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(MAF, unpublished) [10] has now enabled extensive collection of natural isolates of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 <7.10−11, Fisher's exact test;
** p value<3.10−6; * p value<3.10−2.
10.1371/journal.pbio.1000586.g002
Figure 2
Transmission electron micrographs of intestinal cells of C. elegans JU1580 adult hermaphrodites.
(A,D,G) Bleached animals. (B–C, E–F, H) Naturally infected animals. (A–C) The infection
provokes a reorganization of cytoplasmic structures, most visibly the loss of intestinal
lipid storage granules (g). The cytoplasm of infected intestinal cells mostly contains
rough endoplasmic reticulum (rer) and mitochondria (m). * hole in the resin used for
inclusion in electron microscopy. (D–F) A nucleus in a non-infected animal is surrounded
by a nuclear membrane (see inset in D), whereas the nuclear membrane disappears upon
infection (E–F). Absence or incomplete nuclear membrane was observed repeatedly in
infected animals, while the nuclear membrane could be observed on bleached animals
(using both fixation methods). The nuclear material (n) in (F) may represent nucleolar
material and at lower magnification (not shown) matches the shape of elongated nucleoli
as observed by Nomarski optics (Figure 1E–G). The rough endoplasmic reticulum (rer)
on the left of the mitochondrion (m) in (F) may be a remnant of the nuclear envelope.
(G–H) The infection may result in disorganization of the intermediate filament (IF)
network normally located below the apical plasma membrane. On the right of (H) is
shown a higher magnification of the intestinal lumen, showing putative viral particles
(arrowheads). The animals were fixed using high-pressure freezing (A–C, E–F) or conventional
fixation (D, G, H).
One representative, stably infected, strain of each nematode species, C. elegans JU1580
(isolated from a rotting apple in Orsay, France) and C. briggsae JU1264 (isolated
from a snail on a rotting grape in Santeuil, France), were selected for detailed analysis.
Bleaching of adult animals resulted in phenotype-free progeny from both strains, demonstrating
that the phenotype was not vertically transmitted (embryos are resistant to the bleaching
treatment) (Figure 1K). Addition of dead infected animals, or homogenates from infected
animals after filtration through 0.2 µm filters, to plates containing previously bleached
animals recapitulated the morphological phenotype, raising the possibility that a
virus might play a role in inducing the morphological phenotype (Figure 1K). We found
that the infectious agent could be passed on horizontally through live animals by
incubating GFP-labeled animals (strain JU1894, Table 1) with 10 non-GFP-infected worms
(JU1580), checking that the latter did not die before removing them 24 h later. The
GFP-labeled culture displayed the intestinal symptoms after a week. One possibility
is that the intestinal infectious agent is shed from the intestine through the rectum
and may enter the next animal during feeding.
10.1371/journal.pbio.1000586.t001
Table 1
Strain list.
Strain
Genotype
JU1264
C. briggsae wild isolate, Santeuil, France
JU1580
C. elegans wild isolate, Orsay, France
AF16
C. briggsae wild reference isolate, India
N2
C. elegans wild reference isolate, England
CB4856
C. elegans wild isolate, Hawaii
AB1
C. elegans wild isolate, Australia
PB303
C. elegans wild isolate, USA
PB306
C. elegans wild isolate, USA
JU258
C. elegans wild isolate, Madeira
PS2025
C. elegans wild isolate, California, USA
JU1894
mfEx50[let858::GFP, myo-2::DsRed] in JU1580 background
JU1895
mfEx51[let858::GFP, myo-2::DsRed] in N2 background
WM27
rde-1(ne219) V
WM29
rde-2(ne221) I
WM49
rde-4(ne301) III
NL936
unc-32(e189) mut-7(pk204) III
In support of the hypothesis that these wild Caenorhabditis were infected by a virus,
small virus-like particles of approximately 20 nm diameter were visible by electron
microscopy of the intestinal cells (Figures 2H, S1). Such particles were not observed
in bleached animals, nor in C. elegans animals infected by bacteria, which showed
a strong reduction of intestinal cell volume (strain JU1409, unpublished data).
