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Genetic Variation in Caenorhabditis elegans Responses to Pathogenic Microbiota
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The bacterivorous nematode Caenorhabditis elegans is an important model species for understanding genetic variation of complex traits. So far, most studies involve axenic laboratory settings using Escherichia coli as the sole bacterial species. Over the past decade, however, investigations into the genetic variation of responses to pathogenic microbiota have increasingly received attention. Quantitative genetic analyses have revealed detailed insight into loci, genetic variants, and pathways in C. elegans underlying interactions with bacteria, microsporidia, and viruses. As various quantitative genetic platforms and resources like C. elegans Natural Diversity Resource (CeNDR) and Worm Quantitative Trait Loci (WormQTL) have been developed, we anticipate that expanding C. elegans research along the lines of genetic variation will be a treasure trove for opening up new insights into genetic pathways and gene functionality of microbiota interactions.
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The Natural Biotic Environment of Caenorhabditis elegans
Hinrich Schulenburg, Marie-Anne Félix (2017)
Organisms evolve in response to their natural environment. Consideration of natural ecological parameters are thus of key importance for our understanding of an organism’s biology. Curiously, the natural ecology of the model species Caenorhabditis elegans has long been neglected, even though this nematode has become one of the most intensively studied models in biological research. This lack of interest changed ∼10 yr ago. Since then, an increasing number of studies have focused on the nematode’s natural ecology. Yet many unknowns still remain. Here, we provide an overview of the currently available information on the natural environment of C. elegans. We focus on the biotic environment, which is usually less predictable and thus can create high selective constraints that are likely to have had a strong impact on C. elegans evolution. This nematode is particularly abundant in microbe-rich environments, especially rotting plant matter such as decomposing fruits and stems. In this environment, it is part of a complex interaction network, which is particularly shaped by a species-rich microbial community. These microbes can be food, part of a beneficial gut microbiome, parasites and pathogens, and possibly competitors. C. elegans is additionally confronted with predators; it interacts with vector organisms that facilitate dispersal to new habitats, and also with competitors for similar food environments, including competitors from congeneric and also the same species. Full appreciation of this nematode’s biology warrants further exploration of its natural environment and subsequent integration of this information into the well-established laboratory-based research approaches.
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Natural and Experimental Infection of Caenorhabditis Nematodes by Novel Viruses Related to Nodaviruses
Marie-Anne Félix, Alyson Ashe, Joséphine Piffaretti … (2011)
Introduction Model organisms such as D. melanogaster [1],[2] and C. elegans [3],[4] have been increasingly used in recent years to examine features of the host immune system and host-pathogen co-evolution mechanisms, due to the genetic tractability and ease of manipulation of these organisms. A prerequisite to fully exploit such models is the identification of an appropriate microbe capable of naturally infecting the host organism. Analysis in C. elegans of bacterial pathogens such as Pseudomonas, Salmonella, or Serratia has been highly fruitful, in some instances revealing the existence of innate immune pathways in C. elegans that are also conserved in vertebrates [3]. The recent report of natural infections of C. elegans intestinal cells by microsporidia makes it a promising model for microsporidia biology [5]. Efforts to use C. elegans to understand anti-viral innate immunity, however, have been hampered by the lack of a natural virus competent to infect and replicate in C. elegans. In the absence of a natural virus infection system, some efforts to define virus-host responses in C. elegans have been pursued using artificial methods of introducing viruses or partial virus genomes into animals [6],[7]. For example, the use of a transgenic Flock House virus RNA1 genome segment has clearly established a role for RNAi in counteracting replication of Flock House virus RNA [7] and has defined genes essential for the RNAi response [8]. However, this experimental system can only examine replication of the viral RNA and is fundamentally unable to address the host response to other critical aspects of the virus life cycle such as virus entry, virion assembly, or egress. The ability of a host to target steps other than genome replication to control viral infections is highlighted by recent discoveries such as the identification of tetherin, which plays a critical role at the stage of viral egress by blocking the release of fully assembled HIV virions from infected human cells [9]. Furthermore, the artificial systems used to date for analysis of