6K Fig) . On one occasion, microsporidian spores were observed within the host intestinal cell nucleus, whose nucleolus had apparently further degenerated (S6L Fig).

Figures of intranuclear mitosis

Host specificity: natural associations and laboratory infection tests

The pattern of natural association revealed an apparent specificity of a given microsporidian species for a nematode genus, mostly Caenorhabditis versus Oscheius in our collection. Strikingly, N. parisii, N. ausubeli and N. major infections were found in Caenorhabditis species, while N. minor, N. homosporus and Clade IV microsporidia species infections were all found in Oscheius species and not in Caenorhabditis (or, for N. homosporus, in Rhabditella, a closer relative of Oscheius compared to Caenorhabditis; Fig 3). The notable exception in Clade IV was the epidermal P. epiphaga JUm1396, found in C. brenneri. These results suggested a pattern of host-pathogen specificity between nematode and nematode-infecting microsporidia.

We further complemented these natural associations with infections performed in the laboratory. To test for the capacity of a given microsporidia strain to infect a given host, uninfected nematode cultures (cleaned by bleaching) were exposed to microsporidian spores. We used clean spore preparations from seven microsporidian species (see Materials and Methods), namely N. parisii, N. ausubeli, N. major, N. homosporus, E. longa, E. breve and P. philotis. On the host side, we focused on four nematode species of two genera: C. elegans, C. briggsae, O. tipulae and O. sp. 3, all of which reproduce through self-fertilizing hermaphrodites and facultative males [START_REF]Oscheius tipulae[END_REF]20]. We favored wild strains that had been found naturally infected with microsporidia and were thus not generally resistant to microsporidian infections (Table 5).

N. parisii (JUm2816) infected more than 50% of C. elegans (N2, JU2009) and C. briggsae (JU2747, JU2793) individual animals at 72 hpi. However, no microsporidian infection symptom was observed in O. tipulae (JU1483, JU170) nor O. sp. 3 (JU408, JU75) at 72 and 120 hpi (Table 5). O. tipulae strains JU1504, JU1510 and JU2552 were also exposed to N. parisii spores, and none of them became infected either. These infection results indicated that N. parisii was unable to infect O. tipulae nor O. sp. 3 (Table 5). Specificity of N. ausubeli (JUm2009) slightly differed from that of N. parisii. By 72 hpi, half of all Caenorhabditis animals and about 30% of O. sp. 3 showed signs of infection. None of O. tipulae worms were infected even at 120 hpi (Table 5). However, when we made a new N. Wild Caenorhabditis brenneri strain JU1396, with Pancytospora epiphaga infection. Spores are seen in the epidermal cells in the tail that does not contain any gut tissue (posterior to the rectum). Anterior is to the right. The "fur" on the outside of the cuticle is formed by unidentified bacteria (see [START_REF] Hodgkin | Two Leucobacter strains exert complementary virulence on Caenorhabditis including death by worm-star formation[END_REF] ausubeli (JUm2009) spore preparation and used it directly for infection tests, O. tipulae strains JU1510 and JU2552 could be infected, but the preparation lost its ability to infect O. tipulae over storage at -80˚C (see Materials and Methods). We conclude that O. tipulae was far less susceptible to infection than C. elegans, C. briggsae and O. sp. 3 to N. ausubeli infection.

The host spectrum of N. major (JUm2747) was quite similar to that of N. parisii: it infected C. elegans (N2, JU2009) and C. briggsae (JU2507, JU2793), but not O. tipulae (JU1483, JU170) nor O. sp. 3 (JU408, JU75) (Table 5). N. homosporus, however, could infect both Caenorhabditis and both Oscheius species and thus appeared as the most generalist (Table 5). Yet O. tipulae seemed relatively less sensitive than C. elegans, C. briggsae and O. sp. 3 to N. homosporus infection.

Enteropsectra spp. and Pancytospora philotis showed different and even opposite specificities compared to the four tested Nematocida species. Indeed, none could successfully infect any tested Caenorhabditis strains at 120 hpi. Within the two Oscheius species, specific interactions were further observed. Enteropsectra longa (JUm408) only infected O. sp. 3 strains (JU408, JU75), but not O. tipulae (JU1483, JU2551), while E. breve (JUm2551) infected all four O. tipulae and O. sp. 3 strains (Table 5). Pancytospora philotis (JUm1505) only infected O. tipulae (JU1483, JU1505), but not O. sp. 3 strains (JU408, JU75). Since O. sp. 3 is the closest known species to Oscheius tipulae, E. longa and P. philotis are examples of narrow specialization in the host-parasite interaction. We also found that C. elegans N2 could be infected with P. epiphaga (JUm1396), showing epidermal and muscle infection (Table 5; S7F Fig).

The spore morphology of a given microsporidian species was maintained in different nematode species, indicating that host genotype does not affect this pathogen phenotype. For instance, O. tipulae (JU1510) infected with N. ausubeli (JUm2526) displayed two sizes of spores in its intestinal cells as upon Caenorhabditis infection by N. ausubeli (S7A Fig) . Oscheius sp. 3 (JU408) infected with Enteropsectra breve (JUm2551) formed small rod-shaped or crescentshaped spores along the apical side of the worms' intestinal cells, as upon O. tipulae infections (S7D Fig) . C. elegans N2 infected with P. epiphaga (JUm1396) formed long and thin spores in the epidermis and muscles, as upon C. brenneri infection (S7F Fig) .   N. ausubeli elicits a less robust host transcriptional response than other Nematocida species, despite establishing a robust infection Given the capacity of all Nematocida species to infect C. elegans, we next sought to compare the C. elegans response to infection among our newly isolated microsporidia species. N. parisii infection in C. elegans has been shown to induce a broad transcriptional response [START_REF] Bakowski | Ubiquitin-mediated response to microsporidia and virus infection in C. elegans[END_REF]. Among genes that were highly upregulated at all infection timepoints were C17H1.6 and F26F2.1, two genes of unknown function. Two transgenic C. elegans strains, ERT54 and ERT72, were generated as transcriptional reporters for these two genes and have been previously shown to be strongly induced in early N. parisii and N. displodere infection [START_REF] Bakowski | Ubiquitin-mediated response to microsporidia and virus infection in C. elegans[END_REF]. We tested these reporter strains with our new microsporidia species by placing them onto plates with a culture of infected worms and microsporidian spores, then monitoring GFP expression at different timepoints in the reporter strains, as well as monitoring microsporidian meront and spore formation. As expected, N. parisii, N. ausubeli, N. major and N. homosporus could all infect these reporter strains, forming meronts and spores, and induce reporter GFP expression. By contrast, E. longa and Enteropsectra JUm1483 failed to show evidence of proliferative infection and did not robustly induce reporter expression (Fig 9A and 9B;S2 Table). Most interestingly, while N. parisii, N. major or N. homosporus consistently induced the GFP reporters, different strains of N. ausubeli (JUm2009, ERTm2, ERTm6; Fig 9;S2 Table) did not, although this species did robustly infect and proliferate within the C. elegans intestine.

To verify that this differential induction of the GFP reporters matched the transcripts of the endogenous genes, we conducted qRT-PCR after controlled N. parisii (ERTm1) and N. ausubeli (ERTm2) infections of N2 using purified spore preparations that were normalized for an equivalent level of invasion (see Materials and Methods). Indeed, we saw that both C17H1.6 and F26F2.1 transcripts, along with another gene highly induced by N. parisii infection W04B5.5 [START_REF] Cuomo | Microsporidian genome analysis reveals evolutionary strategies for obligate intracellular growth[END_REF], were induced approximately 6-7-fold lower upon N. ausubeli infection compared to N. parisii infection (Fig 9C ), while the levels of pathogen rRNA (indicative of pathogen load) remained similar. Thus, N. ausubeli infection caused a much reduced host response compared to other Nematocida species (Fig 9;S2 Table), despite causing an equivalent, or even more robust infection [START_REF] Balla | Cell-to-cell spread of microsporidia causes C. elegans organs to form syncytia[END_REF]. Considering the phylogenetic relationships of the Nematocida 5. C. Transcript levels for three genes were measured after 4 hours of infection of N2 C. elegans by N. parisii (ERTm1) and N. ausubeli (ERTm2). The fold increase in transcript level was measured relative to uninfected N2 levels. Infection dose was normalized between Nematocida by successful invasion events counted as intracellular sporoplasms at 4 hpi. To independently compare the microsporidian doses in parallel to the transcript quantification, we also measure the levels of Nematocida SSU rRNA after 4 hours of infection of C. elegans in the same experiment: we found that the rRNA level measured after infection with ERTm2 was 1.25-fold higher than that with ERTm1.

Discussion

Independent evolutionary branches of nematode parasitism by microsporidia

Microsporidia are ubiquitous obligate intracellular pathogens that have agricultural and medical significance, but have been difficult to study in the laboratory. Our study provides a collection of microsporidia that can infect bacteriovorous nematodes and can easily be studied in the laboratory in their natural hosts and in related species. These rhabditid nematode-infecting microsporidia seem to have more than one origin within the Microsporidia phylum: at least one origin within Clade II and one or two within Clade IV. We thus here enlarge considerably the spectrum of microsporidia that can be cultured in nematodes, including some that are genetically close to human pathogens in Clade IV. Environmental SSU rDNA microsporidian sequences have been reported from soil, sand and compost samples from North America [START_REF] Ardila-Garcia | Microsporidian diversity in soil, sand, and compost of the Pacific Northwest[END_REF]. (The corresponding species have not been named.) Some of them branch in the SSU phylogeny in the vicinity of the nematode-infecting microsporidia that we isolated (S4 Fig) . Specifically, some branch close to Nematocida homosporus and some may be outgroups to Nematocida or further species of the genus. In Clade IV, one is closely related to the Pancytospora epiphaga JUm1396 sequence.

The clades of nematode-infecting microsporidia that we describe have close relatives that infect arthropods, especially insects. This relationship may be due to deep co-evolution (arthropods and nematodes being close relatives on the animal phylogeny), or to the fact that nematodes share their habitats and interact with insects by using them as hosts or carriers [START_REF] ´lix | The natural history of Caenorhabditis elegans[END_REF], which may have facilitated a host shift or a complex lifecycle with several hosts. The microsporidia described here can be cultured continuously in their nematode hosts, but we cannot rule out the possibility that some of them may use non-nematode hosts as well, including insects. Of note, all of them use a horizontal mode of transmission, despite the fact that many instances of vertical transmission of microsporidia in arthropods, molluscs and fish are known [START_REF] Smith | The ecology and evolution of microsporidian parasites[END_REF][START_REF] Vavra | Microsporidia and 'The art of living together[END_REF]. In addition, Nematocida species are diploid with evidence of recombination and thus possibly a sexual cycle [START_REF] Bakowski | Genome sequence of the microsporidian species Nematocida sp.1 strain ERTm6 (ATCC PRA-372)[END_REF][START_REF] Cuomo | Microsporidian genome analysis reveals evolutionary strategies for obligate intracellular growth[END_REF], which might occur in another host.

N. parisii, N. ausubeli and N. major are relatively common pathogens of Caenorhabditis but not of Oscheius

Our results suggest that infections by N. parisii and N. ausubeli are quite common in wild Caenorhabditis strains, especially in C. elegans and C. briggsae. In our collection, N. parisii, N. ausubeli and N. major infections were found in 30 strains of four Caenorhabditis species. Though we have a sampling bias towards France, N. ausubeli was found in Asia, Europe and Africa, while N. parisii was found mostly in France and once (ERTm5) from Hawaii. N. major was only found from three Caenorhabditis strains of C. briggsae and C. tropicalis, all of which were sampled in tropical areas, despite the fact that that we have sampled many hundreds of C. elegans isolates and that N. major can easily infect C. elegans in our specificity infection tests (Table 5). A possibility is that N. major may be preferentially distributed in the tropics rather than temperate zones, where C. elegans are mostly found (Table 2, Fig 1B) [START_REF] ´lix | The natural history of Caenorhabditis elegans[END_REF].

In addition to C. elegans and C. briggsae strains, we also have a relatively large collection of microsporidia-infected Oscheius strains (10 O. tipulae strains and one O. sp. 3 strain). However, none of these strains was found with Nematocida or N. major infections. In line with their natural associations, N. parisii and Nematocida major were not able to infect any Oscheius strains in the laboratory. These specializations may be due to long-term coevolution and adaptation processes [START_REF] Antonovics | The origin of specificity by means of natural selection: evolved and nonhost resistance in host-pathogen interactions[END_REF].

In addition, one new microsporidian species infecting Caenorhabditis was found in clade IV, Pancytospora epiphaga. As this Clade IV microsporidian species can infect C. elegans, it would be interesting to develop its study as a model system for Clade IV species infection.

Diverse microsporidia infect Oscheius species

Microsporidian species that naturally infect Oscheius species are diverse (Fig 10,green entries). N. minor, found from two O. tipulae strains, forms two distinct sizes of spores, similar to N. parisii, N. ausubeli and N. major. N. homosporus was found from one O. tipulae strain and one R. typhae and is the only species tested here that is able to infect species of three genera Caenorhabditis, Oscheius and Rhabditella, suggesting that N. homosporus may be a relatively less specific pathogen for rhabditid nematodes.

The Clade IV Oscheius-infecting microsporidia are separated into two groups: Enteropsectra species, and Pancytospora philotis. None of those could infect Caenorhabditis and their host specificity is even narrower, distinguishing between Oscheius tipulae and its sister species Oscheius sp. 3. The SSU rDNA genetic distances between E. longa and E. breve are quite small and two other closely related Enteropsectra strains are also available (Tables 2 and3). Overall, Enteropsectra and the Tipulae group of Oscheius species [START_REF]Oscheius tipulae[END_REF][START_REF] ´lix | Three biological species related to Rhabditis (Oscheius) pseudodolichura Ko ¨rner in Osche[END_REF] provide an interesting case to study the evolution of a narrow host specificity.

Evolutionary changes in tissue tropism

Although microsporidia are known to be able to adopt either a horizontal or a vertical transmission [START_REF] Smith | The ecology and evolution of microsporidian parasites[END_REF][START_REF] Dunn | Microsporidian life cycles and diversity: the relationship between virulence and transmission[END_REF], we here only observed infection in somatic tissues and transmission was horizontal. Most of the infections occurred in host intestinal cells, while two independent instances showed infections elsewhere. As reported previously, Nematocida displodere can infect many tissues and cells in C. elegans, including the epidermis, muscle, coelomocytes and neurons, although it appears to invade all cells by firing its polar tube from the intestinal lumen [START_REF] Luallen | Discovery of a natural microsporidian pathogen with a broad tissue tropism in Caenorhabditis elegans[END_REF]. The second independent case is Pancytospora epiphaga can be seen in the epidermis, coelomocytes and muscles. Whether it also enters the nematode's cells through the gut remains to be studied.

Cellular exit strategies

The most striking variation we observed concerns the cellular exit strategies of the spores ( Fig [START_REF] Stentiford | Microsporidia-Emergent pathogens in the global food chain[END_REF]. Nematocida parisii spores acquire an additional membrane around the spore wall and thus exit through a vesicular pathway, using the host exocytosis machinery [START_REF] Szumowski | The small GTPase RAB-11 directs polarized exocytosis of the intracellular pathogen N. parisii for fecal-oral transmission from C. elegans[END_REF]; in addition, clusters of spores with two additional membranes were observed. If the process is similar in N. ausubeli to that in N. parisii, the spore clusters may correspond to re-endocytosis of spores from the lumen [START_REF] Szumowski | Small GTPases promote actin coat formation on microsporidian pathogens traversing the apical membrane of Caenorhabditis elegans intestinal cells[END_REF] or perhaps to autophagy of internal spores using the apical plasma membrane. Of note, the host rough endoplasmic reticulum could often be seen to form concentric patterns in the intestinal cell cytoplasm (S1B and S1E Fig), sometimes wrapping around the sporoblasts (Fig 5E). Whether the reticulum may be a precursor for the additional membranes through an autophagic pathway [START_REF] Tooze | The origin of the autophagosomal membrane[END_REF][START_REF] Sica | Organelle-specific initiation of autophagy[END_REF], is an alternative possibility. By contrast, in Enteropsectra longa, the sporoblasts and mature spores were never seen surrounded by an additional membrane, which rules out exocytosis as an exit route. Instead, the Whether the final release step was by pinching of the plasma membrane at the base or by rupturing it is unclear, although the former is more probable, given that the intestinal cells were not seen to leak out. We observed spore sections in the lumen, far from any intestinal cells in the corresponding section, with either an additional membrane around them or none. A possible scenario is that the spores are first released with a membrane, and that the membrane then disintegrates (Fig 6B). Yet because we did not follow by serial sectioning the length of the spores, we cannot know for sure that those with a membrane were not still attached to the epithelial cell. We thus cannot rule out an alternative mechanism whereby the spores are released through a hole in the plasma membrane-although given spore size, this latter exit mechanism would likely lead to host cell rupture and death, an event that was never observed. Of note, another exit mode was noted in the human gastrointestinal microsporidia Enterocytozoon bieneusi (in Clade IV like E. longa), whereby the infected cell itself is extruded in the lumen [START_REF] Peacock | Histological diagnosis of intestinal microsporidiosis in patients with AIDS[END_REF][START_REF] Mansfield | Identification of an Enterocytozoon bieneusi-like microsporidian parasite in Simian-Immunodeficiency-Virus-inoculated macaques with hepatobiliary disease[END_REF][START_REF] Vavra | Structure of microsporidia[END_REF]. Presumably, the cell then rapidly dies and the spores are released by disintegration of the enterocyte plasma membrane. In the present case of Enterospectra longa, the epithelial intestinal cell remains overall intact and only the spore exits, possibly with the surrounding enterocyte plasma membrane that then disintegrates.

A diverse collection of natural host/microsporidia pairs in C. elegans and related wild-caught nematodes

Beyond access to a diversity of microsporidia, our collection of host-parasite combinations also provides a resource for defining the genetic basis of host resistance. Most current work on C. elegans and N. parisii is performed using the C. elegans reference strain N2 and the N. parisii ERTm1 isolate, yet this strain combination has been shown to lead to a very strong infection where the host does not mount an effective defense response (e.g. in comparison with C. elegans CB4856; [START_REF] Balla | A wild C. elegans strain has enhanced epithelial immunity to a natural microsporidian parasite[END_REF]), thus making it a difficult system in which to identify immune defense pathways. The present collection offers many further possibilities of genetic screens using induced mutations or natural genetic variation for resistance pathways.

Conclusion

Overall, we here considerably enlarged the resources and knowledge on the microsporidia infecting bacteriovorous terrestrial nematodes. These microsporidia are diverse in terms of phylogenetic relationships, spore size and shape, the presence of vesicles containing spores, host specificity pattern, host tissue tropism, host cell intracellular localization and cellular exit route.