While a clear morphological phenotype was visible by microscopy, infection did not
cause a dramatic decrease in adult longevity (unpublished data), nor a change in brood
size (Figure S2A,B). However, progeny production was significantly slowed down during
adulthood, most clearly in the infected C. briggsae JU1264 isolate compared to the
uninfected control (Figure S2D).
Molecular Identification of Two Divergent Viruses
An unbiased high-throughput pyrosequencing approach was used to determine whether
any known or novel viruses were present in the animals. From JU1264, 28 unique sequence
reads were identified initially that shared 30%–48% amino acid sequence identity to
known viruses in the family Nodaviridae. Nodaviruses are bipartite positive strand
RNA viruses. The RNA1 segment of all previously described nodaviruses is ∼3.1 kb and
encodes ORF A, the viral RNA-dependent RNA polymerase. Some nodaviruses also encode
ORFs B1/B2 at the 3′ end of the RNA1 segment. The B1 protein is of unknown function
while the B2 protein is able to inhibit RNAi [12]. The RNA2 segment of all previously
described nodaviruses is ∼1.4 kb and possesses a single ORF encoding the viral capsid
protein. Assembly of the initial JU1264 pyrosequencing reads followed by additional
pyrosequencing, RT-PCR, 5′ RACE, and 3′ RACE yielded two final contigs, which were
confirmed by sequencing of overlapping RT-PCR amplicons. The two contigs corresponded
to the RNA1 and RNA2 segments of a novel virus. The first contig (3,628 nt) encoded
a predicted open reading frame of 982 amino acids that shared 26%–27% amino acid identity
to the RNA-dependent RNA polymerase of multiple known nodaviruses by BLAST alignment.
All known nodavirus B2 proteins overlap with the C-terminus of the RNA-dependent RNA
polymerase and are encoded in the +1 frame relative to the polymerase. No open reading
frame with these properties was predicted in the 3′ end of the RNA1 segment. The second
contig of 2,653 nt, which was presumed to be the near-complete RNA2 segment, encoded
at its 5′ end a predicted protein with ∼30% identity to known nodavirus capsid proteins
(Figure 3A). This contig was ∼1 kb larger than the RNA2 segment of all previously
described nodaviruses and appeared to encode a second ORF of 332 amino acids at the
3′ end. This second predicted ORF, named ORF δ, had no significant BLAST similarity
to any sequence in Genbank.
10.1371/journal.pbio.1000586.g003
Figure 3
Genomic organization and phylogenetic analysis of novel viruses.
(A) Schematic of genomic organization of Santeuil virus. Predicted open reading frames
are displayed in gray boxes. Red bar indicates sequence used to generate double-stranded
DNA probes for Northern blotting. Blue bar indicates sequence used to generate single-stranded
riboprobes. (B) Neighbor-joining phylogenetic analysis of the predicted RNA-dependent
RNA polymerases encoded by the RNA1 segments. (C) Neighbor-joining phylogenetic analysis
of the predicted capsid proteins encoded by the RNA2 segments.
Pyrosequencing of JU1580 demonstrated the presence of a second distinct virus that
shared the same general genomic organization as the virus detected in JU1264. Partial
genome sequences of 2,680 nucleotides of the RNA1 segment and 2,362 nucleotides of
the RNA2 segment were obtained and confirmed by RT-PCR. The putative RNA-dependent
RNA polymerases of the two viruses shared 44% amino acid identity by BLAST analysis.
Like the virus in JU1264, the virus in JU1580 was predicted to encode a capsid protein
at the 5′ end of the RNA2 segment as well as a second ORF in the 3′ half of the RNA2
segment. The ORF δ encoded proteins from the two viruses shared 37% amino acid identity
when compared using BLAST. Thus, the genomic organization of these two viruses, while
sharing substantial commonality with known nodaviruses, also displayed novel genomic
features. Phylogenetic analysis of the predicted RNA polymerase and capsid proteins
demonstrated that the virus sequences in JU1580 and JU1264 were highly divergent from
all previously described nodaviruses and most closely related to each other (Figure
3B,C). We propose that these sequences represent two novel virus species and have
tentatively named them Santeuil virus (from JU1264) and Orsay virus (from JU1580).