virus-nematode interactions cannot be used to examine transmission dynamics of virus infection. These limitations underscore the need to establish an authentic viral infection and replication system in nematodes. Natural populations of C. elegans have proven hard to find until recent years. The identification of C. elegans habitats and the development of simple isolation methods (MAF, unpublished) [10] has now enabled extensive collection of natural isolates of C. elegans. Here we report the discovery of natural populations of C. elegans and of its close relative C. briggsae that display abnormal morphologies of intestinal cells. These abnormal phenotypes can be maintained in permanent culture for several months, without detectable microsporidial or bacterial infection. We show that these populations are infected by two distinct viruses, one specific for C. elegans (Orsay virus), one for C. briggsae (Santeuil virus). These viruses resemble viruses in the Nodaviridae family, with a small, bipartite, RNA(+sense) genome. Infection by each virus is transmitted horizontally. In both nematode species, we find intraspecific variation in sensitivity to the species-specific virus. We further show that infected worms mount a small RNA response and that RNAi mechanisms act as antiviral immunity in nematodes. Finally, we demonstrate that the C. elegans isolate from which Orsay virus was isolated is incapable of mounting an effective RNAi response in somatic cells. We thus find natural variation in host antiviral defenses. Critically, these results establish the first experimental viral infection system in C. elegans suitable for probing all facets of the host antiviral response. Results Natural Viral Infections of C. briggsae and C. elegans From surveys of wild nematodes from rotting fruit in different regions of France, multiple Caenorhabditis strains were isolated that displayed a similar unusual morphology of the intestinal cells and no visible pathogen by optical microscopy. Intestinal cell structures such as storage granules disappeared (Figures 1A–J, 2A–C) and the cytoplasm lost viscosity and became fluid (Figure 1B,I), moving extensively during movement of the animal. The intestinal apical border showed extensive convolutions and intermediate filament disorganization (Figures 1A, 2H, as described in some intermediate filament mutants, [11]). Multi-membrane structures were sometimes apparent in the cytoplasm (Figure 1C). Elongation of nuclei and nucleoli, and nuclear degeneration, were observed using Nomarski optics, live Hoechst 33342 staining, and electron microscopy (Figures 1E–H, 2D–F). Finally, some intestinal cells fused together (Figure 1I). This suite of symptoms was first noticed during sampling of C. briggsae. Indeed, more individuals appeared affected in C. briggsae than in C. elegans cultures, and to a greater extent (Figure 1K). 10.1371/journal.pbio.1000586.g001 Figure 1 Intestinal cell infection phenotypes in wild Caenorhabditis isolates. (A–H) C. briggsae JU1264 and (I,J) C. elegans JU1580 observed by Nomarski microscopy. (A–C, E–G, I) Infected adult hermaphrodites from the original cultures, with the diverse infection symptoms: convoluted apical intestinal border (A), degeneration of intestinal cell structures and liquefaction of the cytoplasm (B, G, I), presence of multi-membrane bodies (C). The animals in (E–H) were also observed in the fluorescence microscope after live Hoechst 33342 staining of the nuclei, showing the elongation and degeneration of nuclei (E′–H′). In (E), the nucleus and nucleolus are abnormally elongated. In (F), the nuclear membrane is no longer visible by Nomarski optics. In (G), the cell cytoplasmic structures are highly abnormal (apparent vacuolisation) and the nucleus is very reduced in size. In (E–H′), arrows denote nuclei and arrowheads nucleoli. The infected animal in (I) displays an abnormally large intestinal cell that is probably the result of cell fusions, with degeneration of cellular structures including nuclei. (D, H, J) Uninfected (bleached) adults. Arrowheads in (J) indicate antero-posterior boundaries between intestinal cells, each of which generally contains two nuclei. Bars: 10 µm. (K) Proportion of worms showing the indicated cumulative number of morphological infection symptoms in at least one intestinal cell, in the original wild isolate (I), after bleaching (bl) and after re-infection by a 0.2 µM filtrate (RI). Note that not all symptoms shown in (A–I) were scored, because some are difficult to score or may also occur in healthy animals. The animals were scored 4 d after re-infection for C. briggsae JU1264, and 7 d after re-infection for C. elegans JU1580, at 23°C. The symptoms are similar in both species, and generally more frequent in JU1264. *** p value on number of worms showing infection symptoms 450 observed individuals for each treatment. (C) Somatic RNAi was tested using bacteria expressing dsRNA specific for GFP. Each point corresponds to the median log2(GFP/DsRed) intensity ratio from one flow cytometry run of strains carrying the let-858::GFP transgene in the JU1580 and N2 backgrounds, after treatment with GFP RNAi or empty vector. Horizontal bars indicate group means. The difference in log2 intensity ratios between GFP RNAi and empty vector is reduced in JU1580 compared to N2 (p<0.001, see Methods). (D) unc-22 dsRNA was administered by injection into the syncytial germline of the mother. 