Materials and Methods

Nematode sampling, isolation and microsporidia strains

Hundreds of samples, mostly from rotting fruits, rotting stems and compost, were collected worldwide over several years, and nematodes were isolated as described [START_REF] Barrière | Isolation of C. elegans and related nematodes[END_REF]. The nematode species was identified as described [START_REF] Barrière | Isolation of C. elegans and related nematodes[END_REF][START_REF] Fe ´lix | A streamlined system for species diagnosis in Caenorhabditis (Nematoda: Rhabditidae) with name designations for 15 distinct biological species[END_REF], using a combination of morphological examination (dissecting microscope and Nomarski optics), molecular identification (18S, 28S or ITS rDNA) and mating tests by crossing with close relatives. Isogenic nematode strains were established by selfing of hermaphrodites or for obligate male-female species from a single mated female. Individuals of strains showing a paler intestinal coloration (Fig 1A) were examined by Nomarski optics. Strains with meronts and spores in the intestinal cells or elsewhere were labeled as suspected to harbor a microsporidian infection. Each nematode strain was then frozen and stored at -80˚C. both genes) as outgroups (Figs 2 and 3; S2 Fig) [START_REF] Vossbrinck | Molecular phylogeny of the Microsporidia: ecological, ultrastructural and taxonomic considerations[END_REF]. For phylogenetic analysis of SSU rDNA genes, 28 out of 33 sequences obtained from this study were compared with 11 sequenced Nematocida isolates (ERTm1, ERTm2, ERTm3, ERTm5, ERTm6, JUm1248, JUm1249, JUm1253, JUm1762, JUm1893, and JUm2807), JUm1396, 60 other microsporidian species chosen from all five major clades of microsporidia [START_REF] Vossbrinck | Molecular phylogeny of the Microsporidia: ecological, ultrastructural and taxonomic considerations[END_REF] and two Rozella species [START_REF] James | Reconstructing the early evolution of Fungi using a six-gene phylogeny[END_REF]. For analysis of β-tubulin genes, only sequences from the six Nematocida species (ERTm1, ERTm2, ERTm3, ERTm5, ERTm6 and JUm2807) and 18 other published microsporidian species were available to be compared with our 32 sequences (S1 Table ). To phylogenetically analyze both genes together, we concatenated the two genes of stains ERTm1~6, JUm2807, our 30 strains, 10 other microsporidia species and two outgroups (B. ranarum and C. coronatus). Sequences were aligned using Geneious v7.1.7 with default parameters and further aligned manually and concatenated if available. The alignments were imported to MEGA 6 [START_REF] Tamura | MEGA6: molecular evolutionary genetics analysis version 6.0[END_REF] to estimate the best DNA evolution models and compute mean genetic distances (1000 bootstrap replicates). Bayesian inference phylogenies were constructed using Mesquite v3.04 [START_REF] Maddison | Mesquite: a modular system for evolutionary analysis[END_REF] and MrBayes v3.2.2 [START_REF] Huelsenbeck | MRBAYES: Bayesian inference of phylogenetic trees[END_REF], with the same DNA models as above [START_REF] Bast | Sequence similarity search, multiple sequence alignment, model selection, distance matrix and phylogeny reconstruction[END_REF] and refined by FigTree v1.4.2 (http:// tree.bio.ed.ac.uk/software/figtree/).

High-pressure freezing and transmission electron microscopy

Worms were frozen in M9 buffer [START_REF] Stiernagle | Maintenance of C. elegans[END_REF] supplemented with 20% BSA (Type V) in the 100 μm cavity of an aluminium planchette, Type A (Wohlwend Engineering, Switzerland) with a HPM 010 (BalTec, now Abra Fluid AG, Switzerland). Freeze substitution was performed according to [59] in anhydrous acetone containing 2% OSO4 + 2% H 2 O in a FS 8500 freeze substitution device (RMC, USA). Afterwards samples were embedded stepwise in Epon. To achieve a good infiltration of spores, the infiltration times in pure resin were prolonged for 48 h compared to the published protocol. After heat polymerization thin sections of a nominal thickness of 70 nm were cut with a UC7 microtome (Leica, Austria). Sections were collected on 100 mesh formvar coated cupper grids and poststained with aqueous 4% uranylacetate and Reynold's lead citrate. Images were taken with a Tecnai G2 (FEI, The Netherlands) at 120 kV and equipped with a US4000 camera (Gatan, USA).

Spore size measurements

Spore size was measured as described [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF]. Briefly, infected nematodes were photographed by Nomarski optics and spores were measured using the Image J software [START_REF] Abràmoff | Image processing with ImageJ[END_REF]. We only took into account spores with a clear outline within the focal plane. In species with two spore size classes, large spores are less numerous than small ones and they are found in groups. When measuring, the spores were first assigned to a size class, in part based on the spatial clustering of large spores. 20 spores were measured for each spore type; except N. ausubeli, for which 42 small ones and 40 large ones were measures.

Microsporidia spore preparation

For the microsporidian spore preparation, we first tried the methods previously established for N. parisii and N. ausubeli [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF][START_REF] Estes | Non-lytic, actin-based exit of intracellular parasites from C. elegans intestinal cells[END_REF]. Because wild nematodes naturally live in habitats with various microbes [START_REF] ´lix | The natural history of Caenorhabditis elegans[END_REF][START_REF] Cutter | Caenorhabditis evolution in the wild[END_REF], the microsporidia-infected nematode cultures generally originally contained other microbes, such as bacteria, fungi, or even viruses. In order to obtain a relatively pure microsporidian spore preparation, we treated the nematode cultures repeatedly with antibiotics (100 ug/ml gentamycin, 50 ug/ml Ampicillin, 50 ug/ml Kanamycin, 20 ug/ml Tetracycline, and 50 ug/ml streptomycin), monitoring the presence of non-E. coli bacteria and fungi on the plate. Nematode strains do not lose the microsporidian infection after antibiotic treatment. After antibiotic treatment, if the appearance of a plate with infected worms looks like those with bleached worms, we considered the plate to be clean and the infected worms were used to extract clean spores. Even though inconspicuous microbes may still be carried over, as we know so far, none of them could prevent the worms from getting infected with microsporidia nor induce similar symptoms as microsporidia.

Antibiotic-cleaned worms without other detectable microbes were harvested in 2-ml microfuge tube and autoclaved silicon carbide beads (1.0 mm, BioSpec Products, Inc.) were added. The tube was then vortexed for 5 min at 2,500 rpm and the lysate of worms filtered through a 5 μm filter (Millipore) to remove large worm debris. Spore concentration was quantified by staining with chitin-staining dye direct yellow 96 (DY96).

This method worked well on N. major and N. homosporus, but spores of Clade IV species extracted this way could not infect any worms. To prepare infectious spores of these species, we used instead a plastic pestle to crush worms manually, and stored these spore preparations at 4˚C.

Nematocida species spore preparations could generally be stored at -80˚C for later infection tests. However, storage at -80˚C could affect the infection efficiency of these spore preparations. Indeed, when we made a fresh N. ausubeli (JUm2009) spore preparation and used it directly for infection tests, it could infect O. tipulae strains JU1510 and JU2552, with meronts and spores found in their intestinal cells at 120hpi. One month later, we used the same batch that had been stored at -80˚C to infect C. elegans (N2), O. tipulae (JU1483, JU170, JU1510 and JU2552). At 120 hpi, 100% of N2 adult worms were infected, while none of the O. tipulae strains became infected. These results suggested that this spore preparation became less infectious after being frozen and stored at -80˚C for one month, which did not compromise infection in C. elegans but did compromise infection of O. tipulae. For further specificity tests, spore preparations of N. major, N. homosporus and Clade IV species were then used within two hours after extraction, without freezing.

Infection assays

20 uninfected L4 or young adults (i.e. prior to first egg formation) were transferred to a 6 cm NGM plate seeded with E. coli OP50. 5 million microsporidian spores in 100 μl distilled water were placed on the E. coli lawn. The cultures were then incubated at 23˚C. The infection symptoms of 20 adults were checked by Nomarski optics at 72 hours after inoculation. If no infection symptoms were found at this timepoint, they were scored a second time at 120 hours post-inoculation.

Assays with reporter strains

Two transgenic C. elegans strains, ERT54 jyIs8[C17H1.6p::gfp; myo-2p::mCherry] and ERT72 jyIs15[F26F2.1p::gfp;myo-2::mCherry] were used in infection assays to test infection specificity and transcriptional response of C. elegans to different microsporidian infections. These two lines express a constitutive fluorescent Cherry marker in the pharyngeal muscles and induce GFP upon infection with N. parisii [START_REF] Bakowski | Ubiquitin-mediated response to microsporidia and virus infection in C. elegans[END_REF]. In the first qualitative assay (23˚C), we focused on the ERT54 strain. First, 10 L4 stage animals from seven naturally infected strains (C. elegans JU1762 with N. parisii infection, C. elegans JU1348 with N. ausubeli, C. briggsae JU2507 with N. major, O. tipulae JU1504 with N. homosporus, R. typhae NIC516 with N. homosporus, O. tipulae JU1483 with Enteropsectra, Oscheius sp. 3 JU408 with E. longa) were transferred to new plates and cultured for two days, in order to release microsporidian spores onto the plates. Then 10 L4 stage worms of the ERT54 strain were added onto these plates and onto a clean plate as control. Two days post-inoculation (dpi), a chunk was transferred to new plate to prevent starvation. One day later (3 days dpi), GFP expression of ERT54 animals (visualized using the Cherry reporter in the pharynx) and infection symptoms were scored. 20 worms showing GFP expression (if any, else the Cherry marker was used) were picked and transferred to a new clean plate. GFP expression was monitored on 8 dpi and 14 dpi. In the second quantitative assay (23˚C), first, 10 L4 stage animals from five naturally infected strains (C. briggsae JU2055 with N. parisii infection, C. elegans JU2009 with N. ausubeli, C. briggsae JU2507 with N. major infection, R. typhae NIC516 with N. homosporus infection, Oscheius sp. 3 JU408 with E. longa infection) and uninfected C. elegans reference strain N2 (as negative control) were transferred to new plates and cultured for three days. Then 200 L4 stage worms of ERT54 or ERT72 were added. GFP expression of 50 worms (if possible) of reporter strains was monitored at five different timepoints (2 hours post inoculation (hpi), 4 hpi, 8 hpi, 28 hpi, 48 hpi) and infection symptoms were scored at 48 hpi.

qRT-PCR of reporter transcripts

For measurements of transcripts levels by quantitative RT-PCR (qRT-PCR) (primers used see S2 Table ), 3000 synchronized N2 C. elegans L1 larvae were infected for 4 hours at 25˚C with 5.0 x 10 5 ERTm1 (N. parisii) spores and 1.5 x 10 6 ERTm2 (N. ausubeli) spores. Prior analysis of serial spore dilutions determined that these ERTm1 and ERTm2 spore doses resulted in an average of 1 sporoplasm per L1 larva at 4 hpi at 25˚C as measured by FISH to Nematocida rRNA. At 24 hpi, animals were harvested and RNA was isolated by extraction with Tri-Reagent and bromochloropropane (BCP) (Molecular Research Center). cDNA was synthesized from 175 ng of RNA with the RETROscript kit (Ambion) and quantified with iQ SYBR Green Supermix (Bio-Rad) on a CFX Connect Real-time PCR Detection System (Bio-Rad). Transcript levels were first normalized to the C. elegans snb-1 gene within each condition. Then transcript levels between conditions were normalized to uninfected N2 for C. elegans transcripts or normalized to ERTm1 rRNA for Nematocida rRNA.

Taxonomic section

Rationale for the description of new microsporidia taxa. We describe here two new genera and nine new species of microsporidia based on rDNA and β-tubulin sequences and phenotypic analyses.

The rDNA (and β-tubulin, when we could amplify it) sequences could be readily grouped in three distinct clades, one including Nematocida parisii and many of our strains in microsporidia clade II, and the two other clades in microsporidia clade IV (Figs 2 and 3;S2 Fig).

All described microsporidian species infecting nematodes are reviewed in [START_REF] Bakowski | Microsporidia infections in Caenorhabditis elegans and other nematodes[END_REF]. Previously described species with associated SSU rDNA sequences are Nematocida parisii [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF], Nematocida displodere [START_REF] Luallen | Discovery of a natural microsporidian pathogen with a broad tissue tropism in Caenorhabditis elegans[END_REF], Sporanauta perivermis [START_REF]Microsporidian infection in a free-living marine nematode[END_REF] and Nematocenator marisprofundi [START_REF] Sapir | Microsporidia-nematode associations in methane seeps reveal basal fungal parasitism in the deep sea[END_REF], the two latter infecting marine nematodes. Compared to the species studied here, S. perivermis is found in another group of clade IV, while N. marisprofundi appears as a distant outgroup [START_REF] Sapir | Microsporidia-nematode associations in methane seeps reveal basal fungal parasitism in the deep sea[END_REF] (Fig 2). Our strains are thus all distinct from the two latter species. In addition, two species were reported without any associated molecular sequence [START_REF] Bakowski | Microsporidia infections in Caenorhabditis elegans and other nematodes[END_REF]. The first one, Thelohania reniformis, infected the intestine of a parasitic nematode with a single class of spores of a size exceeding in length and/or width any of those we describe [START_REF] Bakowski | Microsporidia infections in Caenorhabditis elegans and other nematodes[END_REF]. The second species, of an undefined genus ("Microsporidium" rhabdophilum), infected the pharyngeal glands, hypodermis and reproductive system of Oscheius myriophila [START_REF] Poinar | Microsporidium rhabdophilum sp. n. (Microsporida: Pansporoblastina), a parasite of the nematode, Rhabditis myriophila (Rhabditina: Rhabdidae)[END_REF], and does not match in tissue tropism and spore morphology any of the present species.

The biological species concept cannot be used in describing these microsporidia as their sexual cycle is unknown and thus we cannot test their crossing ability. Microsporidia species have been classically delimited through their morphology and their association with a host. In recent years, DNA sequences have further helped to assess phylogenetic relationships among microsporidia, and to assign strains to a species when morphology was not sufficient [START_REF] Stentiford | Plastic paraistes: Extreme dimorphism creates a taxonomic conundrum in the phylum Microsporidia[END_REF]. Among our strains, as a first example, two close groups of strains in the Nematocida clade correspond to N. parisii and N. sp. 1 in [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF], respectively. These groups are consistently distinct from each other by molecular analysis of rDNA and β-tubulin genes (Table 3;S3 Table) but do not appear very different from spore size and general morphology ( [START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF], this work). Their molecular distance (0.017) is consistent with molecular distances between microsporidian species and even greater than other examples of interspecific distance for this gene [START_REF] Sokolova | Development, ultrastructure, natural occurrence, and molecular characterization of Liebermania patagonica n. g., n. sp., a microsporidian parasite ot the grasshopper Tristira magellanica (Orthoptera: Tristiridae)[END_REF]. Their whole genome sequence [START_REF] Cuomo | Microsporidian genome analysis reveals evolutionary strategies for obligate intracellular growth[END_REF] further shows that the two species are wide apart, with only 62% amino-acid identity between protein orthologs, while strains of the same species are much closer, such as 0.2% difference at the nucleotide level between N. parisii ERTm1 and ERTm3 and 1 SNP every 989 bp for N. sp. 1 ERTm2 and ERTm6 [START_REF] Bakowski | Genome sequence of the microsporidian species Nematocida sp.1 strain ERTm6 (ATCC PRA-372)[END_REF][START_REF] Cuomo | Microsporidian genome analysis reveals evolutionary strategies for obligate intracellular growth[END_REF] We therefore formally describe here N. sp. 1 as a new species and call it N. ausubeli n. sp.

Concerning the other strains in the Nematocida clade, given their greater molecular distance to each other, we define four other Nematocida species that are also distinct from N. displodere [START_REF] Luallen | Discovery of a natural microsporidian pathogen with a broad tissue tropism in Caenorhabditis elegans[END_REF]. In this case, each of them further shows a distinct spore morphology (Table 4;Fig [START_REF] Nkinin | Microsporidian infection is prevalent in healthy people in Cameroon[END_REF]. No other described microsporidian species to our knowledge has a similar sequence nor host distribution. We thus describe them below as four new species of Nematocida, namely N. major n. sp. (two sizes of spores, each slightly larger than the respective class in N. parisii and N. ausubeli), N. minor n. sp. (two sizes of spores, each smaller than the respective class in N. parisii and N. ausubeli), N. homosporus n. sp. (a single class of spores) and N. ciargi n. sp. (a single class of spores, particularly small), each with their reference strain. The two latter species were not found in Caenorhabditis nematodes but in other bacteriovorous terrestrial nematodes. We could not amplify the SSU gene of Nematocida "sp. 7" NICm1041 and therefore refrain from formally describing this putative new species.

The remaining strains of microsporidia in our sampling do not belong to clade II but to clade IV. By blast of the rDNA sequence, they are closest to Orthosomella operophterae, an insect pathogen, and by phylogenetic analysis they form two clades. One clade includes JUm408, JUm1456, JUm1483 and JUm2551, and is sister to Liebermannia species (also arthropod parasites)-but not particularly close in molecular distance (Orthosomella is closer). The other clade includes five strains (JUm1505, JUm1460, JUm1670, JUm2552 and JUm1396) and appears as an outgroup to the four strains + Liebermannia spp. Based on the host phylum, the molecular distances and the monophyletic clade groupings, we describe here two new genera named Enteropsectra n. gen. for the first group of four strains (type JUm408), and Pancytospora n. gen. for the second independent clade (type JUm1505).

In Enteropsectra n. gen., we isolated four strains. Based on genetic distance (Table 3; S3 Table ), spore morphology (Figs 7 and8; S6 Fig; Table 4), and host specificity (Table 5) of JUm408 and JUm2551, we define two species: E. longa JUm408 with large spores (type species of the genus) and E. breve JUm2551 with small spores. We do not assign a species name to the two other strains (JUm1456 and JUm1483) as their molecular relationship depends on the gene (SSU rDNA versus β-tubulin). For example, JUm1483 show small spores, was found infecting Oscheius tipulae and groups with JUm2551 by SSU rDNA, but its β-tubulin sequence is closer to JUm408. We thus prefer to abstain assigning a species name to this strain.

In Pancytospora n. gen., we isolated five strains. Based on genetic distance (Table 3; S3 Table ), host specificity and tissue tropism ( 5), we define two species: Pancytospora philotis n. sp. (type species of the genus) infects Oscheius tipulae intestine, while Pancytospora epiphaga n. sp. was found to infect Caenorhabditis brenneri epidermis and muscles.

Nomenclatural acts. The electronic edition of this article conforms to the requirements of the amended International Code of Zoological Nomenclature, and hence the new names contained herein are available under that Code from the electronic edition of this article. This published work and the nomenclatural acts it contains have been registered in ZooBank, the online registration system for the ICZN. The ZooBank LSIDs (Life Science Identifiers) can be resolved and the associated information viewed through any standard web browser by appending the LSID to the prefix ''http://zoobank.org/" The LSID for this publication is: urn:lsid:zoobank.org:pub:0C31D734-FE13-49F9-8318-ADC6714F316E. The electronic edition of this work was published in a journal with an ISSN. The type strain is ERTm2. Two strains ERTm2 and ERTm6 have been submitted to the American Type Culture Collection (ATCC, https://www.atcc.org) as PRA-371 and PRA-372, respectively. The type host is Caenorhabditis briggsae [START_REF] Dougherty | A new species of the free-living nematode genus Rhabditis of interest in comparative physiology and genetics[END_REF][START_REF] Dougherty | The genera of the subfamily Rhabditinae Micoletzky, 1922 (Nematoda)[END_REF], strain JU1348, which was isolated from a mixed sample of decaying vegetal matter (rotting fruits, leaf litter, soil, bark, flowers). The type locality is Periyar Natural Preserve, Kerala, India. The species was also found in C. briggsae in Germany and Cape Verde, and Caenorhabditis elegans and Caenorhabditis remanei in Europe. The ribosomal DNA sequence, deposited to Genbank under Accession JH604648. The genome of the reference strain has been sequenced [START_REF] Cuomo | Microsporidian genome analysis reveals evolutionary strategies for obligate intracellular growth[END_REF] (accession AERB01000000). The spores are ovoid and measure 2.80 x 0.96 μm (ranges 2.21-3.26 x 0.72-1.30) for the large class and 2.04 x 0.53 μm (ranges 1.36-2.92 x 0.28-0.80) for the small class. Infection is localized to the host intestinal cells. Transmission is horizontal, via the oral-fecal route. The species is named to honor Dr. Frederick Ausubel and his work on innate immunity of C. elegans.