Viral Detection and Confirmation of Viral Infection
RT-PCR assays were used to analyze RNA extracted from JU1580, JU1264, their corresponding
bleached control strains JU1580bl and JU1264bl, and the same strains following reinfection
with viral filtrates. Orsay virus RNA could be detected by RT-PCR in the original
JU1580 culture, disappeared in the bleached strains, and stably reappeared following
re-infection with the corresponding viral filtrate (Figure 4A). The same pattern applied
for the Santeuil virus and JU1264 animals (Figure 4B). JU1580 and JU1264 cultures
continuously propagated for 6 mo by transferring a piece of agar (approx. 0.1 cm3)
to the next plate twice a week continued to yield positive RT-PCR results (unpublished
data).
10.1371/journal.pbio.1000586.g004
Figure 4
Molecular evidence of viral infection.
(A) RT-PCR detection of the Orsay virus in the original JU1580 wild isolate (I), after
bleaching (bl) and after re-infection by a 0.2 µM filtrate after 7 d (RI1) and 3 wk
(RI2) of culture at 23°C. (B) RT-PCR detection of the Santeuil virus in the original
wild isolate (I), after bleaching (bl) and after re-infection by a 0.2 µM filtrate
after 4 d (RI1) and 4 wk (RI2) at 23°C. (C) Northern blots of Santeuil virus RNA1
and RNA2 segments hybridized using a double-stranded DNA probe. (D) Northern blots
of Santeuil virus RNA1 segment using + and − sense riboprobes. (E) RNA FISH with a
probe targeting Orsay virus RNA1 segment. Representative JU1580bl animals following
infection by Orsay virus (top and middle rows) or uninfected (bottom row). S corresponds
to ovary sheath cells, OO is an oocyte, and I is an intestinal cell.
Northern blotting confirmed the presence of Orsay and Santeuil virus RNA sequences
in the infected animals. Hybridization with a DNA probe targeting the RNA1 segment
of Santeuil virus yielded multiple bands in JU1264 animals but not in the corresponding
bleached control strain. The strongest band detected migrated between 3.5 and 4 kb
consistent with the 3,628 nt sequence we generated for the putative complete RNA1
segment (Figure 4C). Multiple higher molecular weight bands were also detected that
may represent multimeric forms of the viral genomic RNAs, which have previously been
described for some nodaviruses [13],[14]. Northern blotting with a probe targeting
the RNA2 segment (Figure 4C) yielded a major band that migrated at ∼2.5 kb as well
as fainter, higher molecular weight bands. Similar patterns were seen for both segments
of Orsay virus (unpublished data).
To demonstrate virus replication in the infected animals, we performed Northern blotting
using strand-specific riboprobes. For positive sense RNA viruses like nodaviruses,
the negative sense RNA is only synthesized during active viral replication. It is
not packaged in virions and typically exists in much lower quantities than the positive
strand. Robust levels of the positive strand of the Santeuil virus RNA1 segment were
detected (Figure 4D). Northern blotting with a riboprobe designed to hybridize to
the negative sense strand detected a band of ∼3.5 kb as well as higher molecular weight
bands and a lower band of ∼1.5 kb (∼30-fold longer exposure than the positive sense
blot; Figure 4D). While the precise nature of the high and low molecular weight species
remains to be defined, the presence of multiple RNA species of negative sense polarity
in JU1264 animals demonstrates bona fide replication of Santeuil virus in JU1264.
In order to determine the localization of Orsay viral RNA in infected animals, we
performed RNA fluorescent in situ hybridization (FISH) using a probe complementary
to the positive sense RNA1 segment of Orsay virus. Viral RNA was robustly detected
in intestinal cells of JU1580bl animals infected 4 d previously with Orsay viral filtrate
(Figure 4E, top panels). Interestingly, some animals also showed localization of viral
RNA in the somatic gonad (Figure 4E, middle panels). JU1580bl animals not treated
with the viral filtrate displayed no fluorescent signal (Figure 4E, bottom panels).