10–14 animals of each genotype were injected and 30 progeny were scored for the twitcher phenotype on each plate. (E) Orsay virus sensitivity of seven wild C. elegans isolates representative of the species' diversity. Morphological symptoms were scored 5 d after infection of clean cultures by the Orsay virus filtrate at 23°C. The JU1580 control was performed in duplicate. Bar: standard error on total proportion. *** p<0.001. The germline RNAi competence of JU1580 together with the presence of Orsay virus RNA1 in the somatic gonad raises the possibility that vertical transmission of viral infection could occur in a strain defective for germline RNAi. To examine this possibility, JU1580bl, N2, and rde-1 were exposed to Orsay virus filtrate. A subset of adult animals from each plate was bleached and their adult offspring collected 4 d later. No evidence for vertical transmission was observed by qRT-PCR for Orsay virus RNA in any strain (Figure S5). We further tested the efficiency of the RNAi response in six other wild C. elegans isolates representative of its worldwide diversity (Figure 8A,B,D). Our results suggest that the somatic RNAi response varies quantitatively in C. elegans and is not correlated with germline RNAi sensitivity. Under experimental conditions that yield efficient infection of JU1580bl by Orsay virus, none of the other strains yielded significant levels of morphological symptoms (Figure 8E). Only JU1580bl and JU258 were positive by RT-PCR (unpublished data). Thus, factors other than RNAi competency also contribute to the sensitivity of C. elegans to the Orsay virus. Discussion The First Viruses Infecting Caenorhabditis Here we report the first molecular description, to our knowledge, of viruses that naturally infect nematodes in the wild. The two novel viruses we identified, while clearly related to known nodaviruses, possess unique genomic features absent from all other previously described nodaviruses. These viruses may thus define a novel genus within the family Nodaviridae or may even represent prototype species of a new virus family (pending formal classification by the International Committee for the Taxonomy of Viruses). The same range of intestinal symptoms was observed in animals that were infected by the Orsay and Santeuil viruses, further suggesting that these viral infections were causing the cellular symptoms. We observed putative viral particles of the size expected for nodaviruses, and a strong RNA FISH signal in intestinal cells and the somatic gonad of infected animals demonstrating that the virus is present intracellularly. It is likely that further sampling of natural populations of Caenorhabditis will yield other viruses of this and other groups. In fact, these symptoms were seen repeatedly in C. briggsae animals sampled from different locations in France, and in one instance, a Santeuil virus variant has been identified (unpublished data). A characteristic feature of these two viruses is the presence of the novel ORF δ. Conservation of sequence length and identity of the ORF δ in these two viruses, and the absence of this ORF in all other described nodaviruses, suggests that this predicted protein is likely to be important for the ability of the virus to infect or replicate in nematodes. Its function is currently unknown, but it is tempting to speculate that this protein may play a role in antagonizing an innate antiviral pathway. A Laboratory Viral Infection of a Small Model Animal The infection of C. elegans by the Orsay nodavirus provides an exciting prospect for studies in virology, host cell biology, and antiviral innate immunity. Genetic screens to identify anti-viral factors in model organisms have been limited in large part by the lack of natural infection systems. Although Drosophila has been used with great success to examine host-virus interactions for various insect viruses [21] and influenza [1], none of these studies has examined viral infection of the host organism by natural transmission routes. Here we present a novel association between C. elegans and a virus that persists in culture through horizontal transmission, causing high damage in intestinal cells yet remarkably little effect on the animal, which continues moving, eating, and producing progeny, although at a lower rate. De novo infection of naïve animals can be affected by the simple addition of either dead infected animals or homogenized lysates made from infected animals to culture dishes. This is sufficient to seed sustained complete cycles of viral replication, shedding, and infection. With this system, it is now possible to embark on whole genome genetic screens to identify host factors that block any facet of the viral life cycle. Using the current experimental conditions, infection of JU1580bl and rde-1 mutants in N2 background was highly reproducible. The fact that the reference wild type N2 strain may only sustain a very low yet detectable viral titer makes it a particularly favorable genetic background in which to screen for genes involved in interaction with the virus. The intestine is a tissue that is particularly exposed to