Nematocida major n. sp. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org:act:4D7C3F14-187A-4AD1-A62F-DA79BED716E0

The type strain is JUm2507. The type material is deposited as a live frozen culture of the infected host at ATCC and in the collection of the corresponding author (MAF; http://www. justbio.com/worms/index.php). The type host is Caenorhabditis briggsae [START_REF] Dougherty | A new species of the free-living nematode genus Rhabditis of interest in comparative physiology and genetics[END_REF][START_REF] Dougherty | The genera of the subfamily Rhabditinae Micoletzky, 1922 (Nematoda)[END_REF], strain JU2507, isolated from rotting figs. The type locality is Khao Sok National Park, Thailand. The species was also found in Guadeloupe in C. briggsae and Caenorhabditis tropicalis. The ribosomal DNA sequence, deposited to Genbank under Accession KX360148. The spores are ovoid and measure 3.4 x 1.2 μm (ranges 2.9-3.8 x 0.8-1.6) for the large class and 2.3 x 0.54 μm (ranges 1.8-2.7 x 0.41-0.77) for the small class. Infection is localized to the host intestinal cells. Transmission is horizontal, presumably via the oral-fecal route. The species is named after the large size of its spores.

Nematocida minor n. sp. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org:act:646590B-C-E5A8-4FD5-9026-96B276A4D159

The type strain is JUm1510. The type material is deposited as a live frozen culture of the infected host at ATCC and in the collection of the corresponding author (MAF; http:// www.justbio.com/worms/index.php). The type host is Oscheius tipulae [START_REF] Lam | Morphology and biology of Panagrolaimus tipulae n. sp. (Panagrolaimidae) and Rhabditis (Rhabditella) tipulae n. sp. (Rhabditidae), from leatherjacket larvae, Tipula paludosa (Diptera: Tipulidae)[END_REF], strain JU1510, isolated from compost. The type locality is Hluboka nad Vlatavou near Budweis, Czech Republic. The species was also found in O. tipulae in Armenia. The ribosomal DNA sequence, deposited to Genbank under Accession KX360147. The spores are ovoid and measure 1.9 x 0.83 μm (ranges 1.5-2.2 x 0.5-1.1) for the large class and 1.3 x 0.55 μm (ranges 1.1-1.7 x 0.35-0.73) for the small class. Infection is localized to the host intestinal cells. Transmission is horizontal, presumably via the oral-fecal route. The species is named after the small size of its spores.

Nematocida homosporus n. sp. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org:act: C959C7AD-DC01-4391-8B18-1B5D02F7349B

The type strain is JUm1504. The type material is deposited as a live frozen culture of the infected host at ATCC and in the collection of the corresponding author (MAF; http://www. justbio.com/worms/index.php). The type host is Oscheius tipulae [START_REF] Lam | Morphology and biology of Panagrolaimus tipulae n. sp. (Panagrolaimidae) and Rhabditis (Rhabditella) tipulae n. sp. (Rhabditidae), from leatherjacket larvae, Tipula paludosa (Diptera: Tipulidae)[END_REF], strain JU1504, isolated from a rotting Arum stem. The type locality is Le Blanc (Indre), France. The species was also found in the nematode Rhabditella typhae in Portugal. The ribosomal DNA sequence, deposited to Genbank under Accession KX360153. The spores are ovoid and measure 2.0 x 0.72 μm (ranges 1.7-2.7 x 0.56-0.94). Infection is localized to the host intestinal cells. Transmission is horizontal, presumably via the oral-fecal route. The species is named after the single class of spore size that can be observed in the host.

Nematocida ciargi n. sp. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org:act:77EF241F-463D-443A-819D-C32B1BC49332

The type strain is JUm2895. The type material is deposited as a live frozen culture of the infected host at ATCC and in the collection of the corresponding author (MAF; http://www. justbio.com/worms/index.php). The type host is Procephalobus sp. strain JU2895 (Cephalobina), isolated from rotting citrus fruits. The type locality is Barcelona, Spain. The ribosomal DNA sequence, deposited to Genbank under Accession KX360152. The spores are ovoid and 1.4 x 0.59 μm (ranges 1.5-2.2 x 0.41-0.84). Infection is localized to the host intestinal cells. Transmission is horizontal, presumably via the oral-fecal route. The species is named after its type locality, close to the Centre de Regulacio ´Genòmica in Barcelona, Spain. The type strain is JUm408. The type material is deposited as a live frozen culture of the infected host at ATCC and in the collection of the corresponding author (MAF; http://www. justbio.com/worms/index.php). The type host is Oscheius sp. 3 [START_REF] ´lix | Three biological species related to Rhabditis (Oscheius) pseudodolichura Ko ¨rner in Osche[END_REF], strain JU408, isolated from a soil sample. The type locality is the Botanical garden of Reykjavik, Iceland. The ribosomal DNA sequence, deposited to Genbank under Accession KX360142. The spores have the shape of a long and thin rod, measuring 3.8 x 0.49 μm (ranges 3.1-5.0 x 0.35-0.68). The polar tube makes one turn at the posterior part of the spore; one or two polar tube sections can be seen in transmission electron microscopy when the spore is cut transversally. Infection is observed in the host epidermis and does not affect the intestinal cells. The spores do not seem to be enclosed as groups of spores in a vesicle. Transmission is horizontal, presumably via the oral-fecal route. The species is named after the long size of its spores.

Enteropsectra breve n. sp. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org:act:236607CA-8C44-414D-916C-802C7C67600D

The type strain is JUm2551. The type material is deposited as a live frozen culture of the infected host at ATCC and in the collection of the corresponding author (MAF; http://www. justbio.com/worms/index.php). The type host is Oscheius tipulae [START_REF] Lam | Morphology and biology of Panagrolaimus tipulae n. sp. (Panagrolaimidae) and Rhabditis (Rhabditella) tipulae n. sp. (Rhabditidae), from leatherjacket larvae, Tipula paludosa (Diptera: Tipulidae)[END_REF], strain JU2551, isolated from a rotting apple. The type locality is an apple orchard in Orsay (Essonne), France. The ribosomal DNA sequence, deposited to Genbank under Accession KX360145. The spores are ovoid and measure 1.8 x 0.66 μm (ranges 1.3-2.1 x 0.42-0.90). Infection is observed in the host intestine and does not affect the intestinal cells. Transmission is horizontal, presumably via the oral-fecal route. The species is named after the short size of the spores. The type strain is JUm1505. The type material is deposited as a live frozen culture of the infected host at ATCC and in the collection of the corresponding author (MAF; http://www. justbio.com/worms/index.php). The type host is Oscheius tipulae [START_REF] Lam | Morphology and biology of Panagrolaimus tipulae n. sp. (Panagrolaimidae) and Rhabditis (Rhabditella) tipulae n. sp. (Rhabditidae), from leatherjacket larvae, Tipula paludosa (Diptera: Tipulidae)[END_REF], strain JU1505, isolated from a rotting peach. The type locality is Le Blanc (Indre), France. The species was also found in O. tipulae in other locations in France. The ribosomal DNA sequence is deposited to Genbank under Accession KX360131. The spores have the shape of a long and thin rod, measuring 3.5 x 0.42 μm (ranges 2.4-4.7 x 0.25-0.52). Infection is localized to the host intestinal cells. Transmission is horizontal. The species is named after its specificity to Oscheius tipulae (abbreviation Oti).

Pancytospora epiphaga n. sp. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org: act:41573063-B494-41C3-B0A8-2417DFEB3DCC

The type strain is JUm1396. The type material is deposited as a live frozen culture of the infected host at ATCC and in the collection of the corresponding author (MAF; http://www. justbio.com/worms/index.php). The type host is Caenorhabditis brenneri. The type locality is a private garden in the vicinity of Medellin, Colombia. The ribosomal DNA sequence, deposited to Genbank under Accession KX424959. The spores are ovoid and measure 3.71 x 0.80 μm (ranges 2.99-4.22 x 0.56-0.92). Infection is observed in the host epidermis and does not affect the intestinal cells. Transmission is horizontal. The species is named after the site of infection in the nematode, mostly the epidermis.

Clément Dubois did his M2 internship in our lab. His work was closely related to this part of my Ph. D. project. I will talk about his results in the Discussion.

II.1. Background

Specificity of infection of Caenorhabditis and Oscheius species by eight microsporidia species was described in the former chapter. Meanwhile, intraspecific variation of C. elegans sensitivity to N. parisii and N. ausubeli was also observed during the maintenance of wild infected strains. Some infected C. elegans strains displayed higher mobility, higher fecundity and greater longevity than other C. elegans infected with the same microsporidia. This host phenotypic variation can be host-based or pathogen-based, or dependent on their interaction.

Host C. elegans strains may have different genetic bases underpinning their microsporidiadefense traits. On the other hand, intraspecific variation for pathogenicity may also exist in microsporidia. Here, we focus on the host C. elegans. We were particularly interested in the genetic basis of C. elegans sensitivity variation to microsporidia infection, which can be further studied by Quantitative trait locus (QTL) analysis.

Previously, the sensitivity of six C. elegans strains to N. parisii infection was investigated by Balla et al. (2015). They notably found that one C. elegans strain, from Hawaii (HW), has enhanced immunity at the young larval stage (L1), which gives to this strain the ability to clear the N. parisii infections. This phenomenon was not observed in the other five strains tested, including N2. The genetic basis of this early-life immunity variation between HW and N2 was further investigated. The results suggested that the special immune ability of the Hawaiian strain was a complex multigenic trait (Balla et al. 2015). Among the six C. elegans strains compared in this study, only one strain ERT002 (derived from CPA24) was naturally found infected with N. parisii (ERTm1). The other five strains may have never encountered any microsporidia in their natural habitats. To study the evolutionary consequences of microsporidia infection pressures on C. elegans, it is relevant to compare C. elegans strains that have been exposed to microsporidia in the wild.

The work described in this chapter is the first description of variation among naturally infected C. elegans strains in sensitivity to microsporidia infection. While comparing the sensitivity of ten wild C. elegans strains to N. ausubeli infection, we proved the natural variation for this trait in C. elegans species. Interestingly, we found contrasted levels of C. elegans tolerance to N. ausubeli infections. Indeed, the two C. elegans strains, JU1249 and JU2825, showed similar levels of infection but highly contrasted levels of sensitivity (indirect fitness assay). These two C. elegans strains will be further used to identify genetic locus/loci involved in C. elegans tolerance to N. ausubeli microsporidia.

II.2. Materials and Methods

II.2.1. C. elegans and Microsporidia strains

In RESULTS I, 19 C. elegans strains had been isolated with N. parisii or N. ausubeli infection. Among these C. elegans isolates, the strains JU1249, JU1762, JU2009, JU2132, JU2287, JU2526, JU2586, JU2816, JU2825 and the lab reference strain N2 were used. The nine naturally infected strains were subsequently bleached to remove microsporidia infection and any other microbes, and maintained in the lab as described in RESULTS I.

The N. ausubeli strain used in the first infection assay on all the ten C. elegans strains was JUm2009, a N. ausubeli strain isolated from the C. elegans strain JU2009. Another N. ausubeli strain JUm2526, two N. parisii strains JUm1249 and JUm2816 were isolated from the C. elegans strains JU2526, JU1249 and JU2816, respectively. JUm2009, JUm2526 and JUm1249 were used on the further infection assays on JU1249 and JU2825, to compare their sensitivity variation upon different microsporidia infections. Finally, N. ausubeli JUm2009 and N. parisii JUm2816 were used to infect the mixed population of JU1249 and JU2825 in the competition assay. Spores were prepared and quantified as described in RESULTS I.

II.2.2. Food consumption assays

A. Infection and maintenance 20 uninfected young adults (i.e. prior to first egg formation) were transferred onto a 55mmdiameter NGM plate seeded with E. coli OP50. 2 million of microsporidian spores suspended in 100 µl of distilled water were dispensed onto the E. coli lawn. The cultures were then incubated at 23 °C. The infection symptoms were monitored by Nomarski optics at 48 hours after inoculation. In order to avoid the possible bias caused by the high or low concentration of spores used in the initial inoculation, and to have a stable infected C. elegans population, 20 young adults were transferred to a new plate every two days. Uninfected C. elegans populations were also maintained and transferred for a few generations. Three replicates were conducted in parallel for each experiment. Infected and uninfected C. elegans populations were maintained for ten days after inoculations. Then the C. elegans isolates were used in food consumption assays (the protocols of the food consumption assay and the intensity normalization method were from by Matthew Rockman and Victoria Cattani).

B. Plate seeding, C. elegans transfer and fluorescence scan

To study the population growth of different C. elegans strains under different conditions, food depletion by each strain was measured in the presence or absence of microsporidia infection.

First, 100 µL of OP50-GFP (a E. coli OP50 strain carrying a GFP marker) were carefully seeded in the center of 6 cm NGM plates. Control 6 cm NGM plates were seeded with 100 µL of regular OP50, which were required for the normalization of OP50-GFP fluorescence intensity. Seeded plates were incubated at room temperature for 24 hours in the dark. Signal intensity of the seeded plates was then scanned and measured using a Typhoon FLA 9000 laser scanner (GE Healthcare Life Sciences). The fluorescence intensity in this first scan was used as the data at timepoint 1. C. elegans individuals from prepared populations were transferred to plates seeded with OP50-GFP soon after the first scan.

From the C. elegans populations maintained for ten days, 10 L4s were transferred to a plate seeded with OP50-GFP, with three replicates per experiment. The fluorescence decay in plates was scanned every 24 hours until the depletion of OP50-GFP. Two types of control were also scanned at each timepoint: three plates seeded with OP50-GFP but without C. elegans (a control for bacterial growth), and three plates seeded with regular OP50 but without C. elegans (a negative control for non-specific fluorescence).

A few plates got contaminated by fungi or other bacteria after several times of scan. These contaminated plates were discarded then.

C. Fluorescence intensity normalization

The mean fluorescence intensity of each plate was corrected by the formula below to obtain the corrected fluorescence intensity (F c ). The food fluorescence intensity on each plate at different timepoints was then analyzed and compared. As the bacteria OP50 could grow in the plate during the experiment, the GFP intensity could initially rise when the bacteria growth rate was higher than the consumption rate of C. elegans.

In the food consumption assays focusing on JU1249 and JU2825, infected C. elegans populations were harvested after the last scan, and stored in acetone at -20 °C for microsporidia infection measurements by FISH.

F c = F i -F backi F 1 -F back1 F nowormi -F backi F noworm1 -F back1
each transfer, animals from old plates were harvested in M9 solution and stored at -80 °C for quantification of the respective proportions of JU1249 and JU2825.

B. Allele quantification

The whole genomes of JU1249 and JU2825 were sequenced using Illumina Hiseq4000 with 20X coverage and paired end 150 bp. For each genome, the raw data were aligned to the reference genome (C. elegans WS243) using a combination of softwares -bwa, samtools, Picard and Genome Analysis Toolkit (GATK) (Li and Durbin, 2009;Li et al., 2009;[START_REF] Mckenna | The Genome Analysis Toolkit: a MapReduce framework for analyzing next-generation DNA sequencing data[END_REF]. From the output files, homozygous single-nucleotide polymorphisms (SNPs) versus the reference genome were called (A pipeline on the genome data analysis was developed in the lab by Fabrice Besnard). A few SNPs between JU1249 and JU2825 were selected manually.

Based on one of the single-nucleotide polymorphisms (SNPs) between JU1249 and JU2825 on chromosome III, pyrosequencing primers III_663310_Forward (5'-GTGACGTACTAGCAACGAGTCGATTTTGGGGATGGA-3'), III_663310_Reverse (5'-CACTAGGCAGGTAGGCATTTTT-3') and III_663310_Sequencing (5'-CCAAACTTTTATAGAGATCA-3') were designed. Allele quantification was performed using pyrosequencing as previously described [START_REF] Félix | Population dynamics and habitat sharing of natural populations of Caenorhabditis elegans and C. briggsae[END_REF]. In brief, harvested C. elegans isolates were centrifuged at 3,000 rpm for 2 minutes to generate a nematode pellet. Lysates of 2 µL of the pellet were used as PCR templates, with the primers III_663310_Forward and III_663310_Reverse. The allele frequency of the PCR product was quantified with the pyrosequencer (PyroMark Q96 ID instrument; Biotage), using primer III_663310_Sequencing and a universal biotinylated primer TAGCAGGATACGACTATC-3').

The accuracy of this quantification method was estimated by measuring the allele frequencies of PCR products that were amplified using C. elegans lysates of known proportions of mixtures of JU1249 and JU2825 individual L4s. On average, 2% of discrepancy existed between expected and observed allele frequencies.

The experiment was continued until all replicates in one treatment reached fixation of either JU1249 or JU2825.

II.3. Results

II.3.1. Natural variation of C. elegans population growth, in the presence or absence of N. ausubeli infection

In the Felix laboratory collection of wild nematode isolates, 19 C. elegans strains were naturally infected with microsporidia (Figure 24). The N. parisii infection has been shown to The strong assumption of our food consumption assay is that a faster worm population growth causes faster food depletion, i.e. there is a good correlation between fitness and food depletion. Note that a faster food depletion rate is not necessarily fully correlated to the nematode population growth rate. For example, it might also be correlated to the food consumption rate per worm. A small population with a high food consumption rate per worm may consume food faster than a relatively larger population with a lower food consumption rate per worm. However, when considering a single C. elegans strain, the impact of microsporidian infection on the host fitness will be reflected by the decay in food consumption caused by the infection. C. elegans food consumption rates might be differentially influenced by microsporidian infections, which could indicate variations in the sensitivity of C. elegans to microsporidia infection. Although the food consumption assay is a rough proxy, it is a practical and convenient method to quantify C. elegans sensitivity to microsporidian infection.

First, we investigated variation in sensitivity among ten C. elegans strains, nine of which were naturally found infected with N. parisii or N. ausubeli, and N2 as a control (Figure 24, marked by "*"). We bleached each strain and infected it with the N. ausubeli strain JUm2009. Then these infected populations were maintained for 10 days (about four generations) to a relatively stable state, in which infected C. elegans individuals infect other uninfected individuals and C. elegans populations continue the infection over generations. From the stable infected population, individual L4s were transferred to plates seeded with E. coli OP50-GFP, the fluorescence intensity of which was monitored at different timepoints. Uninfected C. elegans isolates were also maintained and tested in parallel.

As expected, we find that for nine C. elegans strains (JU2526 will be described in the next chapter), the infected populations consumed food slower than the uninfected populations (Figure 25). This demonstrated that N. ausubeli infection negatively affects C. elegans food consumption rates. Moreover, this negative impact of N. ausubeli infections varies between strains (Figure 25 and Figure 26). For example, with or without infection, JU2287 is the slowest food consumer among the nine strains. However, JU2825, the fastest food consumer when uninfected, only had a medium food consumption rate among the nine strains when they were infected (Figure 26). This demonstrates that the food consumption rates of different C. elegans strains were affected differently when they were infected with N. ausubeli, which suggested that the natural variation in C. elegans response to the N. ausubeli infection.