High Specificity of Infection by Orsay and Santeuil Nodaviruses
We tested whether the Orsay and Santeuil viruses could cross-infect the cured wild
isolate of the other Caenorhabditis species, as well as the reference laboratory strains
of C. elegans and C. briggsae. The Orsay and Santeuil viruses could only infect strains
of C. elegans and C. briggsae, respectively (Figure 5A,B and Figure S3). Furthermore,
each virus showed intraspecific specificity of infection. Indeed, we could not detect
any replication of the Santeuil virus in C. briggsae AF16. The N2 laboratory C. elegans
strain, while infectable by Orsay virus, appeared to be more resistant to viral infection
than JU1580bl. Quantitative RT-PCR demonstrated that viral RNA accumulated in the
N2 strain at levels above background but 50–100-fold lower than in JU1580bl (Figure
5C).
10.1371/journal.pbio.1000586.g005
Figure 5
Specificity of infection by the Orsay and Santeuil viruses.
(A) Specificity of infection by the Orsay virus. Each Caenorhabditis strain (name
indicated below the gel) was mock-infected (−) or infected with a virus filtrate (+).
RT-PCR on cultures after 7 d at 23°C. See Figure S3 for corresponding morphological
symptom scoring. (B) Specificity of infection by the Santeuil virus. RT-PCR results
after 4 d at 23°C. (C) Quantitative variation in viral replication N2 versus JU1580.
N2 and JU1580 were tested by qRT-PCR for infection with Orsay virus extract (n = 10
independent replicates for each strain). By conventional RT-PCR assay, Orsay virus
infection of N2 yielded positive bands in 3 out of 10 replicate infections whereas
7 out of 10 replicate infections of JU1580bl were positive in these conditions. Control
RNA (n = 6) was extracted from JU1580bl animals grown in parallel without virus filtrate,
and to which filtrate was added at the time of sample collection. RNA levels were
normalized to ama-1 and shown as average fold-change relative to JU1580bl. Error bars
represent SEM.
Small RNA Response upon Infection
One key defense mechanism of plants and animals against RNA viruses is the small RNA
response [15]. We therefore determined by deep sequencing of small RNAs whether the
infected animals produced small RNAs in response to viral infection. We generated
small RNA libraries from mixed-stage JU1580 animals infected with the Orsay virus
and from the bleached control strain and analyzed them using Illumina/Solexa high-throughput
sequencing. These libraries represent small RNAs of 18–30 nucleotides in length independent
of their 5′ termini. Small RNAs from infected JU1580 animals that mapped to viral
RNA1 or RNA2 and had no match to the C. elegans genome are shown in Figure 6A and
6B, respectively. Of a total of 1,149,633 unambiguously mapped unique sequences, almost
2% (21,392) mapped to the two RNA segments of Orsay virus. Such RNAs were virtually
absent from a library generated from bleached JU1580 animals (<0.001%) (unpublished
data). Small RNAs that corresponded to the sense strand of the viral RNAs had a broad
length distribution and no 5′ nucleotide preference. These sense small RNAs might
represent Dicer cleavage products or other viral RNA degradation intermediates. In
contrast, most antisense small RNAs were 22 nt long and showed a bias for guanidine
as the first base (Figure 6A,B). This signature is reminiscent of a class of secondary
RNAs named 22G RNAs that are thought to be downstream effectors of exogenous and endogenous
small RNA pathways [16]–[18]. Such RNAs are not associated with transgenes expressed
in the soma of C. elegans from extrachromosomal arrays [17] nor generally a feature
associated with active transcription of endogenous genes [16]–[18]. These data suggest
that JU1580 animals raise a small RNA response to viral infection. We also detected
small RNAs of both sense and antisense polarity that mapped to the Santeuil virus
genome in the JU1264 wild C. briggsae isolate but not in bleached animals (unpublished
data).
10.1371/journal.pbio.1000586.g006
Figure 6
Small RNAs produced upon viral infection.
Number of unique sequences obtained by Illumina/Solexa high-throughput sequencing
of a 5′-independent small RNA library from JU1580 matching a given position in the
Orsay virus segment RNA1 (A) or RNA2 (B). The number of sequences in sense and antisense
orientation are shown on the positive (blue) and negative (red) y-axis, respectively.
Only sequences with a perfect and unambiguous match to the virus genome were considered.
The location of virus protein-coding genes is indicated below each graph as black
bars and the RNA genome as a line. Features of sense and antisense sequences (length
and identity of first nucleotide) are shown to the right of each graph.