microbes through ingestion, and is a main entry point for pathogens in C. elegans as in other animals. In C. elegans, the intestinal cells are large and easily amenable to observations by optical microscopy. The viral parasites affect the organization of the polarized epithelial intestinal cells and will likely provide interesting mechanisms and tools to study their cell biology. Clear reorganization occurs in the intermediate filaments that line the apical brush border, as well as in the lipid storage granules, the nuclear membrane, and other intracellular compartments. The abnormal state of the intestinal cells may slow down progeny production by decreasing the food intake. Alternatively, the presence of viral RNA in the somatic gonad may explain the delay in progeny production, although no gonadal cellular phenotypes have been observed. The presence of viral RNA in the somatic gonad is particularly interesting given the lack of vertical transmission. Targeted Mutant Screens with Orsay Virus Confirm a Role for RNAi in Antiviral Defense Although prior studies have clearly demonstrated a role for C. elegans RNAi in counteracting viral infection, these studies utilized either a transgenic system of viral RNA expression [7] or primary culture cells [22],[23]. The observed susceptibility of Orsay virus RNA to RNAi processing in JU1580 animals provides the first evidence in a completely natural setting, without any artificial manipulations, that RNAi serves an antiviral role in nematodes. Coupled to the increase in accumulation of Orsay virus RNA in RNAi pathway mutant strains as compared to wild type N2, these studies demonstrate that the RNAi pathway is an important antiviral defense against Orsay virus. Moreover, these results demonstrate the feasibility of identifying antiviral genes or pathways in this experimental infection system. The mechanism by which the animals prevent transmission to their offspring is unclear, but our initial results with rde-1 mutants suggest that perturbing germline RNAi is not sufficient to enable vertical transmission. Evolution of Viral Sensitivity and Specificity in Natural Populations The quantitative difference in Orsay nodavirus sensitivity between the N2 and JU1580 wild C. elegans genetic backgrounds will allow the identification of a set of host genes that modulate viral sensitivity during evolution of natural host populations. Based on the defect in exogenous RNAi of the JU1580 strain, we speculate that this set will include, but is unlikely to be limited to, genes involved in exogenous RNAi pathways. Support for the role of other genes outside the RNAi pathway comes from our data on natural isolates. Despite the fact that the magnitude of the somatic RNAi defect of the natural isolate PS2025 was comparable to that of JU1580, no evidence of viral RNA accumulation or morphological symptoms was observed following addition of Orsay virus filtrate. Whether PS2025 lacks one or more crucial receptors for viral infection or has alternative antiviral pathways that suppress viral replication is currently unknown. In addition, the Orsay and Santeuil viruses appear to specifically infect C. elegans and C. briggsae, respectively. Moreover, the C. elegans rde-1 mutation in the N2 background confers susceptibility to the Orsay virus, but not to the Santeuil virus (Figure S3C). The two viruses thus provide a system to study host-parasite specificity and its evolution. With the isolation of additional variants of each virus (our unpublished data), viral evolution studies can also be undertaken. Host-parasite evolutionary and ecological interactions can thus be explored at two evolutionary scales, within and between species of both host and parasite. The rapid life cycle of C. elegans also allows experimental evolution in the laboratory [24],[25]. This model system, which can include both natural and engineered variants of both virus and host, is thus favorable for combining studies of host-pathogen co-evolution in the laboratory and in natural populations. Materials and Methods Nematode Field Isolation Caenorhabditis nematodes were isolated on C. elegans culture plates seeded with E. coli strain OP50 using the procedures described in [10]. JU1264 was isolated from a snail collected on rotting grapes in Santeuil (Val d'Oise, France) on 14 Oct 2007. JU1580 was isolated from a rotting apple sampled in Orsay (Essonne, France) on 6 Oct 2008. When required, cultures were cleared of natural bacterial contamination by frequent passaging of the animals and/or antibiotic treatment (LB plates with 50 µg/ml tetracycline, ampicilline, or kanamycine for 1 h). Infected cultures were kept frozen at −80°C and in liquid N2 as described in [26]. Bleaching was performed as in [26]. Light Microscopy When observed with a transillumination dissecting microscope, infected animals displayed a paler intestine than healthy worms. This lack of intestinal coloration occurred all along the entire intestinal tract in C. briggsae JU1264 and preferentially in the anterior intestinal tract in C. elegans JU1580. Intestinal cells were observed with Nomarski optics with a 63× or 100× objective. The four symptoms used for scoring were 1, the disappearance of gut granules in at least part of a cell; 2, degeneration