To further understand the genetic basis of C. elegans sensitivity variation to microsporidia infection using a quantitative trait locus (QTL) approach, at least two strains showing contrasted phenotypes are required. Among the C. elegans strains we tested, JU1249 and JU2825 were chosen as candidates. In the absence of microsporidia infection, JU2825 consumed food faster than JU1249. In contrast, with N. ausubeli JUm2009 infection, JU1249 consumed food faster than JU2825 (Figure 25, Figure 26). Thus, N. ausubeli JUm2009 infection has a larger negative influence on JU2825 than on JU1249. JU1249 is more resistant to N. ausubeli JUm2009 than JU2825. We further investigated whether the host sensitivity variation between JU1249 and JU2825 is specific to N. ausubeli JUm2009. E. coli OP50-GFP intensity over time for nine C. elegans strains with or without N. ausubeli JUm2009 infection. The Y-axis is the normalized intensity of the fluorescence of OP50-GFP. The X-axis is the scanning time point. Each plate was scanned every 24 hours until full depletion of bacteria on the plate. Data are mean values of three replicates (data marked with "*" are mean values of two replicates), with bars represent standard errors. As the bacteria OP50-GFP could grow on the plate, the intensity increased in the beginning. To further understand why JU1249 consumed food faster than JU2825 with different microsporidia infections, we examined the infection rates and the pathogen loads, using a FISH probe targeting N. parisii and N. ausubeli. We performed this quantitative FISH on the populations used in the Food consumption test after the last scan at 168 hours. When infected with the N. parisii JUm1249, JU1249 displayed a significantly higher pathogen load than JU2825 (p-value is 4e-05). With either of the two N. ausubeli infections, no significant difference in the pathogen load was observed between JU1249 and JU2825. When they were infected with any of the three microsporidia, no significant difference in the infection rate was detected between JU1249 and JU2825. These results suggest that JU1249 and JU2825 have different tolerance abilities. Indeed, JU1249 shows a higher food consumption rate than JU2825 with N. ausubeli infection, yet it did not show a lower infection rate or a lower pathogen load. Together, FISH tests indicate that a different tolerance of JU1249 and JU2825 affected their performances in the food consumption test. C. elegans strains JU1249 and JU2825 were infected with N. parisii JUm1249, N. ausubeli JUm2009 and N. ausubeli JUm2526, respectively. Data are mean values of three replicates, with the standard errors indicated as the error bars. In each replicate, 30 adult C. elegans were measured. A. Pathogen loads of JU1249 and JU2825 upon different microsporidia infections. A t-test was performed to test whether the pathogen loads of the two C. elegans strains significantly differed upon each microsporidia infection. Only upon N. parisii JUm1249 infection, JU1249 displayed a significantly higher pathogen load than JU2825 (p-value is 4e-05, indicated by "**"). No other significant difference between JU1249 and JU2825 was found. B. Infection rate of JU1249 and JU2825 upon different microsporidia infections. A Wilcoxon Rank Sum test was performed to test whether the infection rates of the two C. elegans strains significantly differed upon each microsporidia infection. No significant difference between JU1249 and JU2825 was found.

II.3.3. JU1249 outcompeted JU2825 when they were infected with microsporidia in laboratory competition assays

Competition assays examine the relative fitness between different co-occurring strains. We performed competition assays between JU1249 and JU2825 populations in the presence and absence of microsporidia infection, to compare their sensitivity to microsporidia infection, and to examine the possible evolutionary consequences of this trait in C. elegans and for QTL mapping eventually. In the assay, we bleached and synchronized JU1249 and JU2825, then we mixed the two strains in equal number of individual L4s to seed the populations. C. elegans populations with N. ausubeli infection, N. parisii infection and no infection were assessed, each with five replicate cultures. The populations were maintained by being transferred to fresh plates before starvation. After each transfer, the proportions of JU1249 and JU2825 in the old plates were measured using quantitative pyrosequencing.

Starting from 50% for both strains, in the presence of either microsporidia infection, the resistant strain JU1249 outcompeted the sensitive strain JU2825 (Figure 29). N. ausubeli JUm2009 infection had a stronger selective effect on C. elegans population than N. parisii JUm2816 infection did. Populations infected with JUm2009 reached to fixation of JU1249 at the sixth transfer, while the populations infected with JUm2816 reached to fixation of JU1249 at the eighth transfer. On the other hand, JU2825 outcompeted JU1249 in the uninfected condition. These results confirm that the C. elegans strain JU2825 is more sensitive to microsporidia infection than JU1249. The sensitivity variation among different C. elegans strains to microsporidia infection may alter the outcomes of intraspecific competition in the wild. The proportion of JU2825 and JU1249 is inferred from the proportion by pyrosequencing of their respective private SNP. The mean proportions of five replicates of each treatment at each transfer were indicated. Error bars are standard errors over replicates. The experiment was continued until all replicates in one treatment reached fixation of either JU1249 or JU2825. In the infected populations, JU1249 outcompeted JU2825, whereas in the absence of microsporidia, JU2825 outcompeted JU1249.

II.4. Conclusion

Overall, we find natural diversity for C. elegans sensitivity to N. ausubeli or N. parisii infections. The C. elegans strains JU1249 and JU2825 can be further used to start a QTL analysis on the genetic bases of C. elegans sensitivity variation to microsporidian infection. This experiment was performed in parallel to those in Figure 25.

We further investigated the microsporidia effect on four C. elegans strains with the Mrt phenotype. These strains were tested for the number of generations before becoming completely sterile, in the presence and absence of microsporidia infection. The results demonstrate that both N. parisii and N. ausubeli infections could increase the number of generations of JU2526 and JU775 at high temperature.

III.2. Materials and Methods

III.2.1. C. elegans and Microsporidia strains

The C. elegans strains JU2526, MY10, QX1211 and JU775 were used in this study. All of these four strains have been shown to display the Mrt phenotype. As the Mrt phenotype is temperature sensitive, these four strains were bleached and maintained at 15 °C. The N. parisii strain JUm2816 and the N. ausubeli strain JUm2009 were used in this study. Spores were prepared and quantified as described in RESULTS I.

III.2.2. Mortal germline assay

20 uninfected young adults (i.e. prior to first egg formation) were transferred to a 6 cm NGM plate seeded with E. coli OP50. 5 million microsporidian spores in 100 µl distilled water were placed onto the E. coli lawn. The cultures were then incubated at 15 °C. The infection symptoms were checked by Nomarski optics at 72 hours after inoculation. 10 L4 larvae from the infected population were transferred to a fresh plate and moved to 23 °C. Uninfected individual L4s were also tested in parallel. The generation moved from 15 °C to 23 °C was called the first generation. Ten L4 animals were transferred onto fresh plates at each generation (2-3 days). The transfer was continued until the sterile generation. i.e. when 10 L4 progenies were not available on the plate. The number of generations before becoming sterile was scored.

Three experiments were performed, each with three treatments: N. parisii JUm2816 infection, N. ausubeli JUm2009 infection and uninfected control. In the first experiment, we tested C. elegans strains JU2526 and MY10, both with three replicates; in the second experiment, we tested JU2526 again with five replicates; in the third experiment, we tested C. elegans strains JU775 and QX1211, both with five replicates.

The statistical analysis and the figures in Figure 25 were obtained using Rstudio, as described in Results II. To test whether the number of generations before becoming sterile of each strain with different treatments significantly differed, we used a Wilcoxon Rank Sum test on the generation numbers of different treatments within each strain.

III.3. Results

We tested four C. elegans strains JU2526, MY10, QX1211 and JU775. MY10, JU775 and QX1211 have been shown to have an extremely strong Mrt phenotype at 25 °C (Marie-Anne Félix's data, not shown). In the experimental framework used in my study, at 23 °C, MY10 and QX1211 become sterile in 3 to 4 generations. JU775 and JU2526 exhibit a milder Mrt phenotype. In average, JU775 becomes sterile in about 7 generations, and JU2526 in about 9 generations (Figure 31). With either N. parisii JUm2816 infection or N. ausubeli JUm2009 infection, JU2526 had a significant increase (p-value is 0.01 for both) in the number of generations before the sterility (Figure 31B). Compared to those with N. parisii JUm2816 infection, JU2526 with N. ausubeli JUm2009 infection also showed a significant increase (p-value is 0.04) in the generation number. With either microsporidia infection, JU775 had a significant increase (p-value is 0.01 for both) in the number of generations before the sterility (Figure 31C). Microsporidia infections did not seem to rescue the progressive loss of fertility of MY10 or QX1211. These results indicated that microsporidia infections are able to rescue fertility in C. elegans strains with a moderate Mrt phenotype.

elegans.

Data are mean values and standard errors. In each treatment, A. Three replicates for JU2526 and for MY10; B. Five replicates for JU2526; C. Five replicates for JU775 and QX1211. A Wilcoxon Rank Sum test was performed to test whether the number of generations before becoming sterile of each strain with different treatments significantly differed. All the statistical differences were shown, with * for p-value < 0.05. In A, the differences in generations of JU2526 with or without infection are not significant, which may be due to the limited number of replicates.

As we have already discussed the results in the paper of RESULT I, here I will include again the key points in the discussion of the paper and make further comments on our other results, related studies and some recently reported discoveries.

I. Diverse interactions between rhabditids and microsporidia I.1. The diversity of microsporidia infections in rhabditids

A great diversity of interactions between microsporidia and rhabditid nematodes have been discovered. In addition to N. parisii and N. ausubeli, we found five new Nematocida species, two Enteropsectra species and two Pancytospora species; another two new Nematocida species, Nematocida displodere and Nematocida ironsii, were proposed very recently (Troemel et al., 2008, Luallen et al., 2016;[START_REF] Reinke | Identification of microsporidia host-exposed proteins reveals a repertoire of rapidly evolving proteins[END_REF]. These 13 microsporidia were isolated from 48 rhabditid nematode strains of six Caenorhabditis species, two Oscheius species, Rhabditella typhae and Procephalobus sp. (Troemel et al., 2008, Luallen et al., 2016;[START_REF] Reinke | Identification of microsporidia host-exposed proteins reveals a repertoire of rapidly evolving proteins[END_REF]. In addition, two undefined microsporidia species in the genus Enteropsectra and some environmental SSU rDNA microsporidia sequences may represent further species in the genera Nematocida and Pancytospora (paper Figure S2, S4) (Ardila-Garcia et al., 2013). Altogether, these discoveries demonstrate the great diversity of microsporidia infections in wild rhabditid nematodes.

I.2. How were these microsporidia species described?

As it is difficult to do mating test among microsporidia, these nematode-infecting microsporidia species were mainly defined by molecular methods. N. parisii was defined to be a new species in a new genus of microsporidia because its SSU rDNA sequence was found to be fairly divergent but most closely related to the SSU rDNA of microsporidia Ovavesicula popilliae. N. ausubeli was defined to be an independent Nematocida species because its SSU rDNA is approximately 95% identical to that of N. parisii, which is its most closely related species in the phylogenetic analysis (Troemel et al., 2008).

In our study, the new species N. major, N. minor, N. homosporus and N. ciargi were characterized by SSU rDNA and ß-tubulin. These four Nematocida species all had N. parisii or N. ausubeli as their top blast hit in GenBank, with similarity between 81%-86% of SSU rDNA and 74%-84% of ß-tubulin genes. They formed a single clade with N. parisii and N. ausubeli, with O. popilliae as sister group within Clade II of the microsporidia SSU rDNA phylogeny. The inter-specific genetic distance of SSU rDNA among them is at least 0.06, which is greater than the distance between N. parisii and N. ausubeli. But their intra-specific distances are all 0.00. Therefore, these four Nematocida were defined as new Nematocida species. N. sp. 7 was defined as a new species based on comparison and the phylogeny of ßtubulin genes with other species. E. longa, E. breve, P. philotis and P. epiphaga were described as new species because their SSU rDNA were closest (88-89% identity) to that of Orthosomella operophterae, which is in the Clade IV of the microsporidia phylogeny. The ß-tubulin sequences of the Enteropsectra species were closest (75-76% identity) to Vittaforma corneae, which is a close relative to O.

operophterae. E. longa, E. breve, P. philotis and P. epiphaga were placed in two close groups, with O. operophterae as outgroup in the Clade IV of the microsporidia SSU rDNA phylogeny. Among the four strains in the Enteropsectra group, we defined E. longa (JUm408) and E. breve (JUm2551) as two species, with further considerations on their spore morphology and host specificity. We did not define Enteropsectra JUm1456 and Enteropsectra JUm1483 to a species due to their different placements in the SSU rDNA phylogeny and the ß-tubulin phylogeny. The other group, the Pancytospora group contains five strains, which were defined as two species based on SSU rDNA, host and tissue specificity.

N. displodere and N. ironsii were defined as new Nematocida species based on whole genome sequencing and phylogenomic comparison. N. displodere was found to be a sister group to N. parisii and N. ausubeli in their phylogenomic tree. However, N. parisii and N. ausubeli proteins showed an average amino acid identity of 66% compared to each other, and 48.6% and 48.3% compared to N. displodere proteins, respectively. Thus, N. displodere was defined as a new Nematocida species (Luallen et al., 2016).

The phylogenomic tree includes six strains of four Nematocida species and 17 other microsporidia species. N. displodere is in the basal position of the Nematocida genus. N. ironsii is the closet species to N. parisii. Adapted from [START_REF] Reinke | Identification of microsporidia host-exposed proteins reveals a repertoire of rapidly evolving proteins[END_REF].

Interestingly, our phylogenetic analysis based on the ß-tubulin gene also showed the same groupings of ERTm1, ERTm3 versus ERTm5, as their genomic phylogeny displayed (Figures 32,33B). In the ß-tubulin phylogeny, the N. parisii clade includes two subclades: one subclade includes ERTm1, ERTm3 and six other strains, while the other subclade includes ERTm5 and five other strains (Figure 33B). Thus, the other strains in the same subclade as ERTm5 may also belong to N. ironsii. It will be necessary to further perform genomic and phylogenomic analyses on more strains to understand better the relationships and diversity among different "N. parisii" strains, as well as among N. ausubeli and other nematodeinfecting microsporidia. ausubeli strains. However, the phylogeny of SSU rDNA does not show any of the "parisii" subclades in B. Note that the ß-tubulin gene is more divergent than SSU rDNA, which may explain the better resolution of subclades in B. Branch colors show the posterior probability, with the corresponding color code shown on the left. Scale bar indicates expected changes per site.

I.3. Various tissue tropisms of rhabditid-infecting microsporidia

Only horizontal transmission has been observed in the known rhabditid-infecting microsporidia infections, likely via the oral route. Different nematode tissues are infected by the various microsporidia. In the 13 known rhabditid-infecting microsporidia, 11 of them infect the intestinal cells of their nematode hosts, whereas the other two, N. displodere and P. epiphaga, showed infections in other tissues. N. displodere infection has been found in the epidermis, muscles, coelomocytes and neurons of C. elegans. P. epiphaga infection has been found in the epidermis, muscles and coelomocytes of C. brenneri and at least the epidermis of C. elegans. The various infection niches show that rhabditid-infecting microsporidia have evolved to use different host tissues to maximize their growth.

N. displodere seemed to have some other special features that are related with its diverse tissue tropisms. For example, with a longer polar tube than that of N. parisii (three-fold longer than similar sized spores of the two species), N. displodere was suggested to be able to inject the sporoplasms directly into host epidermis and muscles from the intestinal lumen. Unlike the spores of N. parisii that are continuously shed through the host defecation process, N. displodere spores were observed to accumulate inside the host, mainly in epidermis and muscles, until they are released into the environment through bursting of the host vulva (Luallen et al., 2016). The entry and release mechanisms of P. epiphaga remain unknown yet.

It would be interesting to further study the infection mechanisms of N. displodere and P. epiphaga, and compare them with each other, as well as with the intestine-infecting microsporidia.

I.4. Spread of microsporidia within the infected tissue

A cell-to-cell spread mechanism of Nematocida was reported recently (Balla et al., 2016).

Previously, it was showed that only infected C. elegans with N. parisii spores inside their intestinal cells are infectious, whereas those with only meronts are not infectious to other C. elegans individuals (Troemel et al., 2008). However, within one worm, a single inoculated intestinal cell that only contains N. parisii meronts is found to be contagious to its neighboring intestinal cells (Balla et al., 2016). This cell-to-cell transmission of N. parisii could proceed throughout up to half of all the intestinal cells without any differentiation of spores, and lead to deformation of cell boundaries and syncytia formation in the worm's intestine. Similar to N. parisii, N. ausubeli is able to spread across several intestinal cells before forming spores. N. displodere could spread across different host muscle and hypodermal cells, and even from the main hypodermal syncytium into the anterior epidermal cells. This cell-to-cell spread mechanism was thus suggested to be a conserved mechanism in the genus Nematocida (Balla et al., 2016).

To better determine whether this cell-to-cell spread mechanism is conserved in the genus Nematocida, it is necessary to further examine some or all of the five new Nematocida species we have found. It would be also interesting to examine if similar mechanisms exist in Enteropsectra and Pancytospora species, using the single cell inoculation method in Balla et al. (2016).

I.5. Microsporidia exit mechanism from host intestinal cells

To exit from the intestinal cells of C. elegans, N. parisii first rearranges the host actin (ACT-5) to induce gaps in the terminal web of the host intestinal cells (mentioned in the Introduction III.1.2.) (Troemel et al., 2008;Estes et al., 2011). Then the spore-containing compartment (SCC) will be marked with RAB-11, and then traffic to and fuse with the apical membrane of the intestinal cells. After the fusion, spores gain access to the intestinal lumen and are marked by PGP-1 (Figure 22A, Figure 34, left panel (a)) (Szumowski et al., 2014). A very recent study showed an alternative mechanism that spores may have ACT-5 coats in addition to the PGP-1 marker, when they gain the access to the lumen (Figure 34, left panel (b)) (Szumowski et al., 2016). The ACT-5 coat is not necessary for spore exit, but it may promote spore exocytosis. Moreover, later during infection, spore-filled vesicles coated with CHC-1 (clathrin heavy chain), PGP-1 and ACT-5 were noticed, which was suggested to act as a compensatory endocytosis mechanism for the host cell, due to the massive efflux of intracellular membrane material (Figure 34, right panel) (Szumowski et al., 2016).

In our study, transmission electronic microscopy indicated different exit mechanisms between

Nematocida and Enteropsectra species. N. ausubeli probably has a similar exit mode as N. parisii: some isolated N. ausubeli spores possessed an additional membrane, which likely formed the SCC, and exited from host cells through exocytosis; no additional membranes have been observed around N. ausubeli spores in the intestinal lumen (paper Figures 5H,K, L, 6A). However, E. longa showed a distinct cellular exit mode: no SCCs have been observed homosporus, N. minor has two sizes of spores, which is a feature shared by N. parisii, N. ausubeli and N. major. Our infection studies showed that when O. tipulae or O. sp. 3 were infected with N. ausubeli, which was not known as their natural pathogen, two sizes of spores were observed. It would be interesting to examine the appearance of N. minor spores in a Caenorhabditis host, if the infection can be established.

N. ciargi, which was placed on the basal position in the phylogeny (based on SSU rDNA and ß-tubulin, paper Figure 3) of Nematocida, was isolated from a Procephalobus strain.

Procephalobus sp. is an outgroup nematode, compared to the other nematode host species described here. The study on host specificity of N. ciargi may provide insights on how the host specificity of different Nematocida species evolves.

The two nematode-infecting microsporidia with a non-intestinal tropism, N. displodere and P. epiphaga, were isolated from C. elegans and C. brenneri, respectively. P. epiphaga could also infect C. elegans and C. briggsae. It is of interest to study the specificity of the two species, for example, to see whether the microsporidia with broader tropisms also have broader host spectra.

On the other hand, many other rhabditid nematode species have not been found with microsporidia infection in the wild or tested for their susceptibility to the known microsporidia in the laboratory. For example, in Caenorhabditis, only five species of the Elegans group and C. sp. 42 of the Drosophilae group were involved here. It would be interesting to test microsporidia infection in other Caenorhabditis species. Furthermore, it would be also interesting to test the susceptibility of non-rhabditid nematodes to microsporidia infections. For example, the diplogastrids Pristionchus spp., potential predators of C. elegans and other rhabditids, may also be susceptible to some of the known rhabditidinfecting microsporidia.