RNAi Competency of the Host Is an Antiviral Defense
As viral infection appears to invoke a small RNA response in JU1580 animals, we next
tested if mutations in small RNA pathways could affect replication of the Orsay virus.
Orsay virus infection of the N2 reference strain was reduced compared to JU1580, as
assayed by viral RNA qRT-PCR (Figure 5C) and infection symptoms (Figures S3A and 7B).
Mutation of the rde-1 gene—which encodes an Argonaute protein required for the initiation
of exogenous RNAi [19]—in the N2 background increased viral RNA abundance and morphological
symptoms to levels comparable to JU1580 using both assays (Figure 7A,B). The infected
rde-1 strain produced infectious viral particles, as reinfection of the cured JU1580
strain by filtrates of infected rde-1 animals yielded positive RT-PCR results (unpublished
data). In addition, mutation of other exogenous RNAi pathway genes including rde-2,
rde-4, and mut-7 (Table 1) also led to increased viral RNA accumulation as determined
by quantitative RT-PCR (Figure 7A). We thus conclude that RNAi mechanisms provide
antiviral immunity to C. elegans and that Orsay virus infection of mutant animals
can be used to define genes important for antiviral defense.
10.1371/journal.pbio.1000586.g007
Figure 7
RNAi-deficient mutants of C. elegans can be infected by the Orsay virus.
(A) JU1580bl, N2, rde-1(ne219) (n = 10 independent replicates each), rde-2(ne221),
rde-4(ne301), and mut-7(pk204) (n = 5 independent replicates each) were tested by
qRT-PCR for infection with Orsay virus extract. RNA levels were normalized to ama-1
and shown as average fold-change relative to JU1580bl. Error bars represent SEM. Same
results as displayed in Figure 5C for N2 and JU1580. (B) Scoring of symptoms in two
independent replicates of infection of rde-1 mutant and wild-type N2 animals by the
Orsay virus filtrate, after 4 d.
Natural Variation in Somatic RNAi Efficiency in C. elegans
Since a functional RNAi pathway limits the accumulation of viral RNA in the N2 reference
strain, we assessed the exogenous RNAi competency of the bleached culture of JU1580
(JU1580bl) relative to the reference N2 strain. Using external application of dsRNAs
by feeding, JU1580bl was found to be highly resistant to RNAi of a somatically expressed
gene (unc-22) but competent for RNA inactivation of a germline-expressed gene (pos-1)
(Figure 8A,B). C. elegans wild isolates, such as CB4856, were previously known to
be variably sensitive to germline RNAi [20]. Here we thus observed for the first time
a large variation in sensitivity to somatic RNAi, which does not correlate with germline
RNAi sensitivity and thus cannot be due to inability to intake dsRNA from the intestinal
lumen. We confirmed insensitivity to somatic RNAi of the JU1580bl isolate using a
ubiquitously expressed GFP transgene (let-858::GFP), which was inactivated by GFP
RNAi in the C. elegans N2 reference background, but only modestly repressed in the
JU1580bl isolate (Figure 8C, Figure S4). We confirmed that the insensitivity to somatic
RNAi also applied when unc-22 dsRNAs were directly injected into the syncytial germline
(Figure 8D). Therefore, the robust accumulation of Orsay virus RNA observed in infected
JU1580 may be rendered possible in part by the partial defect in the somatic RNAi
pathway of this wild isolate. The accumulation of small RNAs in response to the virus
in infected JU1580 indicates, however, that its RNAi response is at least partially
active in some tissues, perhaps including the germline.
10.1371/journal.pbio.1000586.g008
Figure 8
Natural variation in somatic RNAi efficacy in C. elegans.
(A) Somatic RNAi was tested using bacteria expressing dsRNA specific for the unc-22
gene (acting in muscle; [37]). The percentage of animals with the corresponding twitcher
phenotype is shown for different C. elegans wild isolates (representative of the species'
diversity; [38]). Bar: standard error over four replicate plates. (B) Germline RNAi
was tested by feeding the animals with bacteria expressing dsRNA specific for the
pos-1 gene. The percentage of animals with the corresponding embryonic-lethal phenotype
is shown for five wild genetic backgrounds of C. elegans. Cbr-lin-12 RNAi is a negative
control. Bar: standard error over six replicate plates (too small to be seen). n>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.