of the nucleus including a very elongated nuclear or nucleolus (when the rest of the nucleus has degenerated) or the apparent disappearance of the nucleus; 3, the loss of cytoplasmic viscosity visible as a very fluid flow of cytosol within the cell; and 4, the fusion of intestinal cells. Some of these traits may sometimes appear in uninfected animals. We systematically tested for a significant increase after infection of the proportion of animals with symptoms (Fisher's exact test). Note that some of these symptoms can also be caused by microsporidial and bacterial infections. Thus, the diagnostic of a viral infection based on the cellular symptoms requires an otherwise clean culture. Live Hoechst 33342 Staining of Nuclei Animals were washed off a culture plate in 10 ml of ddH20, pelleted and incubated in 10 ml of 10 µg/ml Hoechst 33342 in ddH20 for 45 min with soft agitation, protecting the tube from light with an aluminum foil. The animals were then pelleted and transferred to a new culture plate seeded with E. coli OP50. After 2 h, they were mounted and observed with a fluorescence microscope. Electron Microscopy A few adults were washed in 0.2 ml of M9 solution, suspended in 2% paraformaldehyde +0.1% glutaraldehyde, and cut in two on ice under a dissecting microscope for better reagent penetration [27]. Worm pieces were then resuspended overnight in 2% OsO4 at 4°C, washed, embedded in 2% low melting point agar, dehydrated in solutions of increasing ethanol concentrations, and embedded in resin (Epon-Araldite). High-pressure freezing was performed using a Leica PACT2 high-pressure freezer [28]. Progeny Counts The time course was started by isolating single L4 larvae for C. elegans JU1580 and single L3 larvae for C. briggsae JU1264. The parent animal then transferred every day to a new plate until the end of progeny production. The plates were incubated at 20°C for 2 d and kept at 4°C until scoring. The few cases where the parent died before the end of its laying period were not included. Some progeny died as embryos in both infected and non-infected cultures (non-significant effect of treatment; unpublished data). The timing of progeny production was analyzed in R using a Generalized Linear Model using infection status, day, individual (nested in infection status), and Infection Status×Day as explanatory variables, assuming a Poisson response variable and a log link function. Individual, day and Infection Status×Day were the significant explanatory variables for both JU1264 and JU1580 (p<0.001). Infectious Filtrate Preparation and Animal Infections Nematodes were grown on 10 plates (90 mm diameter) until just starved, resuspended in 15 ml of 20 mM Tris-Cl pH 7.8, and pelleted by low-speed centrifugation (5,000 g). The supernatant was centrifuged twice at 21,000 g for 5 min (4°C) and pellets discarded. The supernatant was passed on a 0.2 µm filter. 55 mm culture plates were prepared with 2–5 young adults of N2, rde-1(ne219), or JU1580bl. At the same time (Figures 1K, 4A,B, 5, 7B, and S3), or the following day (Figures 5C, 7A), 30 µl of infectious filtrate was pipetted onto the bacterial lawn. The cultures were incubated at 20°C except otherwise indicated. When both C. elegans and C. briggsae were grown in parallel, an incubation temperature of 23°C (indicated in the figure legends) was used so that both species developed at similar speeds. Maintenance over more than 4 d after re-infection was performed by transferring a piece of agar (approx. 0.1 cm3) every 2–3 d to a new plate with food. High-Throughput Sequencing Phenol-chloroform purified DNA and RNA from infected JU1580 and JU1264 animals were subject to random PCR amplification as described [29]. The amplicons were then pyrosequenced following standard library construction on a Roche Titanium Genome Sequencer. Raw sequence reads were filtered for quality and repetitive sequences. BLASTn and BLASTx were used to identify sequences with limited similarity to known viruses in Genbank. Contigs were assembled using the Newbler assembler. To confirm the assembly, primers for RT-PCR were designed to amplify overlapping fragments of ∼1.5 kb. Amplicons were cloned and sequenced. 5′ and 3′ RACE 5′ RACE was performed according to standard protocols (Invitrogen 5′ RACE kit). 3′ RACE was performed by first adding a polyA tail using PolyA polymerase (Ambion) and then using Qiagen 1-step RT-PCR kit with gene specific primers and an oligo-dT-adapter primer. Products were cloned into pCR4 and sequenced using standard Sanger chemistry. Small RNA Sequencing 4–6 90 mm plates with 15–20 adults (JU1580 or bleached JU1580) were grown for 4 d at 20°C. Mixed stage animals from all plates were collected, pooled, and frozen at −80°C. Total RNA was extracted using the mirVana miRNA isolation kit (Ambion). Small RNAs were size selected to 18–30 bases by denaturing polyacrylamide gel fractionation. A cDNA library that did not depend on 5′-monophosphates was constructed by tobacco acid pyrophosphatase treatment using adapters recommended for Solexa sequencing as described previously [30]. Each sample was labeled with a unique four base pair barcode. cDNA was purified using the NucleoSpin Extract II kit (Macherey & Nagel). Small RNA libraries were sequenced using the