I.7. Diversification of the nematode-infecting microsporidia

Diversification of rhabditid-infecting microsporidia may take place in three routes. Firstly, diversification and speciation of hosts and pathogens may occur in parallel. The correspondent positions of Nematocida and host nematodes (Caenorhabditis and Oscheius) on their phylogenies indicates that microsporidia may coevolve with nematodes, and speciation of microsporidia may occur after the divergence of rhabditid species (paper Figure 3). The genetically close Enteropsectra species and their narrow specificities to Oscheius hosts may be also due to the fact that speciation of microsporidia occurred after the divergence of O. tipulae and Oscheius sp. 3.

Secondly, as rhabditid nematodes share their natural habitats and interact with diverse animals, such as arthropods, mollusks and other nematodes, it is possible that host switch take place among them. For example, the millipede Diploiulus londinensis caeruleocinctus has been found to be infected with Nosema juli, the genus of which is in the Clade IV of the microsporidia phylum [START_REF] Wilson | Nosema juli n. sp., a microsporidian parasite in the millipede Diploiulus londinensis caeruleocinctus (Wood) (Diplopoda: Julidae)[END_REF]. Snails have been found to be infected with microsporidia, some of which are in the Clade IV of the microsporidia phylum [START_REF] Cunningham | Extinction of a species of land snail due to infection with a microsporidian parasite[END_REF]McClumont et al., 2005). It is possible that microsporidia can switch hosts from millipedes or snails to rhabditids. Another possibility is that microsporidia may shift host among different rhabditids, as pathogens might have a higher chance to establish infection in a new host that is closely related to the original host. After the host shifts, pathogen speciation and host specificity may or may not take place to form new microsporidia with narrow ranges of hosts, or with broad ranges of hosts.

Thirdly, both the first and the second diversification routes may have taken place simutaneously to form the diversity of rhabditid-infecting microsporidia.

The diverse natural nematode-microsporidia pairs, especially those with C. elegans, provide many opportunities for studies on the evolution of host shifts and host specificity of pathogens. elegans strains (geographically diverse) to N. ironsii ERTm5 infection was investigated by Balla et al. (2015), with the comparison of host survival and pathogen loads; 2), in our study, nine C. elegans strains were compared for their sensitivity variations to a same N. ausubeli infection (RESULT II). All these nine tested strains were naturally found with N. parisii or N. ausubeli infection (Figure 24); 3), in a related study to RESULT II, Clément Dubois further compared nine C. elegans strains: JU1249 and JU2825 that were tested in RESULT II, the Hawaii strain CB4856 with an enhanced immunity in its L1 larvae (Balla et al. 2015), the other six strains representing genetic and/or geographic diversity. The sensitivity of these nine strains to the infections N. ausubeli JUm2009 and N. parisii JUm2816 were compared.

I.8. An excellent system to study host-pathogen interactions and coevolution

The studies 2) and 3) used the OP50-GFP food consumption tests to compare the host sensitivity upon microsporidia infection. All the three studies found variation in C. elegans sensitivity to microsporidia infection (Figure 23A and B,[START_REF] Sapir | Microsporidia-nematode associations in methane seeps reveal basal fungal parasitism in the deep sea[END_REF][START_REF] Stiernagle | Maintenance of C. elegans[END_REF].

II.2. Interesting pairs of C. elegans strains for further studies

The enhanced immune ability to control and clear N. parisii infection in L1 larvae of CB4856 has been studied with the reference strain N2, which lacks this special immunity (Balla et al. 2015). In our studies, we found that the strains JU1249 and JU2825 showed different sensitivity against both N. parisii and N. ausubeli infections (Figure 27). C. elegans JU1249 is more resistant than JU2825 to N. ausubeli JUm2009, JUm2526 and N. parisii JUm1249 infections. The different sensitivity of JU1249 and JU2825 was reconfirmed in the study by Dubois (Figure 35). Among the nine strains compared by Dubois, JU1249 is the most resistant strain, as the populations of JU1249 consumed the food the fastest upon either of the two microsporidia infections. JU2825, JU2914 and JU751 consumed the food slower compared to other C. elegans strains. These results demonstrated that JU1249 and JU2825 have contrasted phenotypes in the sensitivity to microsporidia. These two C. elegans can be further used in the quantitative trait locus (QTL) approach for detecting the genetic basis of this phenotype variation.

In the study by Dubois, another pair of C. elegans strains, CB4856 and ED3077, showed contrasted variation patterns in sensitivity to different microsporidia infection. With N. parisii JUm2816 infection, ED3077 consumed food faster than CB4856, whereas the variation pattern is reversed when the two strains were with N. ausubeli JUm2009 infection (Figure 35, [START_REF] Szumowski | Small GTPases promote actin coat formation on microsporidian pathogens traversing the apical membrane of Caenorhabditis elegans intestinal cells[END_REF]. ED3077 seems to be more resistant to N. parisii JUm2816 than to N. ausubeli JUm2009, whereas CB4856 appears to be the opposite. These two C. elegans strains provide a good pair to study the interactions between C. elegans and microsporidia.

II.3. The different tolerance of JU1249 and JU2825 to microsporidia infection

Using FISH analyses, we found that though JU1249 consumed food faster than JU2825, populations of JU1249 did not have a lower infection rate nor a lower pathogen load on average than JU2825 populations (Figure 28). In fact, upon N. parisii JUm1249 infection, JU1249 had a significantly higher pathogen load than JU2825. These results indicate a different tolerance of JU1249 and JU2825 to microsporidia infection. By contrast, in the study by Balla et al. (2015), the different mortality rates of the six C. elegans strains are positively correlated with the differences in pathogen loads. This result indicates variation in host resistance rather than tolerance, which was consistent with their further findings that L1 larvae of C. elegans CB4856 displayed a special clearing ability to microsporidia infection. Further studies on brood size of JU1249 and JU2825 with or without microsporidia infection were performed by Dubois (Figure 37). Three N. parisii strains and three N. ausubeli strains were used separately to infect JU1249 and JU2825 in this study. First, the infections of the six strains of two Nematocida species greatly reduced the brood size of both JU1249 and JU2825, which indicates that neither of the two C. elegans strains can clear microsporidia infection like CB4856. Second, the brood sizes of JU1249 and JU2825 were similar when the two strains were uninfected, or infected with any of the three N. parisii or N. ausubeli JUm2799.

With N. ausubeli JUm2526, JU1249 had a larger brood size than JU2825 (p-value is 0.004).

Together, these results demonstrated that upon N. ausubeli JUm2526 infection, C. elegans JU1249 has a higher tolerance in the brood size compared to JU2825. This higher tolerance of JU1249 in the brood size may be one factor that leads to the better performances of JU1249 in the food consumption tests and the final winning in the competition assays with JU2825.

II.4. Future studies on the genetic basis of C. elegans sensitivity variation to microsporidia infection

With the C. elegans strains JU1249 and JU2825 that show contrasted phenotypes in the sensitivity to microsporidia infection, we plan to further study the genetic basis of this natural variation. As proved possible by the competition tests performed between JU1249 and JU2825 in the presence and absence of microsporidia infection, we plan to use Extreme QTL mapping (X-QTL) to detect the underlying genetic loci. X-QTL mapping is a developed bulk segregation technique that employs selection-based phenotyping to large groups of crossed progeny for the trait of interest [START_REF] Ehrenreich | Dissection of genetically complex traits with extremely large pools of yeast segregants[END_REF][START_REF] Chevalier | Efficient linkage mapping using exome capture and extreme QTL in schistosome parasites[END_REF]. X-QTL has been used on yeast, schistosome and Arabidopsis, and proven to be a simple and costeffectively mapping method for traits that are selectable [START_REF] Ehrenreich | Dissection of genetically complex traits with extremely large pools of yeast segregants[END_REF][START_REF] Chevalier | Efficient linkage mapping using exome capture and extreme QTL in schistosome parasites[END_REF][START_REF] Yuan | Extreme QTL mapping of germination speed in Arabidopsis thaliana[END_REF]. It allows a much larger sample size that facilitates detection of multiple loci with small effects.

To assess the genetic basis of C. elegans sensitivity variation to microsporidia infection using X-QTL methods, mainly four steps have to be performed. The first step is to cross JU1249 and JU2825 and then generate bulked segregating populations. In the second step (phenotyping), the bulked segregating populations will be treated with or without microsporidia infection and then maintained for several generations. During the maintenance, individuals with microsporidia-resistant (or tolerant) traits are assumed to be selected and accumulated in the population. As indicated by the competition assays between JU1249 and JU2825, the maintenance should be continued at least for 30 days (about 10 to 12 generations). The third step (genotyping) is to sequence each population as a pool. The final step is to analyze the sequenced data of each pool. The frequencies of genetic markers (i.e.

SNPs) of each pool will be analyzed and compared. The assumption is that the genome regions containing the microsporidia-resistant (or tolerant) loci will show different frequencies between infected populations and uninfected populations, while other genome regions have equal frequencies between populations with different treatments.

II.5. Other future directions

II.5.1. Screening of other C. elegans strains

Together, 22 C. elegans strains were compared for their sensitivity to Nematocida infection in three experiments. Though no C. elegans strain was found to be fully resistant to the tested N.

parisii or N. ausubeli, the natural variation was determined and three interesting pairs of C.

elegans (CB4856 versus N2, JU1249 versus JU2825, CB4856 versus ED3077) were found. It would be interesting to compare the sensitivity among more C. elegans strains, which may provide many windows to study host-pathogen interactions and coevolution.

II.5.2. Natural variation of C. briggsae in sensitivity to microsporidian infection

In addition to C. elegans, natural sensitivity variation can also be studied in C. briggsae. C.

briggsae is one of the most common nematode hosts to microsporidia infections in our collection. Nine C. briggsae strains were naturally found infected with four Nematocida species. C. briggsae was further found susceptible to N. homosporus. It will be interesting to study the natural sensitivity variation among C. briggsae wild isolates.

II.5.3. Natural variation of Nematocida virulence to C. elegans

The virulence difference between N. ausubeli and N. parisii was investigated by single-cell inoculation (Balla et al., 2016). The developmental kinetics and symptom severity to the host of N. ausubeli infection are different from those after N. parisii infection. On average, single N. ausubeli infection spreads faster across host intestinal cells, and forms spores earlier than N. parisii (Figure 38A). Meanwhile, compared to N. parisii, N. ausubeli infection leads to a much stronger negative effect on host growth and fecundity (Figure 38B, C) (Balla et al., 2016). Thus, N. ausubeli seemed more virulent than N. parisii in C. elegans. Nevertheless, to compare the virulence between N. parisii and N. ausubeli, it is necessary to test other strains of both species. In the study of Balla et al. (2016), N. ausubeli ERTm2 was proven to be more virulent than N. parisii ERTm1 to C. elegans. However, RESULT II.3.2 indicated that N. parisii JUm1249 is more virulent than N. ausubeli JUm2009 and N. ausubeli JUm2526 to C. elegans, as N. parisii JUm1249 induced higher pathogen loads in both C.

elegans strains than either N. ausubeli infection did (Figure 28A). Furthermore, the study (DISCUSSION II.3, Figure 37) on brood size of JU1249 and JU2825 with three N. parisii strains and three N. ausubeli strains indicated the variation in virulence of different microsporidia, at both inter-and intraspecific levels. It would be interesting to compare virulence of different microsporidia species and strains using the single cell inoculation method.

In addition, we found that N. ausubeli induced a less robust host transcriptional response than N. parisii and some other Nematocida species in C. elegans, by using transgenic C. elegans strains with GFP reporter genes that are highly upregulated upon N. parisii infection (paper

III. The suppression effect of microsporidia infection on the progressive sterility in C. elegans

Our preliminary tests demonstrated that N. parisii and N. ausubeli infections are able to attenuate the progressive decrease in fertility in C. elegans with the mortal germline phenotype (Mrt). In the four C. elegans strains we tested, two strains (JU775 and JU2526) with a moderate Mrt phenotype had an increased number of fertile generations upon microsporidia infection. This rescue effect represents an exciting type of host-pathogen interaction and is certainly worth further investigations.

Various negative effects of microsporidia infection on C. elegans have been observed and determined. This rescue effect may be a beneficial effect of microsporidia infection on C.

elegans, which may have great evolutionary and ecological significance. Many interesting questions can be further addressed. For example, is this suppression on C. elegans with Mrt phenotype a specific effect of microsporidia infection? Or are other micro-organisms, such as the Orsay virus, bacteria and fungi, also able to display the suppression? Another direction is to identify the molecular mechanisms of this suppression effect, for which reverse, forward and quantitative evolutionary genetic approaches can be further performed.

Conclusion

Our studies revealed the diverse interactions of microsporidia with C. elegans and related nematodes. We found that N. parisii and N. ausubeli are relatively common pathogens of C. elegans and C. briggsae in the wild. We better described N. ausubeli and further found five new Nematocida species. We also defined two new genera of nematode-infecting microsporidia, named Enteropsectra and Pancytospora, which are genetically distinct from

Nematocida. These microsporidia are diverse in terms of phylogenetic relationships, spore size and shape, host tissue tropism, host cell intracellular localization, cellular exit route, host specificity pattern, virulence and negative effects on hosts, etc. These findings considerably enlarged our knowledge on nematode-infecting microsporidia and on the ecology of C.

elegans and related nematodes.

To date, 48 strains of 10 nematode species have been found naturally infected with microsporidia. These nematode species displayed interspecific susceptibility and intraspecific sensitivity variations to different microsporidia infections. Here we further determined the intraspecific variation of C. elegans in sensitivity to N. ausubeli infection. Two C. elegans strains, JU1249 and JU2825, displayed the largest sensitivity difference, which may be due to their different tolerance. These two strains are proven to be good candidates for future studies on the genetic loci associated with C. elegans sensitivity variation to microsporidian infections. Furthermore, an exciting type of host-pathogen interaction was observed.

Microsporidia infection is able to suppress the progressive decrease in fertility in C. elegans with the mortal germline phenotype (Mrt). Overall, the diverse interactions of microsporidia with Caenorhabditis elegans and related nematodes provide great opportunities to study hostpathogen coevolution.

Figure 1 .

 1 Figure 1. Schematic phylogenetic tree showing the position of microsporidia.The phylogeny is based on the analysis of multiple protein-coding genes. Adapted from[START_REF] Keeling | Phylogenetic place of Microsporidia in the tree of Eukaryotes[END_REF].

Figure 2 .

 2 Figure 2. Morphology and structure of microsporidian spores. A. Diagram of a microsporidian spore. The spore coat consists of an electron-dense exospore (Ex), an electron-lucent endospore (En), and a plasma membrane (Pm). The sporoplasm (Sp) contains a single nucleus (Nu), the posterior vacuole (PV), and ribosomes. The polar tube (PT) is attached to the anterior end of the spore by an anchoring disk (AD) and is divided into two regions: the manubroid or straight portion (M) and the posterior region forming five coils around the sporoplasm. The insert depicts a cross section of the polar tube coils (five coils in this spore). B. Ultrastructure of the mature spore. (a) Longitudinally sectioned spore exhibiting the characteristic organelles (arrowhead indicates vacuolar membrane) (Episeptum inversum); (b) polyribosomes attached to membranes (Napamichum dispersus); (c) circularly arranged polyribosomes (Hamiltosporidium magnivora). Scale bars = 0.5 µm (a); 100 nm (b, c). Abbreviations are the same as in A., with, in addition: f, polar tube; Pa, anterior polaroplast region; Pp, posterior polaroplast region; Ps, polar sac; R, polyribosomes. C. Scanning electron micrograph of extruded polar tube of a spore of Encephalitozoon intestinalis piercing and infecting Vero E6 green monkey kidney

Figure 3 .

 3 Figure 3. Diagram of a typical microsporidia life cycle. The life cycle of microsporidia includes extracellular and intracellular phases. Spores are the only life stage that survives outside hosts. The intracellular phase contains the meront stage and the sporont stage. Adapted from Cali and Takvorian (2014).

Figure 4 .

 4 Figure 4. Phylogeny of 71 microsporidia species.The phylogeny is based on SSU rDNA. Font color indicates the host's habitat: blue for freshwater, brown for terrestrial, green for marine and blue green for hosts that spend part of their life cycle in both marine and freshwater habitats. Nematocida parisii, the natural pathogen of C. elegans, is marked by a red frame. Adapted from[START_REF] Vossbrinck | Phylogeny of the Microsporidia[END_REF].

Figure 7 .

 7 Figure 7. Life cycle of C. elegans at 22°C. 0 min is the fertilization. Numbers in blue along the arrows indicate the length of time the animal spends at a certain stage. First cleavage occurs at about 40-minutes postfertilization. Eggs are laid outside at about 150-minutes post-fertilization. The length of the animal at each stage is marked next to the stage name in micrometers (µm).From[START_REF] Hall | Introduction[END_REF].

Figure 8 .

 8 Figure 8. Posterior region of Nematode body. Diagram of the cross section of the posterior region of nematode body. Adapted from Hall and Altun (2009).

Figure 10 .

 10 Figure 10. Phenotypic polymorphisms between N2 and wild C. elegans strains. N2 (left) and CB4857 (right) display distinct clumping (Clp) behavior. Adapted fromBarrière and Félix (2005a).

Figure 11 .

 11 Figure 11. Phylogeny of Nematoda. The position of nematodes in the animal tree is indicated in the inset. The main panel is the phylogeny of Nematoda, which contain three large monophyletic groups: Enoplia, Dorylaimia and Chromadoria. C. elegans belongs to the infraorder Rhabditomorpha (suborder Rhabditina, order Rhabditida, family Rhabditidae), which

Figure 13 .

 13 Figure 13. Phylogeny of Caenorhabditis.The main panel shows the phylogeny of the major Caenorhabditis groups with 27 species (former numerical identifiers in parentheses). The three species with selfing mode of reproduction are in red, while the remaining known Caenorhabditis species are male and female. Geographic sampling locations and number of isolated strains are shown on the right of the phylogeny. Adapted fromCutter (2015);Frézal and Félix (2015).

Figure 14 .

 14 Figure 14. Phylogenetic relationships in the Oscheius genus. From Félix (2006).

  . Gramnegative bacterial pathogens such as Pseudomonas aeruginosa, Salmonella enterica and Salmonella typhimurium, Gram-positive bacterial pathogens Enterococcus faecalis, Straphylococcus aureus and Bacillus thuringiensis could all cause intestinal infections and kill C. elegans (Figure 15) (Aballay and Ausubel, 2002; Darby, 2005).

Figure 15 .

 15 Figure 15. Pathogens of C. elegans and their infection routes and tropisms. Adapted from Engelmann and Pujol (2010).

Figure 16 .

 16 Figure 16. Leucobacter infections in Caenorhabditis. A. The swollen tail of a C. tropicalis JU1635 individual, with bacterial coating on the surface. The arrow indicates the swollen tail. B. L. celer subsp. astrifaciens CBX151 induced a "worms-star" of a quartet of C. elegans individuals held together by their tails. C. Dying C. elegans after 20-hour exposure to L. musarum subsp. musarum CBX152. The anterior gut (arrow) is compressed by swollen vacuoles (asterisks) in the body cavity. D. The swollen tail of C. elegans infected with L. musarum subsp. musarum CBX152. Long arrow indicates the swollen tail; short arrow indicates the bacteria adhered around the anus. Arrowhead indicates the anal opening in A. and D. Adapted from Hodgkin et al. (2013).

Figure 17 .

