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.
Microsporidia are fungi-related obligate intracellular pathogens, with over 1400 described species [1,2]. 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 [3]. 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 [4–6]. 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 [1,7]. 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 [8]. 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.) [9–11]. 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 [12–15]. With an improved understanding of the natural history of Caenorhabditis [16,17], 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 [18]. Oscheius tipulae, a very common bacteriovorous nematode species, also in family Rhabditidae, can be readily isolated from soil and rotting vegetal matter [10,19,20], as well as its closest known relative Oscheius sp. 3, with which it cannot interbreed [21]. 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 [13,14,16]. 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 [22]. Nematocida sp. 1 (described here as Nematocida ausubeli) was further isolated from a wild Caenorhabditis briggsae strain in India [22]. A microsporidian species isolated in C. elegans was found to infect the epidermis and muscles and was named Nematocida displodere [23]. Two microsporidia infecting marine nematodes have also been described, namely Sporanauta perivermis [24] and Nematocenator marisprofundi [25,26]. 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 [22]. 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 β-tubulin 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; [27]), 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.
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 [22], 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 and 2; Fig 1B). The microsporidia strain JUm2807 was isolated during these sampling efforts and described elsewhere as Nematocida displodere, and is not considered here [23]. 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. 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 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 [26] (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) [29]. Note that a recent article assigns this strain to a new species based on genome divergence [28]. 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 [22,30]. This N. sp. 1 group is most closely related to N. parisii in the microsporidian phylogeny and the sequences of both SSU and β-tubulin genes gave the same grouping (Figs 2 and 3; 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 [30]), 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 microsporidia species in our collection of 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. 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) [34]. In addition, the JUm2807 strain that has been recently described as Nematocida displodere [23] 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. As with previously isolated Nematocida, the newly identified microsporidia appeared to be transmitted horizontally, because a bleaching treatment [35] 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 [22], 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 [22], possibly derived from spore re-endocytosis [36]. In this study, as described before [22], all N. parisii and N. ausubeli infections displayed two distinct classes of spore size (Table 1; Fig 4A and 4B; 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 and 4; Fig 4C and 4D). 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 and 4F). 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 and 5L). 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 (Fig 5E–5G; 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 [22]; 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 and 5G). 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 mature spore was seen to contain a posterior vacuole on the side opposite to the anchoring disk (Fig 5K; S1H–S1J Fig). This vacuole seemed to develop from a dense membrane compartment of the sporoblast (S1C and S1D Fig). The spores displayed an external coat with several layers (Figs 5H, 5J, 5K and 6A; S1I–S1L 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 [27], 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). Whereas the Nematocida genus is in Clade II of the microsporidia [22,34], 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 [21] 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 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 Liebermannia species, with O. operophterae as outgroup (Fig 2). Based on molecular sequences, host and tissue specificity, we describe two new species: Pancytospora philotis (JUm1505 as type strain, JUm1505, JUm1670, JUm2552), found in the Oscheius gut, and P. epiphaga (JUm1396) from a C. brenneri strain from Colombia that caused an epidermis and muscle infection (Fig 7; S5 Fig). 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 (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). 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 and 7). 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). Figures of intranuclear mitosis 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. 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 (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). 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). 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 [19,20]. 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. 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 crescent-shaped 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). 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 [38]. 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 [38]. 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 [39], 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 [40]. Considering the phylogenetic relationships of the Nematocida 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.
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 [41]. (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 [16], 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 [42,43]. In addition, Nematocida species are diploid with evidence of recombination and thus possibly a sexual cycle [30,39], which might occur in another host. 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) [16]. 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 [44]. 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. 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 and 3). Overall, Enteropsectra and the Tipulae group of Oscheius species [19,21] provide an interesting case to study the evolution of a narrow host specificity. Although microsporidia are known to be able to adopt either a horizontal or a vertical transmission [42,45], 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 [23]. 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. The most striking variation we observed concerns the cellular exit strategies of the spores (Fig 6). Nematocida parisii spores acquire an additional membrane around the spore wall and thus exit through a vesicular pathway, using the host exocytosis machinery [27]; 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 [36] 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 [46,47], 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 spores pushed out and deformed the apical plasma membrane of the host intestinal cell (Fig 6; S6 Fig). 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 [48–50]. 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. 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; [29]), 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. 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.
Hundreds of samples, mostly from rotting fruits, rotting stems and compost, were collected worldwide over several years, and nematodes were isolated as described [11]. The nematode species was identified as described [11,33], 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. 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) [26,29,30,39,51] (Tables 1 and 2). ERTm4 was previously reported to correspond to N. parisii infection [29], but as no sequence data was available in GenBank, we also sequenced the SSU rDNA and β-tubulin genes for this study. 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 (DreamTaq Buffer 10X, Theromo Fisher), 1 mM MgCl2,0. 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' -CACCAGGTTGATTCTGCCTGAC-3' ) and 1492r (5' -GGTTACCTTGTTACGACTT -3' ) [52] 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' -GGGTACTGGAAATTCCGTGTT-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' -ACAAACAGGNCARTGYGGNAAYCA-3' ) and reverse primer βn1R (5' -TGCTTCAGTRAAYTCCATYTCRTCCAT-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' -CCGGACAATATCGTCTTTGG-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 [53] 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). 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 both genes) as outgroups (Figs 2 and 3; S2 Fig) [34]. 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 [34] and two Rozella species [54]. 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 [55] to estimate the best DNA evolution models and compute mean genetic distances (1000 bootstrap replicates). Bayesian inference phylogenies were constructed using Mesquite v3. 04 [56] and MrBayes v3. 2. 2 [57], with the same DNA models as above [58] and refined by FigTree v1. 4. 2 (http: //tree. bio. ed. ac. uk/software/figtree/). Worms were frozen in M9 buffer [35] 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% H2O 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 was measured as described [22]. Briefly, infected nematodes were photographed by Nomarski optics and spores were measured using the Image J software [60]. 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. For the microsporidian spore preparation, we first tried the methods previously established for N. parisii and N. ausubeli [22,51]. Because wild nematodes naturally live in habitats with various microbes [16,17], 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. 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. 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 [38]. 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. 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 105 ERTm1 (N. parisii) spores and 1. 5 x 106 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.
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.
invertebrates
medicine and health sciences
pathology and laboratory medicine
viral transmission and infection
caenorhabditis
pathogens
microbiology
parasitic diseases
animals
nematode infections
animal models
caenorhabditis elegans
model organisms
microsporidia
experimental organism systems
cellular structures and organelles
fungal pathogens
digestive system
research and analysis methods
mycology
medical microbiology
microbial pathogens
pathogenesis
cell membranes
gastrointestinal tract
host cells
anatomy
cell biology
host-pathogen interactions
virology
nematoda
biology and life sciences
organisms
A Large Collection of Novel Nematode-Infecting Microsporidia and Their Diverse Interactions with Caenorhabditis elegans and Other Related Nematodes