Illumina/Solexa GA2 platform (Illumina, Inc., San Diego, CA). Fastq data files were processed using custom Perl scripts. Reads with missing bases or whose first four bases did not match any of the expected barcodes were excluded. Reads were trimmed by removing the first four nucleotides and any 3′ As. The obtained inserts were collapsed to unique sequences, retaining the number of reads for each sequence. Sequences in the expected size range (18–30 nucleotides) were aligned to the C. elegans genome (WS190) downloaded from the UCSC Genome Browser website (http://genome.ucsc.edu/) [31] and the JU1580 partial virus genome using the ELAND module within the Illumina Genome Analyzer Pipeline Software, v0.3.0. Figure 6 is based on unique sequences (multiple reads of the same sequence were collapsed) with perfect and unambiguous alignment to the Orsay virus genome. Small RNA sequence data were submitted to the Gene Expression Omnibus under accession number GSE21736. Neighbor-Joining Phylogenetic Analysis The predicted amino acid sequences from Orsay and Santeuil nodaviruses were aligned using ClustalW to the protein sequences of the following nodaviruses. Capsid Protein: Barfin1 flounder nervous necrosis virus NC_013459, Barfin2 flounder virus BF93Hok RNA2 NC_011064, Black beetle virus NC_002037, Boolarra virus NC_004145, Epinephelus tauvina nervous necrosis virus NC_004136, Flock house virus NC_004144, Macrobrachium rosenbergii nodavirus RNA-2 NC_005095, Nodamura virus RNA2 NC_002691, Pariacoto virus RNA2 NC_003692, Redspotted grouper nervous necrosis virus NC_008041, Striped Jack nervous necrosis virus RNA2 NC_003449, Tiger puffer nervous necrosis virus NC_013461, Wuhan nodavirus ABB71128.1, and American nodavirus ACU32796.1. 1,000 bootstrap replicates were performed. RNA Polymerase: Barfin flounder nervous necrosis virus YP_003288756.1, Barfin flounder virus BF93Hok YP_002019751.1, Black beetle virus YP_053043, Boolarra virus NP_689439, Epinephelus tauvina nervous necrosis virus NP_689433.1, Flock house virus NP_689444.1, Nodamura virus NP_077730, Pariacoto virus NP_620109.1, Redspotted grouper nervous necrosis virus YP_611155.1, Striped Jack nervous necrosis virus NP_599247.1, Tiger puffer nervous necrosis virus YP_003288759.1, Macrobrachium rosenbergii nodavirus NP_919036.1, Wuhan_Nodavirus AAY27743, and American nodavirus SW-2009a ACU32794.1. 1,000 bootstrap replicates were performed. RT-PCR Nematodes from two culture plates were resuspended in M9 and then washed three times in 10 ml M9. RNA was extracted using Trizol (Invitrogen) (5–10 vol∶vol of pelleted worms) and resuspended in 20 µl in RNAse-free ddH2O. 5 µg of RNA were reverse transcribed using SuperscriptIII (Invitrogen) in a 20 µl volume. 5 µl were used for PCR in a 20 µl volume (annealing temperature 60°C, 35 cycles). For the Orsay nodavirus, the reverse transcription used the GW195 primer (5′ GACGCTTCCAAGATTGGTATTGGT) and the PCR oTB3 (5′ CGGATTCTCGACATAGTCG) and oTB4 (5′GTAGGCGAGGAAGGAGATG). For the Santeuil nodavirus, reverse transcription used oTB6RT (5′ GGTTCTGGTGGTGATGGTG) and PCR oTB5 (5′ GCGGATGTTCTTCACGGAC) and oTB6 (5′ GTCAGTAGCGGACCAGATG). One-Step RT-PCR Animals from one 55 mm culture plate plus viral filtrate (see infection procedure) were washed twice in M9. RNA was extracted using 1 ml Trizol (Invitrogen) and resuspended in 10 µl DEPC-treated H2O. 0.1 µl was used for RT-PCR using the OneStep RT-PCR Kit (Qiagen). Primers annealed to viral RNA1 (GW194 and GW195). qRT-PCR cDNA was generated from 1 µg total RNA with random primers using Superscript III (Invitrogen). cDNA was diluted to 1∶100 for qRT-PCR analysis. qRT-PCR was performed using either QuantiTect SYBR Green PCR (Qiagen) or ABsolute Blue SYBR Green ROX (Thermo Scientific). The amplification was performed on a 7300 Real Time PCR System (Applied Biosystems). Each sample was normalized to ama-1, and then viral RNA1 (primers GW194: 5′ ACC TCA CAA CTG CCA TCT ACA and GW195: 5′ GAC GCT TCC AAG ATT GGT ATT GGT) levels were compared to those present in re-infected bleached JU1580 animals. Northern Blotting For Northern blots, 0.5 µg of total RNA extracted from JU1264 and JU1264bl animals were electrophoresed through 1.0% denaturing formaldehyde-MOPS agarose gels. RNA was transferred to Hybond nylon membranes and then subject to UV cross-linking followed by baking at 75°C for 20 min. Double stranded DNA probes targeting the RNA1 segment of Santeuil nodavirus (nt 1141–1634) and the RNA2 segment of Santeuil nodavirus (nt 1833–2308) were generated by random priming in the presence of α-32P dATP using the Decaprime kit (Ambion). Blots were hybridized for 4 h at 65°C in Rapid hyb buffer (GE Healthcare) and washed in 2XSSC/0.1%SDS 5 min×2 at 25°C, 1XSSC/0.1%SDS 10 min×2 at 25°C, 0.1XSSC/0.1%SDS 5 min×4 at 25°C, and 0.1XSSC/0.1%SDS 15 min×2 at 42°C and 0.1XSSC/0.1%SDS 15 min×1 at 68°C. For strand specific riboprobes, 32P labeled RNA was generated by in vitro transcription with either T7 or T3 RNA polymerase (Ambion) in the presence of α-32P UTP. The target plasmid contained a cloned region of the Santeuil nodavirus RNA1 segment (nt 523–1022) and was linearized with either PmeI or NotI, respectively. For the riboprobes, blots were hybridized at 70°C and then sequentially washed as follows: 