 17 Figure 17. Viral infections in intestinal cells of wild Caenorhabditis isolates. (A-B) C. elegans JU1580. A. An adult hermaphrodite infected with the Orsay virus displays an abnormally large intestinal cell that is probably due to cell fusion. B. An uninfected JU1580 adult. Antero-posterior boundaries between intestinal cells were indicated by arrowheads. (C-E) C. briggsae JU1264 infected with the Santeuil virus show diverse infection symptoms: convoluted apical intestinal border (C), degeneration of intestinal cell structures (D), presence of multi-membrane bodies (E). F. An uninfected JU1264 adult. Scale bars are 10µm. Adapted from Félix et al. (2011).

Figure 18 .

 18 Figure 18. N. parisii infection symptoms in C. elegans. A. The intestine of an uninfected C. elegans. B. Host intestinal granules are displaced by N. parisii meronts, which appear as "grooves" (arrow) in the host intestinal cells. The intestinal lumen is indicated by arrowheads in A and B. C. Large and small rodshaped spores are indicated by larger and smaller arrow, respectively. D. Vesicles containing spores are indicated by an arrow. Scale bar is 10µm. Adapted from Troemel et al. (2008).

Figure 19 .

 19 Figure 19. Ultrastructure of N. parisii infection in the C. elegans intestine. A. An irregular shaped multinucleate meront indicated with an arrow. A host vesicle is marked with an asterisk. Scale bar is 1µm. B. A later stage meront with two nuclei and more regularly shaped plasma membrane. C. and D. are sporonts with electrondense membranes. E. Developing N. parisii spores. F. A small sized spore. G. A larger sized spore. Arrowheads indicate cross-sections of polar tube coils in F and G. H. Vesicles of spores. B-H, scale bar is 500 nm. PP refers to the polaroplast membrane; PV refers to the posterior vacuole. Adapted from Troemel et al. (2008).

Figure 21 .

 21 Figure 21. Phase I of N. parisii exit from C. elegans intestinal cells. A. Diagram of Phase I of N. parisii exit. Green is ACT-5 and orange is IFB-2. Pathogen cells are depicted in red and nuclei depicted in black. B. Transmission electron microscopy shows the intact terminal web (TW) in uninfected C. elegans intestine. C. Gaps (arrows) in the terminal web of C. elegans infected with N. parisii. Double arrows indicate microvilli (MV). Lu indicates lumen. In B and C, scale bar is 1 µm. Adapted from Troemel et al. (2008); Szumowski et al. (2012).

Figure 22 .

 22 Figure 22. Phase II of N. parisii exit from C. elegans intestinal cells. A. Diagram of Phase II of N. parisii exit. See text for details. B. Transmission electron microscopy of an immature N. parisii spore. The spore membrane (SM) is surrounded by an additional membrane (AM). C. A mature spore (asterisk) spanning the luminal membrane. D. Magnification of box from C. Arrows indicate the continuous host apical membrane (HM) that surrounds the spore. (B-E) AJ, apical junction; EN, endospore; EX, exospore; MV, microvilli; PT, polar tube; TW, terminal web. Scale bars: B and D, 500 nm; C, 1 µm. Adapted from Szumowski et al. (2014).

Figure 23 .

 23 Figure 23. Natural variation among different C. elegans strains in the response to N. parisii infection. A. Survival curves of six C. elegans strains upon infection by N. parisii. B. Normalized pathogen load of each strain from (A) at 30 hpi, measured by qRT-PCR targeting an N. parisii SSU rRNA. C. Survival curves of uninfected animals, and infected animals inoculated at the L1 or L4 stage. D. Lifetime fecundity of uninfected animals, and infected animals inoculated at the L1 or L4 stage. Adapted from Balla et al. (2015).

  Luallen and Emily R. Troemel from University of California at San Diego. A Large Collection of Novel Nematode-Infecting Microsporidia and Their Diverse Interactions with Caenorhabditis elegans and Other Related Nematodes Gaotian Zhang 1,2 , Martin Sachse 3 , Marie-Christine Prevost 3 , Robert J. Luallen 4 , Emily R. Troemel 4 , Marie-Anne Fe ´lix 1 * 1 Institut de Biologie de l'Ecole Normale Supe ´rieure, CNRS, Inserm, ENS, PSL Research University, Paris, France, 2 School of Life Sciences, East China Normal University, Shanghai, China, 3 Ultrapole, Institute Pasteur, Paris, France, 4 Division of Biological Sciences, Section of Cell and Developmental Biology, University of California San Diego, La Jolla, California, United States of America * felix@biologie.ens.fr

Fig 1 .

 1 Fig 1. Isolation of nematode-infecting microsporidia. A. Morphological screen for infected worms (here C. elegans JU2816). Compared to uninfected worms, infected adult worms have a paler body color. Note that the pale body color may result from many environmental conditions, and thus these animals were further screened by Nomarski optics for microsporidian infections. Scale bar: 0.2 mm. B. Geographic distribution of our collection of nematode-infecting microsporidia. Sampling locations are represented by differently colored symbols based on microsporidian species. Black symbols were used when different microsporidian species were found in the same location. The world map is cited from http://www.d-maps.com/carte.php?num_car=3267&lang=en. The France map is cited from http:// www.d-maps.com/carte.php?num_car=2813&lang=en. doi:10.1371/journal.ppat.1006093.g001

Fig 2 .Fig 3 .

 23 Fig 2. Bayesian inference SSU rDNA phylogeny of microsporidia species. SSU rDNA sequences from 45 nematode-infecting microsporidia species and 60 other microsporidia species in the databases were used. The tree was generated using MrBayes v3.2.2 and refined by FigTree v1.4.2. Model Kimura 2-Parameter (K2P) was applied. Branch colors show the posterior probability, with the corresponding color code shown on the left. The light green boxes designate microsporidia infecting terrestrial nematodes and lightblue rectangles designate those infecting marine nematodes. Scale bar indicates expected changes per site. Branches of species with more than one strain were compressed. doi:10.1371/journal.ppat.1006093.g002

  0.335 0.326 0.289 0.249 0.236 0.222 0.122 0.040 0.039 0.039 0.039 0.035 0.045 0.035 0.040 0.038 0.039 0.031 0.037 0.065 9 Enteropsectra longa n. sp. (1) a

Fig 4 .

 4 Fig 4. Spore morphology of the different Nematocida species by Nomarski optics. A. Wild Caenorhabditis elegans strain JU1249, with Nematocida parisii infection. B. Wild C. elegans strain JU2520, with Nematocida ausubeli infection. C. Wild C. briggsae stain JU2747, with N. major infection. D. Wild Oscheius tipulae strain JU1510, with N. minor infection. A ~D, large and small spore classes are indicated by larger and smaller arrows, respectively. Spores in each class are smaller than those in A-C in the corresponding class. E. Wild Rhabditella typhae strain NIC516, with N. homosporus infection. A single class of spore size is observed, often clustered inside vesicles as indicated with arrows. F. Wild Procephalobus sp. strain JU2895, with N. ciargi infection. A single class of spore size is observed, often clustered inside vesicles as indicated with arrows. Scale bar: 10 μm. doi:10.1371/journal.ppat.1006093.g004

cFig 5 .

 5 Fig 5. Ultrastructural observations of Nematocida ausubeli. Transmission electron micrographs of N. ausubeli strain JUm2009 after high-pressure freezing/freeze substitution. A. N. ausubeli meront with two nuclei. B. A multinucleated meront. C. Late stage meront. D. Formation of sporoblasts by polysporous sporogony. E. Cluster of sporonts after sporogony; the arrowheads indicate the nascent polar tube and the arrow indicates the dense membrane structure. F. Sporoblast with a maturing anchoring disk and the dense membrane structure on the future posterior side of the spore (large arrow). Four nascent polar tube coil cross-sections (arrows) are visible, suggesting that this sporoblast may form a spore of large size. G. Late stage sporoblast. The arrow indicates the polar tube. H. Mature spore with surrounding additional membrane (arrows). The internal side of this membrane is coated. I. Mature spore with polar tube indicated by arrow. The

Fig 6 .

 6 Fig 6. Cell exit modes of Nematocida ausubeli and Enteropsectra longa. A. Nematocida ausubeli. The top panel is an electron microscopy image of a Nematocida ausubeli spore (large arrow) exiting from the intestinal cell into the lumen. Arrows indicate the apical membrane of the host intestinal cell. The hypothetical diagram below illustrates the exit of N. ausubeli spores from intestinal cells by exocytosis. As in N. parisii[START_REF] Szumowski | The small GTPase RAB-11 directs polarized exocytosis of the intracellular pathogen N. parisii for fecal-oral transmission from C. elegans[END_REF][START_REF] Szumowski | Small GTPases promote actin coat formation on microsporidian pathogens traversing the apical membrane of Caenorhabditis elegans intestinal cells[END_REF], spores appear surrounded by a membrane that fuses with the apical membrane of the host intestinal cell, resulting in the release of spores. We also observed apparently mature spores without an additional membrane and do not know whether they will later acquire a membrane or exit in another manner. Earlier stages were omitted here for simplicity. B. Enteropsectra longa. The top panel is an electron micrograph of Enteropsectra longa spores exiting from the intestinal cell into the lumen, with the host intestinal cell membrane folding out around the E. longa spores (arrow). The diagram below illustrates the exit of E. longa spore from the intestinal cell. The host intestinal cell membrane folds out around the spore until the whole spore exits the cell, after which the host membrane around the spore seems to disappear. Meronts and sporoblasts are not represented in either panel. Scale bars: 500 nm.

  Fig 6. Cell exit modes of Nematocida ausubeli and Enteropsectra longa. A. Nematocida ausubeli. The top panel is an electron microscopy image of a Nematocida ausubeli spore (large arrow) exiting from the intestinal cell into the lumen. Arrows indicate the apical membrane of the host intestinal cell. The hypothetical diagram below illustrates the exit of N. ausubeli spores from intestinal cells by exocytosis. As in N. parisii[START_REF] Szumowski | The small GTPase RAB-11 directs polarized exocytosis of the intracellular pathogen N. parisii for fecal-oral transmission from C. elegans[END_REF][START_REF] Szumowski | Small GTPases promote actin coat formation on microsporidian pathogens traversing the apical membrane of Caenorhabditis elegans intestinal cells[END_REF], spores appear surrounded by a membrane that fuses with the apical membrane of the host intestinal cell, resulting in the release of spores. We also observed apparently mature spores without an additional membrane and do not know whether they will later acquire a membrane or exit in another manner. Earlier stages were omitted here for simplicity. B. Enteropsectra longa. The top panel is an electron micrograph of Enteropsectra longa spores exiting from the intestinal cell into the lumen, with the host intestinal cell membrane folding out around the E. longa spores (arrow). The diagram below illustrates the exit of E. longa spore from the intestinal cell. The host intestinal cell membrane folds out around the spore until the whole spore exits the cell, after which the host membrane around the spore seems to disappear. Meronts and sporoblasts are not represented in either panel. Scale bars: 500 nm. doi:10.1371/journal.ppat.1006093.g006

  (JUm1396) was found to infect epidermis and muscles of C. brenneri (Fig 7D; S5D, S5E and S5F Fig), thus sharing its tissue tropism with N. displodere, although on a different evolutionary branch. P. epiphaga could also infect C. elegans (N2 reference background) (S7F Fig) and C. briggsae (AF16).

  could be seen at this stage, with intranuclear microtubules and spindle plaques at the nuclear membrane (Fig 8E; S6A Fig). Signs of sporogenesis then developed, with a nascent polar tube (Fig 8F-8H; S6B Fig). The spore membrane and nascent wall appeared wrinkled (Fig 8G) before becoming smooth in mature spores (Fig 8H-8J). The spore wall with its endospore and exospore layers could be clearly observed (Fig 8J). An anchoring disk formed (Fig 8I and 8K), but the polaroplast membranes were less developed than in Nematocida species. In most spores, the polar tube presented a single section (Fig 8G, 8H and 8J; S6D Fig). The polar tube however could be seen to turn on the posterior side of the spore (S6C Fig) and occasionally two polar tube sections could be counted in the same spore section (S6E and S6F Fig). The polar tube thus likely folds back anteriorly on the posterior side of the spore on a short part of its length.

  Fig 6B; S6G-S6I Fig). We further focused on spore sections in the intestinal lumen and saw both spores with a surrounding membrane (S6I Fig) and spores without any membrane (Fig 8K).

Fig 7 .

 7 Fig 7. Spore morphology of Enterospectra and Pancytospora species by Nomarski optics. A. Wild Oscheius sp. 3 strain JU408, with Enteropsectra longa infection. Gut lumen was indicated. Arrow indicates spores, here long and thin spores that are aligned along the apical side of intestinal cells. B. Wild O. tipulae strain JU2551, with Enteropsectra breve infection. Arrow indicates spores, here small spores along the apical side of intestinal cells. C. Wild O. tipulae strain JU1505, with Pancytospora philotis infection. Spores are found throughout the intestinal cell. D.Wild Caenorhabditis brenneri strain JU1396, with Pancytospora epiphaga infection. Spores are seen in the epidermal cells in the tail that does not contain any gut tissue (posterior to the rectum). Anterior is to the right. The "fur" on the outside of the cuticle is formed by unidentified bacteria (see[START_REF] Hodgkin | Two Leucobacter strains exert complementary virulence on Caenorhabditis including death by worm-star formation[END_REF] for another example). Scale bar: 10 μm in A-D. E. Two-dimensional diagram of Oscheius sp. 3 intestine infected with Enteropsectra longa. The intestine is formed of polarized epithelial cells. Enteropsectra longa starts to form spores along the apical side of the intestinal cells.

Fig 8 .

 8 Fig 7. Spore morphology of Enterospectra and Pancytospora species by Nomarski optics. A. Wild Oscheius sp. 3 strain JU408, with Enteropsectra longa infection. Gut lumen was indicated. Arrow indicates spores, here long and thin spores that are aligned along the apical side of intestinal cells. B. Wild O. tipulae strain JU2551, with Enteropsectra breve infection. Arrow indicates spores, here small spores along the apical side of intestinal cells. C. Wild O. tipulae strain JU1505, with Pancytospora philotis infection. Spores are found throughout the intestinal cell. D.Wild Caenorhabditis brenneri strain JU1396, with Pancytospora epiphaga infection. Spores are seen in the epidermal cells in the tail that does not contain any gut tissue (posterior to the rectum). Anterior is to the right. The "fur" on the outside of the cuticle is formed by unidentified bacteria (see[START_REF] Hodgkin | Two Leucobacter strains exert complementary virulence on Caenorhabditis including death by worm-star formation[END_REF] for another example). Scale bar: 10 μm in A-D. E. Two-dimensional diagram of Oscheius sp. 3 intestine infected with Enteropsectra longa. The intestine is formed of polarized epithelial cells. Enteropsectra longa starts to form spores along the apical side of the intestinal cells. doi:10.1371/journal.ppat.1006093.g007

  endoplastic reticulum folding around the microsporidia. H. Late stage sporoblast in the center, mature spore on the top left; arrows indicate polar tubes. I. Mature spore with the anterior part of the polar tube, including the anchoring disk. J. Cross-section of mature spores. The exospore and endospore layers are shown in the inset. Arrowheads indicate polar tubes. K. Two mature spores in the intestinal lumen that do not show an additional membrane around them. Low magnification inset shows the positions of the two spores in the lumen and arrowhead indicates host microvilli. L. Low magnification view of cross-section of host, with the intestinal lumen in the center. E. longa spores (arrowheads) concentrate around the apical membrane of intestinal cell, while meronts and early sporonts are on the basal side. Scale bar is 500 nm, unless indicated otherwise. A, anchoring disk; Chr: chromatin; Ex, exospore; En, endospore; Lu, lumen; M, meront; Mi: host mitochondrion; Mv, host microvilli; Nu, nucleus; HNu host nucleus; Pt, polar tube; RER, rough endoplastic reticulum; St: sporont. doi:10.1371/journal.ppat.1006093.g008

Fig 9 .

 9 Fig 9. Responses of C. elegans strains with transcriptional reporters C17H1.6p::GFP and F26F2.1p::GFP to exposure by different microsporidia. Strains ERT54 carrying C17H1.6p::GFP (A) and ERT72 carrying F26F2.1p::GFP (B) were analyzed for GFP induction at different time points after infection with different microsporidia and the proportion of animals with GFP induction is shown. GFP was reproducibly induced in ERT54 and ERT72 upon infection with N. parisii, N. major and N. homosporus, while GFP signal was rarely observed in ERT54 and ERT72 inoculated with N. ausubeli or E. longa or the negative control. N. ausubeli did infect the C. elegans reporter strains, as monitored by DIC as in Table5. C. Transcript levels for three genes were measured after 4 hours of infection of N2 C. elegans by N. parisii (ERTm1) and N. ausubeli (ERTm2). The fold increase in transcript level was measured relative to uninfected N2 levels. Infection dose was normalized between Nematocida by successful invasion events counted as intracellular sporoplasms at 4 hpi. To independently compare the microsporidian doses in parallel to the transcript quantification, we also measure the levels of Nematocida SSU rRNA after 4 hours of infection of C. elegans in the same experiment: we found that the rRNA level measured after infection with ERTm2 was 1.25-fold higher than that with ERTm1.

doi: 10 .

 10 1371/journal.ppat.1006093.g009 species(Figs 2 and 3; S2 Fig), this evolutionary change can be polarized: N. ausubeli seems to have lost the capacity to activate these transcriptional reporters as strongly as its closest relatives, or has acquired the capacity to inhibit their activation. Thus, although morphologically quite similar and both are able to infect C. elegans, N. parisii and N. ausubeli elicit distinct host responses.

Fig 10 .

 10 Fig 10. Summary of the interactions between rhabditid nematodes and microsporidia in the wild and in laboratory. A mosaic green color means that the corresponding natural infection was found. Plain green means that the infection worked in the laboratory and red means that the infection did not work in the laboratory. White: not determined. doi:10.1371/journal.ppat.1006093.g010

  Fig 7; S5 Fig; Table

  Taxonomic descriptions. Phylum Microsporidia Balbiani 1882 Nematocida ausubeli n. sp. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org:act:6-D7E3D0D-3348-4885-BF64-DF1EE1B7EEBA

  Enteropsectra n. gen. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org:act:33CE2667-0109-44DA-9878-34FAD4A2F96B This is a novel microsporidian lineage within microsporidian clade II (ref), with Orthosomella, Liebermannia as the closest relatives, based on SSU rDNA phylogenetic analyses. The type species is Enteropsectra longa n. sp. Zhang & Fe ´lix 2016. The genus is named Enteropsectra (feminine, from the Greek "psectra": brush) after the morphological aspect of the spores at the apical side of the intestinal cells of the nematode host, resembling a bottle brush. Enteropsectra longa n. sp. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org:act: FC304EDC-B2CA-4486-BF8F-095BE0B60E45

  Pancytospora n. gen. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org:act:1ADAC856-ED00-49C9-9906-634BAF38B355 This is a novel microsporidian lineage within microsporidian clade IV, with Orthosomella, Liebermannia and Enteropsectra n. gen. as the closest relatives, based on SSU rDNA phylogenetic analyses. The type species is Pancytospora philotis n. sp. Zhang & Fe ´lix 2016. The genus is named Pancytospora (feminine) after the distribution of the spores throughout the cells. Pancytospora philotis n. sp. Zhang & Fe ´lix 2016. LSID urn:lsid:zoobank.org: act:5EA01A65-F5F4-4EE6-A1B6-2726F9CE8579

F

  i = Mean fluorescence intensity of the plate at timepoint i; F backi = Mean fluorescence intensity of the plate with regular OP50 without C. elegans at timepoint i; F nowormi = Mean fluorescence intensity of the plate with OP50-GFP without C. elegans at timepoint i.

Figure 25 .

 25 Figure 25. Variation in sensitivity against N. ausubeli infection among nine C. elegans wild isolates.

Figure 26 .

 26 Figure 26. OP50-GFP intensity of nine C. elegans strains at two timepoints. The OP50-GFP intensity of the plates with uninfected C. elegans (at 72h) and infected C. elegans (at 120h) is shown. Data are mean values of three replicates (data marked with "*" are mean values of two replicates), with bars represent standard errors. The nine strains on the X axis were ordered from left to right by their normalized OP50-GFP intensity at 72 hours. JU2825 and JU1249, which displayed contrasted phenotype, were highlighted with green frames.

Figure 27 .

 27 Figure 27. JU2825 is more sensitive than JU1249 to microsporidia infection. Food consumption of C. elegans strains JU1249 and JU2825 was monitored over time in the presence and absence of microsporidia. Two N. ausubeli strains (JUm2009 and JUm2526) and one N. parisii strain (JU1249) were used in the infection. Data are mean values of three replicates, with bars represent standard errors.

Figure 28 .

 28 Figure 28. FISH analysis of the pathogen load and the infection rate of JU1249 and JU2825 upon three different microsporidia infections.

Figure 29 .

 29 Figure 29. Influence of microsporidian infection in the experimental competition between C. elegans JU1249 and JU2825.

Figure 30 .

 30 Figure 30. Food consumption of JU2526 with or without microsporidia infection.

Figure 31 .

 31 Figure 31. The rescue effect of microsporidian infection on the Mrt phenotype in C.

Figure 33 .

 33 Figure 33. Phylogenies of different N. parisii, N. ausubeli and N. ironsii strains. A. SSU rDNA phylogeny. B. ß-tubulin phylogeny. The phylogeny with the ß-tubulin gene shows further relationships among different N. parisii strains, as well as among different N.

Figure 34 .

 34 Figure 34. Diagram of N. parisii exiting strategy from C. elegans intestinal cells. The model for actin regulation of N. parisii spore exocytosis and compensatory endocytosis in C. elegans intestinal cells. Left panel: At 43 hpi, the RAB-11 labeled SCCs transfer to and fuse with the apical membrane of the intestinal cells, and acquire (a) a PGP-1 membrane marker or (b) both PGP-1 and ACT-5 markers. Formation of ACT-5 coats is dependent on the Rho GTPases ced-10 and cdc-42, as well as rab-5 and rab-11. Right panel: At 48 hpi, spores has overtaken the host cell, with a lot of exited spores. Endocytic spore-filled vesicles were coated with CHC-1 (clathrin heavy chain), PGP-1 and ACT-5. Actin in the microvilli and the terminal web is represented in bright red; the apical plasma membrane with PGP-1 is in green. Adapted from Szumowski et al. (2016).

1 .

 1 N. parisii was the first known natural intracellular pathogen of C. elegans. C. elegans is a successful model organism, with several special features and numerous developed tools.Since the discovery of N. parisii, exciting progresses using N. parisii and C. elegans haveII. Natural variation of Caenorhabditis elegans in sensitivity to microsporidian infection II.Intraspecific variation of C. elegans sensitivity to microsporidia infectionThe natural sensitivity variation of C. elegans to microsporidia infections has been studied in different sets of C. elegans strains to different microsporidia: 1), the sensitivity of six C.

Figure 35 .

 35 Figure 35. Variation in sensitivity against N. parisii and N. ausubeli among nine C. elegans wild isolates. The Y-axis is the normalized intensity of the fluorescence of OP50-GFP. The X-axis is the scanning time point. Each plate was scanned every day until full depletion of bacteria on the plate. Data are mean values of three replicates, with bars represent standard errors. The analysis and the figure were performed by Clément Dubois.

Figure 36 .

 36 Figure 36. Contrasted sensitivity variation of ED3077 and CB4856 upon different infections. Food consumption test (a repeated test compared to Figure 35) of ED3077 and CB4856 upon N. parisii JUm2816 and N. ausubeli JUm2009. Data are mean values of three replicates, with bars represent standard errors. The figure was performed by Clément Dubois.

  Several reasons may lead to the different results from the two experiments. Firstly, different infection protocols were used in the two experiments, as we measured the fitness of C. elegans from a population with stable microsporidia infection (ten days post-inoculation), whereasBalla et al. (2015) measured the mortality rates of infected C. elegans immediately (two days post-inoculation). Secondly, we used food consumption tests to compare the sensitivity of different strains. The correlation of the performance of C. elegans in the food consumption test and its mortality is not clear yet. Thirdly, different C. elegans strains were compared in the two experiments. Different C. elegans strains may display different defensive strategies. Finally,Balla et al., 2015 used N. ironsii ERTm5 to apply the inoculation, whereas we used N. parisii and N. ausubeli. The responses of C. elegans to different microsporidia infections may be different.

Figure 37 .

 37 Figure 37. The brood size of C. elegans JU1249 and JU2825 with or without microsporidia infection.In each treatment, 5 L4 stage hermaphrodite C. elegans from a stable population (ten days post infection) were transferred to a fresh plate and the number of their progenies were counted. The total number of progenies per hermaphrodite is showed in the figure. Data are mean values of three replicates, with bars represent standard errors. In the pairwise comparisons, JU1249 and JU2825 showed a significant (p-value is 0.004, marked with "**") brood size difference only upon the infection of N. ausubeli JUm2526. The analysis and the figure were performed by Clément Dubois.

Figure 38 .

 38 Figure 38. Growth and virulence variations of N. parisii and N. ausubeli (N. sp. 1). Infections were initiated by single-cell inoculation in N2. The N. parisii strain ERTm1 and the N. ausubeli strain ERTm2 were used here. A. Growth kinetics showed by copy number (solid lines) and spore formation (dashed lines). B. Sizes of animals over time with different infections and no infection. C. Eggs laid by worms with different infections or no infection, at 60 hpi. Adapted from Balla et al. (2016).

Figure 9 )

 9 Figure 9). Further studies byTroemel et al. showed that N. ausubeli inhibited the induction of host reporter genes by N. parisii in co-infection experiments. These results suggested that N. ausubeli and N. parisii are different in their virulence. It would be interesting to compare virulence of different microsporidia species and strains using transgenic C. elegans strains, and further study the competition of N. ausubeli and N. parisii in the same host.

Table 1 . Collection of wild nematode-infecting microsporidia strains A. Nematocida parisii infection Host Strain Country / Year Sample composition Spore feature

 1

	Caenorhabditis elegans	ERTm1 a	France / 2004	compost	2S c
		ERTm3 a	France / 2007	rotting apple	2S c
		JUm1248 a b	France / 2007	rotting apple	n. d.
		JUm1249 b	France / 2007	rotting apple	2S
		JUm1253 b	France / 2007	rotting apple	n. d.
		ERTm4 a	France / 2008	compost	2S c
		JUm1762 b	France / 2009	rotting stem	2S
		JUm1893 b	France / 2009	rotting apple	2S
		JUm2106	France / 2011	compost	2S
		JUm2131	France / 2011	rotting stem	2S
		JUm2132	France / 2011	rotting stem	2S
		JUm2287	France / 2013	compost	2S
		JUm2816	France / 2014	compost	n. d.
	Caenorhabditis briggsae	JUm1254	France / 2007	rotting apple	2S
		ERTm5 a,d	Hawaii / 2011	rotting fruit	2S
		JUm2590	France / 2013	rotting apple	2S
		JUm2793	France / 2014	compost	n. d.
	Host	Strain	Country / Year	Sample composition	Spore feature
	Caenorhabditis elegans	JUm2009	France / 2010	rotting apples	2S
		JUm2520	Portugal / 2013	rotting fruit	2S
		JUm2526	Portugal / 2013	rotting fruit	2S
		JUm2586	France / 2013	rotting pear	2S
		JUm2671	France / 2011	rotting apple	2S
		JUm2825	France / 2014	rotting stem	2S
	Caenorhabditis briggsae	ERTm2 a	India / 2007	rotting mix	2S c
		ERTm6 a	Cape Verde / 2009	soil	2S
		JUm2799	Germany / 2014	rotting fruits	2S
	Caenorhabditis remanei	JUm2796	Germany / 2014	rotting stems	n. d.

B. Nematocida ausubeli (N. sp. 1) infection

The reference strain of each newly found species is in bold. 2S: two distinct sizes of spores; n. d.: not determined. a Strains and names published previously b 18S sequences courtesy of Aurore Dubuffet and Hinrich Schulenburg c Morphological feature described in

Table 2 . Collection of other microsporidia species infecting wild nematodes Microsporidia Nematode host Strain Country / Year Sample composition Spore feature

 2 Caenorhabditis and other nematodes are Nematocida parisii and Nematocida ausubeli n. sp., with 17 and 10 strains, respectively. They were found in several species of the Elegans group of Caenorhabditis nematodes.

	Nematocida	C. briggsae	JUm2507	Thailand / 2013	rotting fruit	2S
	major n. sp.	C. briggsae	JUm2747	Guadeloupe / 2014	rotting fruit	2S
		C. tropicalis	JUm2751	Guadeloupe / 2014	rotting fruit	n. d.
	Nematocida	Oscheius tipulae	JUm1510	Czech Republic / 2008	rotting apple	2S
	minor n. sp.	O. tipulae	JUm2772	Armenia / 2014	rotting fruit	n. d.
	Nematocida	O. tipulae	JUm1504	France / 2008	rotting stem	1S
	homosporus n. sp.	Rhabditella typhae	NICm516	Portugal / 2013	rotting apple	1S
	Nematocida	Procephalobus sp.	JUm2895	Spain / 2015	rotting fruit	1S
	ciargi n. sp.
	Nematocida	C. sp. 42	NICm1041	French Guiana / 2014	flower	n. d.
	sp. 7
	Enteropsectra	Oscheius sp. 3	JUm408	Iceland / 2002	compost	1S, LT, AP
	longa n. sp.
	Enteropsectra	O. tipulae	JUm2551	France / 2013	rotting apple	1S, SR, AP
	breve n. sp.	O. tipulae	JUm1483	France / 2008	rotting plum	1S, SR,AP
		O. tipulae	JUm1456	France / 2008	rotting fruit	n. d.
	Pancytospora	O. tipulae	JUm1505	France / 2008	rotting apple	LT
	philotis n. sp.	O. tipulae	JUm1460	France / 2008	rotting snail	LT
		O. tipulae	JUm1670	France / 2009	rotting apple	LT
		O. tipulae	JUm2552	France / 2013	rotting stem	LT
	Pancytospora	C. brenneri	JUm1396	Colombia / 2008	rotting fruit	LT
	epiphaga n. sp.

The reference strain of each newly found species is in bold. 2S: two distinct sizes of spores; 1S: one size of spores LT: long, thin rod; SR: small rod (see dimensions in Table

3

); AP: form spores first along the apical side of the intestinal cells. n. d.: not determined doi:10.1371/journal.ppat.1006093.t002 microsporidia species in our collection of

Table 3 . Molecular distances of microsporidia SSU rDNA.

 3

	Species	1	2	3	4	5	6	7	8	9	10	11	12	13	14	15	16	17	18	19	20	21
	1 Nematocida
	parisii (17) a

  The lower left part shows the mean genetic distances between groups and the upper right part is the standard error (SE), with Kimura 2-Parameter model+G, 1000 bootstraps. Mean intra-species, intra-genus or intra-clade divergences are shown in the diagonal if available, with the number of sequences indicated in the parentheses after the name of each group (

	12 Orthosomella	0.392 0.393 0.423 0.395 0.389 0.394 0.362 0.307 0.091 0.084 0.089	/	0.017 0.022 0.023 0.035 0.047 0.046 0.048 0.058 0.084
	operophterae
	(1) a
	13 Pancytospora	0.384 0.378 0.418 0.427 0.374 0.378 0.346 0.269 0.115 0.108 0.113 0.091 0.005 0.021 0.020 0.048 0.052 0.048 0.054 0.070 0.098
	philotis n. sp.
	(4) a
	14 Pancytospora	0.460 0.452 0.490 0.451 0.450 0.455 0.429 0.352 0.141 0.133 0.139 0.124 0.112	/	0.023 0.049 0.056 0.056 0.061 0.068 0.110
	epiphaga n. sp.
	(1) a
	15 Liebermannia	0.405 0.406 0.404 0.406 0.351 0.353 0.329 0.284 0.118 0.118 0.120 0.133 0.123 0.152 0.011 0.037 0.042 0.047 0.051 0.056 0.080
	spp. (3) b
	16 CLADE_IV (19) c 0.510 0.512 0.511 0.512 0.480 0.464 0.455 0.383 0.330 0.331 0.329 0.324 0.405 0.423 0.332 0.280 0.039 0.049 0.038 0.039 0.060
	17 CLADE_III (12) c 0.443 0.443 0.413 0.428 0.428 0.409 0.387 0.355 0.384 0.395 0.389 0.398 0.430 0.465 0.371 0.409 0.223 0.038 0.037 0.040 0.061
	18 CLADE_I (14) c	0.462 0.456 0.459 0.420 0.417 0.396 0.353 0.347 0.404 0.396 0.400 0.389 0.400 0.462 0.404 0.474 0.371 0.169 0.039 0.043 0.068
	19 CLADE_? (3) c	0.417 0.411 0.400 0.394 0.352 0.337 0.319 0.272 0.414 0.420 0.416 0.402 0.446 0.483 0.432 0.392 0.371 0.358 0.255 0.028 0.053
	20 CLADE_V (3) c	0.477 0.471 0.449 0.447 0.403 0.380 0.361 0.310 0.419 0.434 0.423 0.441 0.502 0.513 0.433 0.380 0.373 0.385 0.260 0.127 0.039
	21 Rozella spp. (2) b 0.482 0.478 0.408 0.417 0.410 0.362 0.346 0.473 0.543 0.548 0.541 0.598 0.658 0.712 0.587 0.525 0.528 0.561 0.433 0.322 0.042

0.435 0.436 0.447 0.455 0.413 0.417 0.384 0.318 / 0.004 0.002 0.017 0.020 0.023 0.021 0.036 0.045 0.049 0.049 0.054 0.076 10 Enteropsectra breve n. sp. (1) a 0.420 0.421 0.463 0.439 0.398 0.403 0.370 0.311 0.005 / 0.002 0.016 0.019 0.022 0.021 0.037 0.046 0.048 0.049 0.056 0.077 11 Enteropsectra species (2) 0.424 0.424 0.451 0.443 0.402 0.406 0.374 0.312 0.004 0.004 0.008 0.017 0.019 0.023 0.021 0.036 0.045 0.048 0.049 0.054 0.076 a: species; b: genus; c: larger clade). For a detailed table with all strains, see S3 Table. doi:10.1371/journal.ppat.1006093.t003 Diversity and Specificity of Nematode-Infecting Microsporidia PLOS Pathogens | DOI:10.1371/journal.ppat.1006093 December 12, 2016

Table 4 . Spore sizes of each nematode-infecting microsporidia species, as determined by Nomarski optics.

 4

	Nematode-infecting	Spore sizes (μm)
	Microsporidia	Length	Width
	Nematocida parisii a	3.17 ± 0.22	1.31 ± 0.15
		2.18 ± 0.15	0.80 ± 0.08
	Nematocida displodere b	2.38 ± 0.26	1.03 ± 0.18
	Nematocida ausubeli JUm2009 d	2.80 ± 0.29 (2.21 ~3.26) c	0.96 ± 0.14 (0.72 ~1.30)
		2.04 ± 0.32 (1.36 ~2.92)	0.53 ± 0.10 (0.28 ~0.80)
	Nematocida major JUm2507	3.44 ± 0.23 (3.04 ~3.79)	1.20 ± 0.21 (0.90 ~1.53)
		2.35 ± 0.23 (2.02 ~2.66)	0.57 ± 0.14 (0.41 ~0.77)
	Nematocida major JUm2747	3.29 ± 0.25 (2.87 ~3.69)	1.20 ± 0.23 (0.79 ~1.60)
		2.21 ± 0.26 (1.85 ~2.60)	0.53 ± 0.06 (0.44 ~0.63)
	Nematocida minor JUm1510	1.93 ± 0.18 (1.54 ~2.23)	0.83 ± 0.13 (0.51 ~1.09)
		1.31 ± 0.17 (1.09 ~1.73)	0.55 ± 0.12 (0.35 ~0.73)
	Nematocida homosporus JUm1504	2.07 ± 0.30 (1.69 ~2.50)	0.77 ± 0.12 (0.56 ~0.94)
	Nematocida homosporus NICm516	1.95 ± 0.14 (1.67 ~2.16)	0.69 ± 0.11 (0.56 ~0.91)
	Nematocida ciargi JUm2895	1.39 ± 0.20 (1.54 ~2.24)	0.59 ± 0.13 (0.41 ~0.84)
	Enteropsectra longa JUm408	3.76 ± 0.38 (3.15 ~4.99)	0.49 ± 0.06 (0.35 ~0.68)
	Enteropsectra breve JUm2551	1.81 ± 0.22 (1.34 ~2.14)	0.66 ± 0.12 (0.42 ~0.90)
	Pancytospora philotis JUm1505	3.46 ± 0.48 (2.42 ~4.65)	0.42 ± 0.06 (0.25 ~0.52)
	Pancytospora epiphaga JUm1396	3.71 ± 0.31 (2.99 ~4.22)	0.80 ± 0.09 (0.56 ~0.92)

a

[START_REF] Troemel | Microsporidia are natural intracellular parasites of the nematode C. elegans[END_REF]

b

[START_REF] Luallen | Discovery of a natural microsporidian pathogen with a broad tissue tropism in Caenorhabditis elegans[END_REF]

Table 5 . Nematode-infecting microsporidia specificity.

 5

		N. parisii	N. ausubeli	N. major	N. homosporus	E. longa	E. breve	P. philotis	P. epiphaga
		(JUm2816)	(JUm2009) (JUm2747)	(NICm516)	(JUm408) (JUm2551) (JUm1505)	(JUm1396)
		infected/	infected/	infected/	infected/	infected/	infected/	infected/	infected/
	Species strain Replicate	total	total	total	total	total	total	total	total
	C. elegans	14/20	8/20	11/19	4/12	0/20	\	0/20	10/18 a
	N2	17/20	8/17	10/19	8/18	0/20	\	0/20	15/19 b
	C. elegans	10/19	10/20	14/20	2/20	0/20	0/20	0/20	\
	JU2009	11/19	5/19	5/20	5/15	0/20	\	0/20	\
	C. briggsae	10/17	6/13	\	\	0/20	0/20	0/20	\
	JU2747	14/19	7/17	\	\	0/20	\	0/20	\
	C. briggsae	\	\	13/20	4/15	\	\	\	\
	JU2507	\	\	12/20	9/15	\	\	\	\
	C. briggsae	11/17	11/20	12/20	9/21	0/20	\	0/20	\
	JU2793	12/20	13/20	14/19	10/18	0/20	\	0/20	\
	O. tipulae	\	\	\	13/16	\	\	\	\
	JU1504	\	\	\	13/17	\	\	\	\
	O. tipulae	\	\	\	\	\	6/14	11/20	\
	JU1505	\	\	\	\	\	\	7/20	\
	O. tipulae	0/20	0/20	0/20	0/17	0/20	4/17	3/20	\
	JU1483	0/20	0/20	0/20	1/14	0/20	5/15	8/17	\
	O. tipulae	0/20	0/20	0/20	1/14	\	\	\	\
	JU170	0/20	0/20	0/20	4/20	\	\	\	\
	O. tipulae	\	\	\	\	0/20	9/19	\	\
	JU2551	\	\	\	\	0/20	7/15	\	\
	O. sp. 3	0/20	8/17	0/20	9/20	11/17	3/18	0/20	\
	JU408	0/20	3/17	0/20	9/17	16/16	5/18	0/20	\
	O. sp. 3	0/20	8/18	0/20	11/17	12/16	3/18	0/20	\
	JU75	0/20	4/15	0/20	11/19	12/15	3/13	0/20	\
	R. typhae	\	\	\	5/12	\	\	\	\
	NIC516	\	\	\	3/13	\	\	\	\

Experiments in the same column are in parallel. The same batch of spores was used on the four nematode species, each with two different strains and two replicates each. At 72 hours post inoculation (hpi) or 120 hpi at 23˚C, nematodes found with meronts and/or spores in their intestinal cells were considered infected; otherwise, they were considered uninfected. a Infection were checked at 72 hpi, out of 18 worms, 10 were found infected with meronts, but not spores. b Infection were checked at 96 hpi, out of 19 worms, 15 were found infected with meronts and spores. doi:

10.1371/journal.ppat.1006093.t005

PLOS Pathogens | DOI:10.1371/journal.ppat.1006093December 12, 2016

Acknowledgments

We are very grateful to all sample collectors (M. Asahina, H. Baylis, F. Besnard, A. Zalmanski) and A. Barrière, C. Braendle and L. Fre ´zal for isolating infected strains. We thank M. Botts, K.