2XSSC/0.1%SDS 5 min×2 at 68°C, 1XSSC/0.1%SDS 10 min×2 at 68°C, 0.1XSSC/0.1%SDS 10 min×2 at 68°C, and 0.1XSSC/0.1%SDS 20 min×1 at 73°C. The Santeuil RNA1 segment migrates at approximately the same position as the 28S ribosomal RNA. Under the extended exposure time (72 h) needed to visualize the negative sense genome, low levels of non-specific binding to the 28S RNA become apparent (Figure 4D). RNA Interference For pos-1 and unc-22 RNAi using bacteria as the dsRNA source, bacterial clones from the Ahringer library expressing dsRNAs [32] (available through MRC Geneservice) were used to feed C. elegans on agar plates. For the pos-1 experiment, bacteria were concentrated 10-fold by centrifugation prior to seeding the plates. A C. briggsae Cbr-lin-12 fragment [33] was used as a negative control as it does not match any sequence in C. elegans. Three or four L4s were deposited on an RNAi plate, singly transferred the next day to a second RNAi plate, and their progeny scored after 2 d (pos-1) or 3 d (unc-22) at 23°C. For unc-22 dsRNA synthesis and injection, the unc-22 fragment in the Ahringer library clone was amplified by PCR using the T7 primer and in vitro transcribed with the T7 polymerase using the Ambion MEGAscript kit, according to the manufacturer's protocol [34]. Cel-unc-22 dsRNAs were injected at 50 ng/µl into both gonadal arms of young hermaphrodite adults of the relevant strain. The animals were incubated at 20°C. The adults were transferred to a new plate individually on the next day, and the proportion of twitching progeny scored 3 d later, touching each animal with a platinum-wire pick to induce movement. For GFP RNAi, transgenic N2 and JU1580 strains were generated expressing the ubiquitously expressed let-858::GFP and the pharyngeal marker myo-2::DsRed as an extrachromosomal array. Bacteria expressing dsRNA against GFP cDNA were used to feed animals on agar plates. An empty vector was used as a negative control. Two or three L4s were deposited on a 55 mm RNAi plate, grown at 20°C for 3 d, and the GFP/DsRed expression levels in their offspring measured using flow cytometry (Union Biometrica) as described previously [35]. Offspring from two RNAi plates were combined for sorting. Each combination of RNAi vector and strain was repeated in at least triplicate. GFP and DsRed intensities were obtained from 14 wormsorter runs including 3–4 replicate runs for N2 and JU1580 after treatment with GFP RNAi or empty vector. A larger proportion of N2 animals showed reporter expression compared to JU1580 animals (Figure S4, top). To control for this difference between strains, animals with no reporter expression were excluded by requiring DsRed intensities to exceed a cutoff set to the median 99th percentile from three control runs of animals with no array present (Figure S4). A linear regression model was fitted to the median log2(GFP/DsRed) intensity ratios including strain, treatment, and an interaction term as explanatory variables. The interaction term was significantly different from zero at p<0.001. RNA Fluorescent In Situ Hybridization (FISH) A segment of Orsay virus RNA1 was generated with primers GW194 and GW195 and cloned into pGEM-T Easy (Promega). Fluorescein labeled probe was generated from linearized plasmid using the Fluorescein RNA Labeling Mix (Roche) and MEGAscript SP6 transcription (Ambion). JU1580bl animals were infected with Orsay virus filtrate and grown for 4 d at 20°C on 90 mm plates. Control animals were grown under the same conditions in the absence of virus. In situ hybridization was performed essentially as previously described [36]. The fluorescent RNA probe was visualized directly on an Olympus FV1000 Upright microscope. Genbank Sequences: Accession numbers for Orsay and Santeuil virus contigs: HM030970–HM030973. Small RNA sequencing data at GEO: GSE21736. Supporting Information Figure S1 Putative viral particles in transmission electron microscopy of intestinal cells of infected C. elegans JU1580 adult hermaphrodites. On the right are shown higher magnifications of the parts delimited by a black rectangle in the left micrograph, showing putative viral particles (arrows). (A) Putative viral particles are found in an intracellular multi-membrane compartment. The particles in the upper left of the inset of (A) are ribosomes; the putative viral particles in the lower two-thirds are characterized by a slightly larger and more regular ring appearance (ca. 20 nm diameter). (B) Putative viral particles similar to those in Figure 2H are visible in the intestinal lumen, close to microvilli. These particles are clearly larger and distinct from the ribosomes (r) seen on the lower left of the inset. The animals were fixed using conventional fixation in (A) and high-pressure freezing in (B). (3.41 MB PPT) Click here for additional data file. Figure S2 Infection does not alter brood size, but results in delayed progeny production in C. briggsae JU1264. (A,B) Boxplots of the distribution of brood size in naturally infected and bleached (“bl”) cultures of C. elegans JU1580 (A) and C. briggsae JU1264 (B) at 20°C. The line indicates the median, the box the lower and upper quartiles, and the whiskers the 10th and 90th percentiles. Brood size is not significantly different in infected versus non-infected