This work was supported by: Ecole Normale Supe ´rieure, Centre National de la Recherche Scientifique, Inserm; Fondation pour la Recherche Me ´dicale DEQ20150331704; Fondation Bettencourt-Schueller Coup d'Elan 2011; Fellowship from the China Scholarship Council; National Science Foundation Graduate Research fellowship, NIH R01GM114139 and Burroughs Wellcome Fund fellowship; and National Science Foundation Graduate Research Opportunities Worldwide. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Sequences have been submitted to GenBank under accession numbers KX352724-KX352733, KX360130-KX360167 and KX378155-KX378171.

Author Summary

Microsporidia are microbial parasites that live inside their host cells and can cause disease in humans and many other animals. The small nematode worm Caenorhabditis elegans has recently become a convenient model host for studying microsporidian infections. In this work, we sample Caenorhabditis and other small nematodes and 47 associated microsporidian strains from the wild. We characterize the parasites for their position in the evolutionary tree of microsporidia and for their lifecycle and morphology. We find several new species and genera, especially some that are distantly related to the previously known Nematocida parisii and instead closely related to human pathogens. We find that some of these species have a narrow host range. We studied two species in detail using electron microscopy and uncover a new likely mode of exit from the host cell, by budding off the host cell plasma membrane rather than by fusion of a vesicle to the plasma membrane as in N. parisii. We also find a new species that infects the epidermis and muscles of Caenorhabditis rather than the host intestinal cells and is closely related to human pathogens. Finally, we find that one Nematocida species fails to elicit the same host response that other Nematocida species do. These new microsporidia open up many windows into microsporidia biology and opportunities to investigate host-parasite coevolution in the C. elegans system. For this study, these frozen nematodes were thawed and maintained on nematode growth media (NGM) seeded with E. coli OP50 at 23˚C. The microsporidian strain was identified after the strain identifier of its host nematode strain (itself identified according to C. elegans community rules; http://www.wormbase.org/about/userguide/nomenclature), with an additional "m" between the letters and the numbers for the microsporidia. For instance, a microsporidian strain from the nematode strain JU1762 was named JUm1762. Previously published nematode-infecting microsporidian strains keep their names: ERTm1 (from strain CPA24), ERTm2 (from JU1348), ERTm3 (from JU1247), ERTm4 (from JU1395), ERTm5 (from JU2055) and ERTm6 (from JU1638) [START_REF] Bakowski | Microsporidia infections in Caenorhabditis elegans and other nematodes[END_REF][START_REF] Balla | A wild C. elegans strain has enhanced epithelial immunity to a natural microsporidian parasite[END_REF][START_REF] Bakowski | Genome sequence of the microsporidian species Nematocida sp.1 strain ERTm6 (ATCC PRA-372)[END_REF][START_REF] Cuomo | Microsporidian genome analysis reveals evolutionary strategies for obligate intracellular growth[END_REF][START_REF] Estes | Non-lytic, actin-based exit of intracellular parasites from C. elegans intestinal cells[END_REF] (Tables 1 and2). ERTm4 was previously reported to correspond to N. parisii infection [START_REF] Balla | A wild C. elegans strain has enhanced epithelial immunity to a natural microsporidian parasite[END_REF], but as no sequence data was available in GenBank, we also sequenced the SSU rDNA and β-tubulin genes for this study.

SSU rDNA and β-tubulin gene sequencing

We sought to amplify by PCR and sequence fragments of two microsporidian genes (SSU rDNA and β-tubulin genes) from all potentially infected rhabditid isolates. Ten infected worms were placed in a PCR tube with 10 μl single worm lysis buffer (1X PCR buffer (Dream-Taq Buffer 10X, Theromo Fisher), 1 mM MgCl 2 , 0.45% Tween 20 and 170 ng/μl proteinase K), which was then treated at 60˚C for 60 min, followed by 95˚C for 15 min. This DNA extract was then used as DNA template. To amplify microsporidian SSU rDNA, primers v1f (5'-CA CCAGGTTGATTCTGCCTGAC-3') and 1492r (5'-GGTTACCTTGTTACGACTT -3') [START_REF] Weissli | Utility of microsporidian rRNA in diagnosis and phylogeny: a review[END_REF] were used to amplify strains JUm408, JUm1254, JUm1483, JUm1504, JUm2009, JUm2106, JUm2131, JUm2132, JUm2287, JUm2520, JUm2526, JUm2551, JUm2586, JUm2590, JUm2671 and NICm516. We used v1f and 18SR1492 (5'-GGAAACCTTGTTACGACTT-3') to amplify sequences of JUm1456, JUm1460, JUm1505, JUm1510, JUm1670, JUm2552, JUm2793, JUm2796, JUm2799, JUm2816, JUm2825 and JUm2895. We designed a new pair of primers SPF (5'-GATACGAGGAATTGGGGTTTG-3') and SPR (5'-GGGTACTGGAAATTCCGT GTT-3') for JUm2507, JUm2747, JUm2751 and JUm2772. We failed to amplify SSU rDNA for JUm1501 and NICm1041.

To amplify the microsporidian β-tubulin gene, newly designed forward primer βn1F (5'-A CAAACAGGNCARTGYGGNAAYCA-3') and reverse primer βn1R (5'-TGCTTCAGTRAA YTCCATYTCRTCCAT-3') were used. To obtain the β-tubulin gene sequence of JUm2551 and JUm1456, nested PCR was performed using first primers βn1F and βn1R then βnOF (5'-C CGGACAATATCGTCTTTGG-3') and βnOR (5'-CAGCTCCTGAATGCTTGTTG-3') (S1 Table ). PCR products showing a positive signal by gel electrophoresis were sequenced on both strands on ABI 3730XL sequencing machines (MWG). SSU rDNA of five additional N. parisii strains (JUm1248, JUm1249, JUm1253, JUm1762, JUm1893) were provided by Aurore Dubuffet and Hinrich Schulenburg. The results were analyzed using Geneious v7.1.7 [START_REF] Kearse | Geneious Basic: an integrated and extendable desktop software platform for the organization and analysis of sequence data[END_REF] and compared by BLAST with the NCBI database (http://blast.ncbi.nlm.nih.gov/Blast.cgi). Note that some PCR products could not be amplified (S1 Table ). Especially, we failed to amplify the β-tubulin gene in several Oscheius infections. Both genes fail to amplify for the putative microsporidian infection of the O. tipulae strain JU1501 and this infection could not be characterized.

SSU rDNA and β-tubulin gene sequences have been submitted to GenBank under accession numbers KX352724-KX352733, KX360130-KX360167 and KX378155-KX378171 (S1 Table ).

Phylogenetic analysis

SSU rDNA and β-tubulin gene sequences of microsporidia from this study were analyzed with those of other published microsporidian species and fungi (Rozella spp. for SSU rDNA, Basidiobolus ranarum and Conidobolus coronatus for β-tubulin and concatenated sequences of respectively. The spores are on the apical side of the cell (and are not well infiltrated in these sections). In panel A, on the lower right is seen a nuclear division of a sporont. The mitotic spindle is indicated by an arrowhead. In panel B, the longitudinal section of the polar tube of a spore is indicated by an arrowhead. The white halo that can be seen between the mature spore wall and the cytoplasm is not due to the presence of a membrane but to incomplete infiltration during the preparation of the samples for electron microscopy. Such light-appearing areas are also seen occasionally on the internal side of the spore wall in both observed species. C. Mature spore with inset indicating the turn of the polar tube on the posterior end of the spore. In Table 2, dividing lines were introduced during typesetting that affect the readability of the table. The publisher apologizes for the error. Please see the corrected Table 2

II. Natural variation of Caenorhabditis elegans in sensitivity to microsporidian infection

While studying the diverse microsporidian infections found in the wild nematodes, we also observed natural phenotypic variation in sensitivity of different wild C. elegans strains to the same microsporidian infection. In this chapter, I will focus on the comparison of the sensitivity of ten C. elegans strains upon the same N. ausubeli infection. We chose to focus on the N. ausubeli instead of N. parisii infection in C. elegans with two reasons. First, our previous results showed that C. elegans may have a different transcriptional response to the infection by N. ausubeli than to the infections by three other Nematocida species, including N. parisii, N. major and N. homosporus (see the former chapter). Second, the sensitivity of six C.

elegans strains to N. parisii infection has been studied by Balla et al. (2015). It will be of interest to compare N. ausubeli with N. parisii in their interactions with C. elegans.

Variation in C. elegans sensitivity to N. ausubeli infection was determined by food consumption tests in the presence or absence of microsporidia. Two C. elegans strains, JU1249 and JU2825, displayed the largest sensitivity difference to N. ausubeli infection.

JU1249 was found to be the most resistant strain, whereas JU2825 was the most sensitive one.

Quantitative fluorescence in situ hybridization (FISH) analysis on microsporidian infection in the two strains suggested that the JU1249 and JU2825 strains differ in their tolerance.

Competition assays further supported that JU1249 and JU2825 are good candidates for studies on the genetic loci associated with C. elegans sensitivity variation to microsporidian infections.

A surprising effect of microsporidia infection on C. elegans was also found in this study, which will be described in detail in the next chapter.

My contributions in this study:

I performed all the assays and analyses in this part.

II.2.3. Measurements of microsporidia infection by FISH

A. FISH measurements

The infection rate, the pathogen load and the average infected area of C. elegans were measured by FISH staining. We used the FISH probe MicroB conjugated to a Cyanine5 (Cy5) dye to stain the small subunit rRNA of N. parisii and N. ausubeli (Troemel et al. 2008). FISH was performed as previously described (Troemel et al. 2008). Briefly, 5ng/µL of MicroB was used in the hybridization at 46 °C. After overnight hybridization, C. elegans populations were washed and incubated at 48 °C for 30 minutes.

Stained C. elegans isolates were mounted on glass slides and imaged using the fluorescence microscope under the Cy5 filter with a 10X objective and the Metamorph software. Images were analyzed semi-manually using ImageJ software. The body area, the infected area and the intensity of the infected area in 30 adult C. elegans were measured for each replicate. The infection rate, and the average pathogen load were computed and analyzed.

B. Statistical analysis on FISH data

Rstudio (v 0.99.903) and two packages (ggplot2, plyr) were used for the statistical analysis and the Figure 28. The pathogen load was analyzed by a linear model to explain the logarithm of fluorescence intensity (pre-normalized) by the C. elegans genotypes. To test whether the pathogen loads of the two C. elegans strains significantly differed upon each microsporidia infection, we performed a t-test. To test whether the infection rates of the two C. elegans strains significantly differed upon each microsporidia infection, we used a Wilcoxon Rank Sum test.

II.2.4. Competition assays

A. Infection and competition

The C. elegans strains JU1249 and JU2825 were competed with or without microsporidia infection. Ten uninfected JU1249 L4 larvae and ten uninfected JU2825 L4 larvae were transferred to a 10 cm NGM plate seeded with E. coli OP50. These 20 L4s founded the initial mixed population in the competition assays, with five replicates per experiment. 5 million microsporidian spores in 150 µL distilled water were placed on the E. coli lawn. The cultures were then incubated at 23 °C. The infection symptoms were checked by Nomarski optics at 48 hours after inoculation. To avoid starvation, each plate was visually checked twice a day for the amount of OP50 left in the plate; if starvation may occur before the next visual check, a small fraction of the population (about 200 to 400 animals) was transferred to a fresh culture plate. As microsporidia infection delays the growth of C. elegans populations, infected populations were transferred fewer times than uninfected populations in the experiment. After

II.3.2. Host response of JU1249 and JU2825 to infection by different microsporidia

N. ausubeli JUm2009 has a greater negative impact on JU2825 than on JU1249. We then investigated the impacts of another N. ausubeli strain JUm2526 and a N. parisii strain JUm1249 on the food consumption rates of JU1249 and JU2825. Populations with JUm2009 infection and uninfected populations were also tested in parallel. As in the previous experiment, without infection, JU1249 consumed food in a similar or a slightly slower rate, compared to that of JU2825. In all the infected populations, JU1249 was faster than JU2825 in the food consumption rate (Figure 27). These results confirmed that JU2825 is more sensitive than JU1249 to microsporidia infection.

III. The effect of microsporidia infection on Caenorhabditis elegans with the Mortal germline phenotype

Among the ten C. elegans strains used in the food consumption assay, JU2526 showed an intriguing result: the infected JU2526 population consumed food faster than the uninfected JU2526 population. We examined JU2526 and found that it displays a mortal germline phenotype (Mrt). Further studies indicated that microsporidia infection could suppress the Mrt phenotype of some C. elegans strains, i.e. extend the number of generations before reaching complete sterility.

Here, I describe my preliminary studies on this serendipitous observation.

III.1. Background

The germ line is an immortal cell lineage, which is passed down indefinitely from generation to generation. In C. elegans, the germ line can be passed over generations by self-fertilization.

The mortal germline phenotype (Mrt) corresponds to a progressive decline in fertility of the hermaphrodite from one generation to the next, until full sterility. Several C. elegans mutants (N2 genetic background), such as mrt-2, hrde-1and prg-1, have been described as Mrt, some of them showing a temperature-dependent phenotype [START_REF] Ahmed | MRT-2 checkpoint protein is required for germline immortality and telomere replication in C. elegans[END_REF][START_REF] Buckley | A nuclear Argonaute promotes multigenerational epigenetic inheritance and germline immortality[END_REF][START_REF] Simon | Reduced insulin/IGF-1 signaling restores germ cell immortality to Caenorhabditis elegans Piwi mutants[END_REF]. In our lab, we also noticed that several wild C. elegans strains exhibited the temperature-sensitive progressive sterility. At 23 °C or 25 °C, some strains with a strong Mrt phenotype become sterile after three to four generations. The molecular mechanism of the progressive sterility among wild C. elegans strains remains unknown and is being investigated by Lise Frézal in the lab.

When we investigated the sensitivity variation of different C. elegans strains to microsporidia infection, we first compared the food consumption rates of different C. elegans strains in the presence and absence of microsporidia infection. All infected strains consumed food slower than when they were uninfected, except JU2526 (Figure 30). Surprisingly, in this food consumption experiment, the uninfected JU2526 populations were only composed of few individuals, whereas the infected JU2526 had a much larger population size than the uninfected JU2526 population. Further tests showed that JU2526 has a Mrt phenotype at 23 °C, the temperature at which we did our food consumption test. As all JU2526 individuals tested in food consumption tests were from the same original population, the larger size of the infected population suggested that microsporidia infection may be able to prevent or attenuate the progressive decrease of population growth in JU2526 when cultured at 23°C.

Discussion

type strain of N. ironsii is ERTm5, which was previously identified to be a N. parisii strain based on rRNA sequence (Balla et al., 2015;[START_REF] Reinke | Identification of microsporidia host-exposed proteins reveals a repertoire of rapidly evolving proteins[END_REF]. The genome of ERTm5 was sequenced recently and compared with N. parisii ERTm1 and ERTm3. The average genome nucleotide identity between N. parisii ERTm1 and ERTm5 is 92.3%, whereas the ERTm1 and ERTm3 strains share 99.8% identical genomes (Cuomo et al., 2012;[START_REF] Reinke | Identification of microsporidia host-exposed proteins reveals a repertoire of rapidly evolving proteins[END_REF].

Based on a 95% average genome nucleotide identity standard, ERTm5 was considered as an independent species, named Nematocida ironsii [START_REF] Konstantinidis | The bacterial species definition in the genomic era[END_REF][START_REF] Reinke | Identification of microsporidia host-exposed proteins reveals a repertoire of rapidly evolving proteins[END_REF]. In genomic phylogeny, N. ironsii ERTm5 is the closest species to the clade of N.

parisii ERTm1 and ERTm3 (Figure 32). Compared to molecular analysis with a few gene regions, genomic comparisons among strains and species may provide a better and more accurate method to distinguish species and study the relationships among species. However, note that the thresholds are artificially set in both cases. in the E. longa infection, precluding exocytosis as an exit mode; E. longa spores were seen to protrude on the apical side of the host cell and push out the host cell membrane; some E. longa spores appeared to be surrounded with an additional membrane in the intestinal lumen.

The exit mode of E. longa spores is likely associated to the fact that E. longa tends to form its spores first along the apical side of host epithelial cells (paper Figures 7A,E,8L). This polarization may facilitate the exit of E. longa, as E. longa spores do not need to traffic from other places in the cell to the apical membrane of the intestinal cells, which is probably required for most N. parisii and N. ausubeli spores. Furthermore, it is likely that because E. longa does not exit in the form of SCCs through exocytosis, the host cells will not lose much intracellular membrane material. Thus no compensatory endocytosis mechanism that occurs in N. parisii infection is needed for hosts with E. longa infection, one proof of which is that no spore-containing vesicles have been observed in the E. longa infection. It would be interesting to study the molecular mechanism of the spore exit in Enteropsectra species, as well as other new rhabditid-infecting microsporidia.

I.6. The host specificity of various microsporidia

The different microsporidia displayed a range of host specificity. In the genus Nematocida, N. parisii and N. major are relatively specialists as they infect Caenorhabditis, but not Oscheius.

N. ausubeli, naturally found in C. elegans and C. briggsae, is also able to infect Oscheius sp. In the future, it would be interesting to study the specificity of N. minor, N. ciargi and the two microsporidia with broader tropisms. In the genus Nematocida, N. minor (both strains) and N.

homosporus (one of the two strains) were naturally found in O. tipulae. Different from N. The sequenced microsporidia genomes could highly facilitate studies on host-pathogen coevolution. To date, the genome of six Nematocida strains has been sequenced: the N. parisii ERTm1 and ERTm3, the N. ausubeli ERTm2 and ERTm6, the N. displodere JUm2807 and the N. ironsii ERTm5 (Cuomo et al., 2012;Bakowski et al., 2014c;Luallen et al., 2016;[START_REF] Reinke | Identification of microsporidia host-exposed proteins reveals a repertoire of rapidly evolving proteins[END_REF]. Based on the analysis of these genomes, seven Nematocida large gene families (NemLGF1-7) were defined. From these Nematocida large gene families, 82 hostexposed proteins were identified, which may function in manipulations of host metabolism and defenses. These host-exposed proteins were further found to evolve rapidly, which suggested a common strategy for microsporidia to interact and adapt to the nematode hosts [START_REF] Reinke | Identification of microsporidia host-exposed proteins reveals a repertoire of rapidly evolving proteins[END_REF]. For further studies, it would be necessary to sequence the genomes of all the new rhabditid-infecting microsporidia species, especially Enteropsectra and Pancytospora species.