cultures (Wilcoxon test p = 1.0 for JU1264, p = 0.81 for JU1580). (C,D) Progeny number over time in JU1580 (C) and JU1264 (D). Infection results in a significant change in the timing of progeny production (Generalized linear model, Treatment×Day: p<0.001 in both cases). Note that time 0 corresponds to the L4 stage in the experiment in (A,C) and the L3 stage in (B,D). (0.33 MB PDF) Click here for additional data file. Figure S3 Scoring of morphological symptoms after exposure of various wild isolates and rde-1 mutants. (A) Specificity of infection by the Orsay nodavirus. Each Caenorhabditis strain was mock-infected (−) or infected with a virus filtrate (+). The proportion of worms with morphological infection symptoms after 7 d at 23°C is shown for the same experiment as in Figure 5A. (B) Specificity of infection by the Santeuil nodavirus. The proportion of worms with morphological infection symptoms after 4 d at 23°C is shown for the same experiment as in Figure 5B. (C) Santeuil virus sensitivity of rde-1 mutants in the C. elegans N2 background. Morphological symptoms were scored 5 d after infection at 23°C by the Santeuil virus filtrate. * p<0.05; *** p<0.001. (0.29 MB PDF) Click here for additional data file. Figure S4 Quantification of GFP transgene expression and silencing. (Top) Reporter genes are expressed in a larger proportion of animals from N2 compared to JU1580. Shown are the number of animals according to binned log2 DsRed intensities. Each line corresponds to one flow cytometry run with colors indicating strain and treatment as explained in the color legend. The extrachromosomal array was inherited more efficiently in N2 than in JU1580, making it necessary to analyze only those animals carrying the array. (Bottom) Each data point corresponds to the difference between treatment with GFP RNAi and empty vector in N2 (blue) and JU1580 (red) observed for animals with log2 DsRed intensity in a given bin. Dots indicate the difference between the means of median log2(GFP/DsRed) ratios for treatment with GFP RNAi and empty vector. Vertical bars indicate standard errors. The cutoff for DsRed intensities is indicated in both panels by a vertical dotted line. (0.29 MB AI) Click here for additional data file. Figure S5 Quantitative analysis of vertical transmission of Orsay virus. N2, JU1580bl, and rde-1 (5 replicates each) were infected with Orsay virus. After 4 d 25 adults from each plate were bleached onto a new, uninfected plate. The remaining adults were collected for RNA extraction. The offspring from the bleached adults were collected for RNA extraction after 4 d. As a control, virus was added to plates and incubated in the absence of animals for 4 d. Viral RNA levels were determined by qRT-PCR and normalized to gapdh. Viral RNA level is shown on a log scale, using a reference value of 1 for the infection of JU1580. (0.27 MB AI) Click here for additional data file.
- Record: found
- Abstract: found
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Caenorhabditis elegans responses to bacteria from its natural habitats.
Buck S. Samuel, Holli Rowedder, Christian Braendle … (2016)
Most Caenorhabditis elegans studies have used laboratory Escherichia coli as diet and microbial environment. Here we characterize bacteria of C. elegans' natural habitats of rotting fruits and vegetation to provide greater context for its physiological responses. By the use of 16S ribosomal DNA (rDNA)-based sequencing, we identified a large variety of bacteria in C. elegans habitats, with phyla Proteobacteria, Bacteroidetes, Firmicutes, and Actinobacteria being most abundant. From laboratory assays using isolated natural bacteria, C. elegans is able to forage on most bacteria (robust growth on ∼80% of >550 isolates), although ∼20% also impaired growth and arrested and/or stressed animals. Bacterial community composition can predict wild C. elegans population states in both rotting apples and reconstructed microbiomes: alpha-Proteobacteria-rich communities promote proliferation, whereas Bacteroidetes or pathogens correlate with nonproliferating dauers. Combinatorial mixtures of detrimental and beneficial bacteria indicate that bacterial influence is not simply nutritional. Together, these studies provide a foundation for interrogating how bacteria naturally influence C. elegans physiology.
Author and article information
Journal
Journal ID (nlm-ta): Microorganisms
Journal ID (iso-abbrev): Microorganisms
Journal ID (publisher-id): microorganisms
Title:
Microorganisms
Publisher:
MDPI
ISSN
(Electronic):
2076-2607
Publication date
(Electronic):
24
April
2020
Publication date Collection:
April
2020
Volume: 8
Issue: 4
Electronic Location Identifier: 618
Affiliations
Author notes
Author information
Yuqing Huang
https://orcid.org/0000-0002-4469-6746
Jan E. Kammenga
https://orcid.org/0000-0003-4822-4436
Article
Publisher ID: microorganisms-08-00618DOI: 10.3390/microorganisms8040618
PMC ID: 7232262
PubMed ID: 32344661
SO-VID: a6d101ca-dad1-4494-b568-5cdd9133df10
Copyright © © 2020 by the authors.
License:
Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license ( http://creativecommons.org/licenses/by/4.0/).
History
Date
received
: 01
April
2020
Date
accepted
: 22
April
2020
Categories
Subject:
Review
Data availability: