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Open Access

Peer-reviewed

Research Article

Hunting Drosophila viruses from wild populations: A novel isolation approach and characterisation of viruses

Abstract

Metagenomic studies have demonstrated that viruses are extremely diverse and abundant in insects, but the difficulty of isolating them means little is known about the biology of these newly discovered viruses. To overcome this challenge in Drosophila, we created a cell line that was more permissive to infection and detected novel viruses by the presence of double-stranded RNA. We demonstrate the utility of these tools by isolating La Jolla virus (LJV) and Newfield virus (NFV) from several wild Drosophila populations. These viruses have different potential host ranges, with distinct abilities to replicate in five Drosophila species. Similarly, in some species they cause high mortality and in others they are comparatively benign. In three species, NFV but not LJV caused large declines in female fecundity. This sterilization effect was associated with differences in tissue tropism, as NFV but not LJV was able to infect Drosophila melanogaster follicular epithelium and induce follicular degeneration in the ovary. We saw a similar effect in the invasive pest of fruit crops Drosophila suzukii, where oral infection with NFV caused reductions in the fecundity, suggesting it has potential as a biocontrol agent. In conclusion, a simple protocol allowed us to isolate new viruses and demonstrate that viruses identified by metagenomics have a large effect on the fitness of the model organism D. melanogaster and related species.

Author summary

There is known to be an enormous diversity of viruses that infect insects, but very few of these have been cultured in the laboratory, meaning that we know little about their biology. To help overcome this challenge we have generated cultured insect cells that are easily infected by viruses, and used a simple test that detects a wide variety of different viruses. This allowed us to isolate La Jolla virus and Newfield virus from Drosophila fruit flies. We found that in some species of Drosophila these are harmful pathogens that can kill or sterilise the insects that they infect, including the crop pest Drosophila suzukii. Therefore, a simple technique allowed us to isolate new viruses that are likely important pathogens of these insects in nature, and these viruses have the potential to be used to control pest species.

Citation: Bruner-Montero G, Luque CM, Cesar CS, Ding SD, Day JP, Jiggins FM (2023) Hunting Drosophila viruses from wild populations: A novel isolation approach and characterisation of viruses. PLoS Pathog 19(3): e1010883. https://doi.org/10.1371/journal.ppat.1010883

Editor: Elizabeth A. McGraw, Pennsylvania State University - Main Campus: The Pennsylvania State University - University Park Campus, UNITED STATES

Received: September 15, 2022; Accepted: March 8, 2023; Published: March 30, 2023

Copyright: © 2023 Bruner-Montero et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: Viral genome sequences have been submitted to GenBank (Accession Numbers: OP263078-OP263084). The RNAseq reads have been deposited in the NCBI Sequence Read Archive (bioproject PRJNA872469, accessions SAMN30466843- SAMN30466854). The raw data and scripts to reproduce figures and statistical analyses have be submitted to the Cambridge Data Repository https://doi.org/10.17863/CAM.94460.

Funding: This work was supported by the Leverhulme Trust grant RPG-2020-236 to FJ. G.B.-M. was supported by SENACYT-IFARHU. CSC is funded by the São Paulo Research Foundation (FAPESP) grants 2019/03997-2 and 2021/13166-0. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Over recent years metagenomics has revolutionized the discovery of new viruses, revealing extraordinary viral diversity and advancing our understanding of virus evolution [1]. However, the isolation of new viruses has not kept pace with this endeavor, thanks to a lack of appropriate culture techniques, frequently resulting from the strict requirement of a living host cell for replication. This means that the impact of these viruses on the health of their hosts is unknown and their biology largely unstudied. This pattern is exemplified by research on Drosophilidae, where metagenomic approaches have allowed the discovery of over 100 viruses, including many that infect D. melanogaster [2–5]. However, only a handful of these viruses have subsequently been isolated and studied in the laboratory [6–8]. Most virus research in Drosophila relies on viruses isolated decades ago [9], sometimes from other species of insect.

A striking result from metagenomic and PCR surveys has been that the prevalence of virus infection is extremely high in many natural populations of Drosophila [5,10]—in D. melanogaster most individuals in the wild are thought to be infected with one or more virus [5,10,11]. The effect of these viruses on flies is mostly unknown, but where they have been isolated these viruses commonly reduce both the survival and reproductive success of flies, suggesting they are important determinants of the fitness. Drosophila C virus (DCV), which is widely studied but rare in nature, is virulent, with infections in laboratory stocks causing the death of larvae and pupae, and flies injected with the virus dying within days [12]. Other viruses like Drosophila A virus (DAV) and the DNA viruses Drosophila innubila Nudivirus (DiNV) and Kallithea virus reduce host fecundity and kill their hosts, albeit at a slower rate than typically reported for DCV [8,13,14]. Some viruses, such as galbut virus and sigma virus, have less obvious symptoms [7,15]. However, indirect estimates have suggested that even the sigma virus causes an ~20% reduction in the fitness of infected flies in the wild [16]. The impact of viruses on host fitness is corroborated by the observation that genes involved in defending flies against viruses commonly evolve rapidly [17], and natural selection has favored genetic variants that increase virus resistance [18,19]. Together, these results demonstrate that viruses are common and sometimes virulent pathogens of D. melanogaster.

Insect pathogenic viruses are promising biological control agents. Baculoviruses, which are large double-stranded DNA viruses, are the only group that have been widely deployed commercially, mainly for the control of lepidopteran pests [20]. However, field trials have shown that entomopathogenic RNA viruses also have promise in biocontrol [21]. The large number of new viruses being discovered through metagenomics suggests that there may be many biocontrol agents awaiting discovery, with the first step being to isolate and characterize these viruses.

In this study we have developed an accessible protocol to detect and isolate a wide range of viruses of wild Drosophila populations. This is based on the development of a cell line that was more permissive to viral infection. We then follow O’Brian and colleagues in using the presence of double stranded RNA (dsRNA), which is produced by many viruses during their replication, as a tool to detect novel viruses in these cells [22]. We then demonstrate the utility of this approach by isolating two Drosophila viruses, the iflavirus La Jolla virus (LJV) and the permutotetravirus Newfield virus (NFV), which we show have distinct impacts on the fitness of several Drosophila species.

Results

A cell culture system to isolate novel viruses

RNAi is a major antiviral defence of insects. In Drosophila cells, viral double-stranded RNA (dsRNA) is processed into small RNAs, and these in turn target the RNAi machinery to degrade viral RNA with the complementary sequence [23]. When the expression of RNAi pathway genes is knocked down in Drosophila cell lines or mutated in mosquito cell lines, viruses replicate faster and reach higher titres [23,24]. Furthermore, viruses frequently produce viral suppressors of RNAi (VSRs) that make cells more permissive to viral replication. We exploited this to facilitate the isolation of viruses by creating a Drosophila cell line where RNAi was suppressed, increasing its permissiveness to infection. To achieve this, we stably transfected DL2 cells (a subline of S2 cells) with a plasmid expressing V5-tagged B2, a VSR produced by Flock House virus (FHV) that acts by binding dsRNA [23,25]. We named the resulting cell line DL2-B2. We used immunofluorescent microscopy with anti-V5 antibodies to confirm that the cells expressed the B2 protein in the cytoplasm after inducing expression with CuSO4 for 24 h (S1 Fig).

Double-stranded RNA is produced by positive-strand RNA viruses, double-strand RNA viruses and DNA viruses [26]. We therefore used a dsRNA enzyme-linked immunosorbent assay (ELISA) based on the J2 monoclonal antibody as a tool to detect novel viruses (see also [22]). To validate this approach and confirm that the DL2-B2 cells were more permissive to viral infection, cells were infected with the dicistrovirus DCV at different concentrations. Consistently more dsRNA was detected in the infected cells at 48 hours post infection (hpi) than in the uninfected cells, confirming that this virus produces dsRNA (Fig 1A). Furthermore, there was more dsRNA in the cells expressing the B2 protein, indicating that this succeeded in making the cells more permissive. This effect was significant at all but the highest and lowest concentrations of the virus (Fig 1A; Tukey’s HSD Test comparing cell types, p<0.01 at all viral doses)—the decline in dsRNA levels at the highest viral concentrations in DL2-B2 cells is likely caused by the virus causing extensive cell death. In our controls, there was no significant difference in dsRNA concentration between uninfected DL2-B2 and DL2 cells (Fig 1A; Tukey’s HSD Test: t = 0.31, d.f. = 79, p = 0.76).

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Fig 1. A cell culture system for isolating viruses.

(A-B) DL2 cells and DL2 cells expressing the B2 suppressor of RNAi (DL2-B2) were inoculated with a range of doses of DCV and FHV, and dsRNA was detected using ELISA based on J2 monoclonal antibodies 48 hpi. The plot shows mean fluorescence intensity (arbitrary units, a.u.) normalized to cell seeding density. (C) Cell density 3, 6 and 9 days post infection (dpi) with DCV in DL2 and DL2-B2 cells. Error bars represent standard errors. TCID50 was estimated using DL2 cells.

https://doi.org/10.1371/journal.ppat.1010883.g001

To confirm these results with a different virus, we repeated the experiment with FHV, an alphanodavirus. Again, expressing the B2 protein in cells led to higher dsRNA levels (Fig 1B; ANOVA cell line x TCID50: F = 21.3, d.f. = 3,24, P = 6 x 10−7). Again, there was no significant difference in dsRNA concentration between uninfected DL2-B2 and DL2 cells (Tukey’s HSD Test: t = 0.53, d.f. = 24, p = 0.60).

Many viruses also cause morphological changes to cells (cytopathic effect) and cell death, providing a second way to detect viruses. We therefore repeated the infections with DCV and counted the number of cells over a time-course. When uninfected, the density of cells increased over time, and expressing the B2 protein had no significant effect on cell growth (Fig 1C; ANOVA, main effect cell type: F = 0.01, d.f. = 1,20, p = 0.38). When infected with a low concentration of virus, the cell density declined at a considerably faster rate in cells expressing B2 (Fig 1C; ANOVA, time × cell type: F = 14.1, d.f. = 1,32, p = 0.0007). When infected with a high concentration of virus, cell numbers were strongly reduced in both cell lines, but again this effect was strongest when expressing B2 (Fig 1C; ANOVA, main effect cell type: F = 11.6, d.f. = 1,32, p = 0.002). Therefore, cell death provides another indicator of viral infection, and this effect is enhanced by expressing B2.

Isolation of novel viruses

Several species of wild-caught Drosophila were collected from populations across Europe and pooled in groups of 5–10 of the same sex and species. These were homogenized, filtered with a 0.22μm PVDF filter to remove microbial contamination, and then added to DL2-B2 cells. After 8–10 days, we inspected these samples under a microscope and used ELISA to test for the presence of dsRNA. We selected 19 samples that both exhibited clear cytopathic effect and had high dsRNA concentrations, and tested them for common Drosophila viruses using qPCR (S1 Table). Nine samples were infected with La Jolla virus (LJV), two were coinfected with the DNA virus Kallithea and Newfield virus (NFV), one was coinfected with NFV and LJV, and two with DCV. As DCV is widely used in our laboratory and is uncommon in the wild [5], these two samples were discarded. We did not detect known viruses in the remaining five samples.

As these isolates sometimes consisted of a mixture of viruses, we attempted to clone single viruses using three rounds of end-point-dilution culture. In every round we performed eight replicates of 12 serial ten-fold dilutions, and selected virus from the most dilute sample that resulted in infection. At the end of this we only detected the presence of NFV in the samples that were previously coinfected with Kallithea virus or LJV.

We selected eleven samples that showed consistent CPE and used RNA sequencing (RNA-seq) to examine whether samples were infected by previously unknown viruses. To check for viral contamination, we also sequenced RNA from the DL2-B2 cell line. Excluding 86% of the reads that were rRNA, there was a mean of 810,478 reads per library. When mapping to known viruses from Drosophila flies or cell culture, reads mapped only to LJV, NFV or a virus called Drosophila melanogaster American nodavirus (ANV) [27]. ANV was first detected in S2 cells [27], and has since been shown to be widespread in Drosophila cell culture [5]. As it occurred in all our samples, including our control that was not inoculated with LJV or NFV, it is presumably a contaminant in the cell line. However, across the sample with LJV or NFV, only 0.09% of the reads mapping to virus were ANV, indicating that it is present at low titres compared to the new viruses we isolated. We also noted a low level of LJV reads in the NFV samples, but as LJV was not detected in these samples in subsequent experiments using quantitative PCR (qPCR) we attributed this to contamination during the preparation of the libraries. After assembling RNA-seq reads that did not map to the D. melanogaster genome or known Drosophila viruses, we used BLAST (Basic Local Alignment Search Tool) to identify possible viruses. Among the viral contigs with an open reading frame encoding at least 40 amino acids, the top BLAST hit was always La Jolla virus, Newfield virus, or Hubei permutotetra-like virus 3. As Hubei permutotetra-like virus 3 is a close relative of Newfield virus [28], we concluded there was no evidence of previously unknown viruses in our samples. We note that this approach will only detect abundant viruses where there are sufficient reads to efficiently assemble.

To obtain genome sequences of the newly isolated viruses, we used our sequence reads to modify a reference genome sequence of these viruses, and these were aligned with published sequences from these viruses to reconstruct their phylogenetic relationships. NFV is a positive-sense single-stranded RNA virus with a genome size of ~4.7 kb belonging to an unclassified genus within the Permutotetraviridae family [5]. Our three isolates of NFV were all from samples of D. melanogaster collected in Cambridge, UK, and they formed a clade that was related to two isolates from D. melanogaster from Australia (Fig 2A). LJV is an abundant virus in several species of Drosophila [4,5]. It has a positive-sense single-stranded RNA genome of ~9.7 kb and belongs to an unclassified genus within the Iflaviridae family [5]. Our four isolates came from two pools of D. simulans from Cyprus, a pool of D. melanogaster from Cyprus, and a pool of D. repleta from Spain. They fell into two clades on the tree, and are closely related to sequences that were mostly obtained from D. melanogaster and D. simulans collected in Europe and America (Fig 2B).

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Fig 2. Phylogeny and morphology of LJV and NFV.

Phylogeny was reconstructed using (A) whole-genome sequences of NFV and (B) partial sequences of the polyprotein gene of LJV. Nodes are labelled with Bayesian posterior probabilities (%) and the tree is midpoint rooted. Virus isolates from this study are red, and taxa are labelled by host species (Dmel, D. melanogaster; Dsim, D. simulans) and collection location. Accession numbers are in S2 Table. Electron microscopy imaging of cells infected with (C) LJV isolate GBM-15052019-305 and (D) NFV isolate GBM-09102019-393 after negative staining. Arrows mark structures seen only in infected cells.

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We used electron microscopy to examine the morphology of LJV and NFV virions. LJV showed an icosahedral viral particle (Fig 2C), consistent with other positive-sense single-stranded RNA viruses in the Iflaviridae family and previous reports of LJV [29]. Surprisingly, NFV repeatably showed elongated and spherical-shaped structures, which is consistent with this virus having pleomorphic virions (Fig 2D). Other viruses in the Permutotetraviridae have symmetrical icosahedral virions [30], so further work is required to confirm these are virions.

Species-specific variation in susceptibility to NFV and LJV

To investigate the host range of LJV and NFV, we infected five species of Drosophila and measured viral titers after four days. The viral titers varied considerably depending on the combination of host and virus (Fig 3A; ANOVA, species × virus interaction: F = 50.0, d.f. = 4,40, p<10−14). For example, mean titres of LJV were more than 400 times greater in D. simulans than D. sechellia, despite these species being very closely related (Fig 3A). However, this pattern was reversed for NFV, where mean titres were over 30 times greater in D. sechellia than D. simulans (Fig 3A). There were similarly large differences among the other three species.

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Fig 3. Host range and virulence of LJV and NFV.

(A) Viral titre 4 dpi. Each point is a pool of 10–12 flies, and viral RNA levels are measured relative to RpL32. (B) Survival of flies after viral infection. Each line is from a mean of 98 flies. (C) Egg production after infection. Each point is the number of eggs laid of two females over 24h.

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NFV and LJV reduce lifespan

To evaluate the effect of viral infection on lifespan, we followed the survival of the five species of Drosophila after infection. With the exception of NFV in D. simulans, in all cases both NFV and LJV led to significantly increased mortality compared to uninfected flies (Fig 3B; Cox’s proportional hazards mixed effects model with Tukey’s adjustment: p<0.02 in all cases). However, the magnitude of this effect depended on the specific combination of host and virus. In D. erecta and D. sechellia, NFV caused significantly greater mortality than LJV (Fig 3B; Cox’s model with Tukey’s adjustment: p<0.002 in both cases). However, in the other three species this pattern was reversed, with LJV causing flies to die fastest (Fig 3B; Cox’s model with Tukey’s adjustment: p<0.001). In some species infection proved very virulent—for example, LJV had killed most D. simulans by 10 dpi (Fig 3B). In others, such as NFV infecting D. melanogaster, the virus caused only a modest increase in mortality. To examine whether these effects are likely caused by the virus we isolated, we also measured titres of the cell culture contaminant ANV by quantitative PCR. ANV titres were low—ten days post infection we estimated that there was 3.7 million times more LJV than ANV (t = 57.6, d.f. = 9.0, p<10−12).

There is an imperfect association between mortality rates and viral loads (comparison of Fig 3A and 3B). For example, D. simulans has very low titres of NFV and this virus did not reduce the survival of this species. However, this virus had higher titres in D. melanogaster than D. erecta, and yet caused greater mortality in the latter species.

NFV can sterilize female flies

NFV infection caused strong reductions in female fecundity (Fig 3C). Combining data across the three timepoints we investigated, NFV caused a reduction in the number of eggs laid of 87% in D. melanogaster, 94% in D. erecta, 83% in D. suzukii and 41% in D. simulans. With the exception of the 5 dpi timepoint in D. simulans, these reductions were all statistically significant (GLM, contrasts with Tukey’s correction: p< = 0.05 at every timepoint in each species). It is notable that this effect was markedly smaller in D. simulans, which is the species that had the lowest NFV titre (Fig 3A and 3C).

In three of the species, LJV caused considerably smaller reductions in fecundity than NFV—summing across timepoints, infection caused D. melanogaster to lay 8% fewer eggs, D. erecta 11% fewer, and D. suzukii 21% fewer (Fig 3C). The exception to this pattern was D. simulans, which suffered a 91% reduction in fecundity, and was completely sterile at 10 and 15 dpi (Fig 3C; GLM, contrasts with Tukey’s correction: p<0.001 at all timepoints). It is notable that LJV also causes very high mortality in D. simulans at the ages when we were measuring fecundity (Fig 3A).

NFV infects the follicular epithelium in the ovary and induces follicular degeneration

These results suggest that in some species NFV can have a specific impact on fecundity. For example, in D. melanogaster NFV causes less mortality than LJV, but causes a far greater reduction in fecundity. To investigate this further we examined the distribution of the two viruses in D. melanogaster ovaries using the monoclonal antibody J2 to detect viral dsRNA. A high level of dsRNA signal was observed in the ovaries of female flies infected with LJV and NFV (Fig 4). Interestingly, while LJV infection appears limited to the external muscular layers (epithelial sheet) that cover the ovariole, NFV infects the follicular epithelium, evidenced by the dsRNA signal detected within numerous follicle cells (Fig 4). Moreover, degenerated ovarian follicles with high levels of dsRNA signal can be observed at later stages, suggesting that oogenesis is severely affected in NFV-infected flies. Taken together, these results suggest that the viruses infect different tissues of the fly ovary producing different adverse effects.

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Fig 4. Immunostaining of ovaries showing the location of viral dsRNA.

D. melanogaster ovaries were stained 10 dpi with DAPI to visualise DNA (blue), phalloidin to visualise the actin cytoskeleton (red), and mouse monoclonal antibody J2 to detect viral dsRNA (green), and imaged with a confocal microscope. From left to right, columns show individual ovarioles of females uninfected (CTRL), and infected with LJV and NFV. Top panels are a maximum intensity projection of 6 confocal planes taken from the same ovariole. Middle and bottom panels are individual confocal planes showing a transverse section of the ovariole (middle) or a superficial section (bottom). Note the dsRNA signal is absent from the CTRL, present at the ovariole surface (LJV, yellow arrows) or within the follicular epithelium (NFV, pink arrows). Letters represent oocyte (O), nurse cell (NC), follicle (F) and follicle cells (FC). In the case of NFV infection, infected follicles fail to progress through oogenesis and degenerate completely (white arrowheads). Scale bar indicates 100 μm.

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NFV is a potential biological control agent of D. suzukii

Drosophila suzukii is a major pest of commercially grown fruit, so the finding that NFV can partially sterilize this species means it is a potential biological control agent. We therefore tested whether these results held when flies were orally infected. When D. suzukii females were fed with yeast paste containing NFV, the virus replicated rapidly and reached a high titre (Fig 5A). These results suggest that NFV could establish infections in populations of D. suzukii in crop fields.

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Fig 5. Effect of NFV on the fertility of the crop pest D. suzukii.

(A) Viral titre after oral feeding. Each point is a pool of 3–5 flies, and viral RNA levels are measured relative to RpL32. Samples where NFV did not amplify are shown as below the detection limit and these points are offset vertically. Time zero is immediately after removal from the food containing NFV. (B) Fecundity of D. suzukii orally infected with NFV. Each point is the number of eggs laid by 5 females over 24h. (C) Blueberry fruit (i) before and (ii) after an infestation with D. suzukii, which uses a serrated ovipositor that pierces (yellow triangle) the skin to deposit eggs. The larvae consume the fruit from the inside (iii), and cause the proliferation of opportunistic microorganisms such as fungi (iv, red triangle). (D) The number of adult flies that hatch from blueberry fruits after being infected orally with NFV or fed a control solution. Each point is the number of flies that emerge from two fruits.

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We next tested whether NFV reduces the fecundity of D. suzukii after the flies have been infected orally. We found that NFV caused a 51% reduction in the number of eggs laid (Fig 5B; GLM, main effect infection status: χ2 = 89, d.f. = 1, p<10−15). This effect on fecundity was constant through time (Fig 5B; GLM, infection status × time interaction: χ2 = 1, d.f. = 3, p = 0.79).

Finally, we investigated whether NFV-induced infertility could reduce the impact of D. suzukii on blueberry crops by exposing the fruit to infected and uninfected flies. As is the case in agriculture, we found that D. suzukii readily oviposit in blueberries and damage the fruit (Fig 5C). However, when blueberries were exposed to D. suzukii females that were orally infected with NFV, under half the number of flies emerged compared to fruit exposed to uninfected flies (Fig 5D; mean of 2.7 vs 6.95 flies/fruit; t = 5.18, p < 0.001). Taken together, the capacity of NFV to be orally acquired, and its subsequent effect on the number of adult flies emerging from fruit suggests it has the potential to be used as a biological control agent.

Discussion

The discovery of an enormous diversity of insect viruses using metagenomics makes their isolation a pressing priority. We used a simple protocol to isolate viruses from Drosophila, which could easily be applied to any species where cell lines exist, and in any laboratory equipped for routine molecular- and micro-biology. We made a cell line that was considerably more permissive to viral replication by suppressing anti-viral RNAi using the B2 protein. As B2 acts by binding viral RNA rather than a host factor [25], this should work in other insect species. After homogenizing wild-caught insects and adding them to these cells, we followed an approach that has been used in mosquitoes [22] and detected novel viruses from the presence of dsRNA, together with cytopathic effect in the infected cells. This allowed us to readily isolate LJV and NFV from multiple populations.

A key question is whether our protocol will allow the isolation of the wider diversity of viruses found in Drosophila and other insects. In these experiments we recurrently isolated NFV, LJV and Kallithea virus (although Kallithea virus was lost as it co-occurred with other viruses). However, metagenomic and PCR surveys suggesting that other viruses will almost certainly have been present in other samples we collected [5]. The reason we repeatedly isolated the same three viruses is presumably in part that we only selected only the cells with the strongest cytopathic effect and highest dsRNA concentration. A key question for future work will be whether relaxing these criteria we will allow us to isolate a wider diversity of viruses. Furthermore, such work could attempt to isolate viruses using the DL2 and DL2-B2 cells side-by-side, to test the extent to which using the more permissive DL2-B2 increases the rate at which viruses are isolated. We also pooled flies and then used serial dilution to isolate a single virus from the pool, potentially biasing the procedure towards high titre viruses. This could be avoided by isolating viruses from single flies.

By isolating viruses in cell culture, our samples can become contaminated by persistent viruses in those cells. We detected one such virus (ANV), although it was at much lower titres than LJV or NFV. In the future, purified virus could be obtained using density gradient centrifugation. However, we would note that when flies were infected with our virus extracts LJV and NFV reached far greater titres than ANV.

Both the viruses we isolated are positive-strand RNA viruses and belong to families of viruses that are widespread in insects. Using metagenomic sequencing and PCR screening, several studies have found that LJV is common in many wild Drosophila populations [5,7,11,31–33]. It is a member of the Iflaviridae family, all members of which infect arthropods, including mosquitoes [34,35], leafhoppers [36] and hymenopterans [37–39]. The pathology of other members of this family can range from harmful to asymptomatic [40]. Interestingly, LJV has been found in the Ethiopian honey bee (Apis mellifera simensis) [41], Australian honey bee (A. mellifera) [42], and the Asian hornet (Vespa velutina) [43], suggesting a relationship with hymenopteran viruses.

NFV is prevalent in both laboratory fly stocks and Drosophila cell cultures, but less common in wild populations [5]. Only two metagenomic studies have reported NFV in Drosophila wild populations. It belongs to the Permutotetraviridae family, which includes viruses that infect Lepidoptera [44], mosquitoes [45,46], Hymenoptera [39], beetles [47], leafhoppers [48] and thrips [5,49]. Viral sequences closely related to NFV have been detected in a mosquito cell culture and Aedes albopictus mosquito samples collected from Sarawak, Borneo [50,51].

While viral infection is known to be extremely common in natural Drosophila populations, for many viruses it is not known if these infections cause significant disease. We found that both NFV and LJV were virulent pathogens, capable of reducing both the survival and reproductive success of infected flies. This agrees with previous work on LJV in D. suzukii, which found a substantial reduction in the lifespan of infected flies. Given that LJV is estimated to infect ~9% of wild D. melanogaster, it is likely an important determinant of fitness [5].

There was a great variation in the effect of NFV and LJV viruses between species. Both viruses reduced survival. Female fertility was dramatically affected by NFV in all species except D. simulans. Conversely, LJV strongly reduced the fertility of D. simulans but had modest effects in other species. The mechanisms that explain these species-specific outcomes of host-virus interactions are unknown. However, the coevolution of hosts and pathogens can increase the natural variation of resistance, and drive differences between populations and species [52–54]. For example, the host range of the Nora virus is constrained by specific interactions between the host antiviral RNAi machinary and the viral suppressors of RNA silencing [55].

With the exception of D. simulans where titers are low, NFV had a strong sterilizing effect on females. In D. melanogaster this was associated with its ability to infect follicular epithelium in the ovaries and cause ovarian follicles to degenerate. Many parasites have specific effects on host fertility, a trait often called parasitic castration [56]. This is sometimes interpreted as an adaptation of the parasite. If a parasite kills it host, it will die too. However, if it castrates the host, this may redirect resources to the parasite without reducing the period that the host is infectious [56]. Whether the sterilizing effect of NFV is an incidental byproduct of infection or an adaptation is unknown.

In surveys of natural Drosophila populations, it is common to find that virus infect multiple species but their prevalence varies [5], and this could be caused by ecology or differences in host susceptibility is unknown. We found that the viral load varied extensively, even among closely related species. The same was true for virulence, with the impact on fecundity and survival differing greatly among host species. More extensive surveys of DCV and sigma virus have revealed similarly large differences in the susceptibility of Drosophila species to infection [57–59]. Together, these results suggest that host genetics may be a key factor shaping the community of viruses in different Drosophila species.

NFV reduced the fecundity of D. suzukii, which is an invasive pest of fruit crops around the world. It was able to infect D. suzukii orally and reduce the number of flies that emerged from blueberry fruits, suggesting it merits investigation as a potential biological control agent. In a similar experiment it was recently shown that LJV can readily infect D. suzukii orally, and causes similar reductions in lifespan to those we observed after inoculating the flies with the virus [60]. Furthermore, the virus remained infectious after exposure to a broad range of environmental conditions [60]. Other insect viruses have shown to cause mortality in D. suzukii, including Cricket Paralysis virus (CrPV), FHV, DCV and DAV [6,29,33,61,62]. While these experiments suggest viruses might be suitable as biological control agents, more research is needed to determine whether producing and delivering RNA viruses is practical, or whether their effects on D. suzukii are sufficient to reduce crop damage.

Our results provide an alternative method to isolate Drosophila viruses from wild populations by suppressing Drosophila cells RNAi antiviral defence, detection of dsRNA, CPE inhibition assay and RNA sequencing. Additionally, we achieved the isolation of several isolates of LJV and NFV. We demonstrate that the virulence of these viruses varies greatly among closely related Drosophila species and depends on the specific combination between host species and virus. Drosophila is an important genetic and molecular model for the study of host-virus interactions, that together with these new viruses, may represent a useful tool for understanding the genetic and evolutionary basis of viral diseases.

Materials and methods

Fly collection

Drosophila flies were collected in the field at Limassol and Nicosia (Cyprus) in 2019, Budapest (Hungary) in 2018, Granada and Mallorca (Spain) during 2018–2019, Cambridge (United Kingdom) during 2019. Flies were caught using an inverted cone fly trap containing banana baits sprinkled with dried yeast and ~300 μl of apple cider vinegar, or directly collected from decaying fruits using an entomological aspirator. Every 20–25 flies were transferred into a vial containing standard cornmeal diet and maintained at room temperature. Cyprus flies were transported to the Chrysoula Pitsouli’s laboratory at the University of Cyprus, where flies were anaesthetized using CO2 and split by species level under a stereomicroscope. D. melanogaster and D. simulans females are hard identify morphologically, so that the females for these species were pooled together. All vials were transported to the Department of Genetics at the University of Cambridge for further analysis.

Virus isolation

Flies were anaesthetized using CO2, labelled, and sexed under the stereomicroscope. A pool of 5–10 females or males of the same species per sample were homogenized using a sterile pestle in an Eppendorf tube (1.5 ml) containing 200 μl of sterilized Ringer solution. Then, the tube was briefly spun and supernatant (~180 μl) was placed into a PVDF filter tube (Ultrafree-MC, 0.22 μm) and centrifuged at 1000g for 3 min to remove bacterial contamination and debris. The filtered solution was stored at -80°C for future analysis. Ten μl of the filtrate was inoculated into a well (96-well flat-bottomed plate) containing 190 μl of standard Schneider medium with DL2-B2 cells and incubated at 25°C. Three inoculations were performed as technical replicates per sample. The CPE of the cells was monitored daily for 10 days. On day 10, the plate was briefly spun (1000g for 30s) and 100 μl of the supernatant was stored at -80°C. The percentage of CPE per total well area (32.16 mm2) was scored from minimum (0) to maximum (3), where 0, 1, 2, and 3 represent 0–25%, 26–50%, 51–75%, and 75–100% of CPE, respectively. The remaining 100 μl was used in an enzyme-linked immunosorbent assay (ELISA) to detect double-stranded RNA (dsRNA). Samples with the highest CPE and dsRNA concentrations were retained.

Enzyme-linked immunosorbent assay

One hundred μl of the cell culture was transferred into a 96-well microplate provided by the In-Cell ELISA kit (Abcam, ab111541), and treated following the manufacturer’s recommendations. For detecting dsRNA, the mouse monoclonal antibody J2 (Scicons) diluted 1:400 was used as primary antibody. As secondary antibody, goat anti-mouse IgG (IRDye 800CW) diluted 1:1000 was used. The plate was stored at 4°C, and subsequently scanned on the Odyssey Infrared Imaging System using a fluorescent signal intensity, both 700 and 800 nm channels, and 4 mm focus. The fluorescence intensity of the image was analyzed on LI-COR Image Studio 4.0 Software for Odyssey. The microplate was normalized using the Janus green staining protocol as recommended by manufacturer instructions. Briefly, the microplate was emptied and 50 μl of 1 × Janus Green stain was added to every well for 5 min at room temperature, then the plate was washed 5 times with deionized water. Finally, 200 μl of 0.5 M hydrochloric acid was added to every well and incubated for 10 min. The microplate OD at 595 nm was measured in a microplate spectrophotometer SpectraMax iD3 (Molecular Devices).

RNA extraction and quantitative polymerase reaction

Two ml of DL2-B2 cell culture (100,000 cell/ml) was grown in plastic culture flasks (see below, cell culture maintenance), and inoculated with 10 μl of the filtered solution from a sample positive for virus. After 8 days, the cell culture was transferred into a tube (15 ml) and briefly spun. Two hundred-fifty μl of the supernatant was used for total RNA extraction using the chloroform isopropanol TRIzol method following the manufacturer’s protocol (Life Technologies), the remaining supernatant was placed in cryogenic vials (Cryovial) and stored at -80°C. One μl of RNA per sample was reverse-transcribed with Promega GoScript reverse transcriptase, followed of random hexamer primers, and then diluted 1:10 with nuclease free water. The complementary DNA (cDNA) was screened for the presence of the 14 Drosophila viruses: La Jolla virus, Kalithea virus, Drosophila C virus, Grom virus, Thika virus, galbut virus, Vera virus, Nora virus, Motts Mill virus, Craigies Hill virus, Drosophila melanogaster sigma virus, Drosophila A virus, Newfield virus and Chaq virus. Additionally, the samples were screened for FHV and CrPv, which are occasionally found in fly stocks as contamination. For the qPCR, 1 μl of cDNA per sample was used to quantify the viral load by amplifying conserved regions of each virus genome (see primers, S1 Table), and the fly gene RPL32 was used to normalize the expression. qPCR was performed on an Applied Biosystems StepOnePlus system using Sensifast Hi-Rox Sybr kit (Bioline) with the following PCR cycle: 95°C for 2min followed by 40 cycles of: 95°C for 5 secs followed by 60°C for 30 secs. Two technical replicates per reaction were carried out per sample with both the target virus and reference genes. To estimate the relative viral load, the formula ΔΔCt = ΔCtrpl32 –ΔCttarget virus was used, where ΔCtrpl32 and ΔCttarget virus represent the mean of the technical replicates.

Purification of viral particles

To purify the viral particles, we used the end-point-dilution. The stock samples with virus detected were thawed on ice, and 10 μl of it inoculated into a well (96-well flat-bottomed plate) containing medium and DL2-B2 cells. Eight replicates of each sample were 1:10 serially diluted 12 times, and the CPE was monitored daily for 10 days. This process was repeated twice. The serially diluted samples were then grown in 6 ml of cell culture (6-well flat-bottomed plate) for 5 days.

Library preparation

One Illumina sequencing library per sample was prepared, including one library for the virus-free DL2-B2 cells used in the isolation protocol as control. One μg of total RNA was used for the NEBNext Ultra Directional RNA Library Prep Kit (New England Biolabs) without removing ribosomal RNA. cDNA synthesis was performed according to NEB protocol, and NEBNext Adaptor with hairpin loop structure were ligated to prepare for hybridization. Size selection was performed using NEBNext Sample Purification Beads (New England Biolabs), and indexes were added using NEBNext Multiplex Oligos for Illumina (New England Biolabs). The library quality was evaluated with the Agilent Bioanalyzer system. Libraries were transported to the Cancer Research UK Genomics Core Facility and sequenced on NovaSeq 6000 system (Illumina).

Bioinformatics

The raw data quality assessment and adapter removal were performed with TrimGalore. Then, sequencing reads were mapped to both the D. melanogaster genome and all published Drosophila associated-virus sequences available in the National Center for Biotechnology Information (NCBI) [47] using STAR [48]. When counting sequence reads mapping to known viruses, we mapped reads to a manually curated database [63]. Subsequently, the unmapped reads were mapped to the SILVA rRNA database [49] to remove all rRNA using bowtie2 [50]. De novo assembly of the unmapped reads was performed with TRINITY software [64], together with the utility TransDecoder to identify coding regions within the transcripts with ORFs higher than 30 amino acids. These contigs were then queried against NCBI RNA virus proteins using blastx to identify putative viruses. The virus-related contigs were confirmed by querying the non-redundant protein database RefSeq by blastx.

To obtain the genome sequence of viruses that had been isolated we mapped reads to published genomes as described above and genetic variants were called with the program Freebayes. These variants were then used to modify the published genome sequence.

Phylogenetic analysis

All sequences of NFV and LJV were downloaded from NCBI for phylogenetic analysis. Complete genome sequences of NFV (~4761 bp) and LJV partial length sequences (~690 bp) of the polyprotein gene were aligned with MUSCLE [65] using default settings. The best-fit nucleotide substitution model for the aligned sequences was determined by jModeltest [66]. Bayesian inference phylogenetic analysis was performed in MrBayes version 3.2.2 [67], with a General Time Reversible (GTR) DNA substitution model (lset nst = 6) Gamma distributed (rates = gamma), 4 independent chains (nchains = 4) and recording the tree and parameters every 1000 generations (samplefreq = 1000). Several runs were performed to reach the optimal range of convergence adjusting the temperature parameter for heating the chains (temp = 0.1). All analyses consisted of 10 million generations. Sixty-four sequences were added to the analysis from the GenBank database from previous studies for comparison.

Transmission electron microscopy

Virus samples with LJV and NFV were inoculated in DL2-B2 cell culture for 4 days. Two hundred μl of the culture was placed into a PVDF filter tube (Ultrafree-MC, 0.22 μm) and centrifuged at 1000g for 3 min. The supernatant was centrifuged twice using different filter tubes to eliminate the cell debris. Aliquots of the supernatant samples were transported to the Cambridge Advanced Imaging Centre for imaging processing. Samples were adsorbed onto glow-discharged 400 mesh copper/carbon film grids (EM Resolutions) for about 1 min. Then, transmission electron microscopy (TEM) grids were passed over two drops of deionized water to remove any buffer salts and stained in 2% (w/v) aqueous uranyl acetate for about 30 seconds. Uranyl acetate dye was drained off the TEM grid using filter paper and grids were allowed to air dry. Samples were viewed using a Tecnai G20 transmission electron microscope (FEI/ThermoFisher Scientific) run at an accelerating voltage of 200 keV using a 20-μm objective aperture to improve contrast.

Cell culture maintenance

Drosophila cultured cell lines DL2 and DL2-B2 were used in the experiments. DL2 refers to a subline of S2 cells that was supplied by Peter Christian for use in viral culture [68]. Cells were grown in plastic culture flasks containing Schneider medium supplemented with 10% heat-inactivated fetal bovine serum (FBS) and streptomycin 100 μg/ml and penicillin 100 U/ml to inhibit bacteria and fungus contamination. Culture flasks were passaged every 6 days and incubated at 25°C.

Amplification and digestion of FHV B2 open reading frame

FHV cDNA was synthesized from RNA isolated from infected flies using GoScript Reverse Transcriptase system (Promega), the resulting cDNA was diluted 2x before use in a PCR reaction. The open reading of the FHV B2 viral suppressor of RNAi [23] was amplified from FHV cDNA using FHV-B2-not_Fw (5’GCACGCGGCCGCACCATGCCAAGCAAACTCGCGCTAATC’3) and FHV_B2_not_Rv (3’GCACGCGGCCGCCCCAGTTTTGCGGGTGGGGGGTC’5) primers. FHV_B2_not_Fw incorporated a NotI restriction site and consensus Kozak sequence at the 5’ end of the open reading frame. FHV_B2_not_Rv incorporated a NotI restriction site, 2 extra bases to put the open reading frame in-frame with the V5 and His epitope tags in the vector and removed the stop codon at the 3’ end. Q5 hot start DNA polymerase (New England Biolabs) was used in the amplification reaction with a touchdown thermocycling protocol. A 5μl aliquot of the resulting 345 bp PCR product was analyzed by agarose gel electrophoresis. The remainder was purified using the QIAquick PCR Purification Kit and the B2 amplicon was eluted in 90μl of 1 mM Tris-HCl pH 8.0. The fragment was then digested by adding 10μl of 10x sure cut restriction enzyme buffer and 1.5μl (15 units) of NotI restriction enzyme (New England Biolabs) and incubating at 37°C for 1:45 h. The digested fragment was purified by gel electrophoresis and the QIAquick gel extraction kit (Qiagen) being careful to not expose the DNA to the fluorescent light for more time than was necessary and eluted in 20 μl of 1mM Tris-HCl pH 8.0.

Plasmid digestion

Meanwhile, 150 ng of pMT-puro plasmid was digested with NotI restriction enzyme in a total reaction volume of 100 μl at 37°C for 1:45 h. To prevent re-annealing of the empty plasmid during ligation the 5’ ends of the digested plasmid were dephosphorylated by adding 1ul of alkaline phosphatase and incubation at 37°C for 1 h. The digested plasmid was then purified using a PCR purification kit (Qiagen) according to the manufacturer’s recommendations and eluted in 20μl of 1 mM Tris-HCl pH 8.0.

Ligation and transformation

Plasmid and B2 insert were ligated together using a rapid ligation kit (Promega). Ligation reactions contained 2.5μl 4x buffer, 0.5μl T4 ligase, 1μl vector, 1μl insert and were incubated at room temperature for 15 min. A control ligation that contained plasmid only was also set up. The ligations were transformed into 50 μl of chemically competent sub-cloning grade E. coli DH5α strain (New England Biolabs). Cells were thawed on ice then 2μl of the ligations were added to the cells, which were then carefully mixed using gentle agitation and returned immediately to ice and incubated for 30 min. The cells were then heat shocked for 30 s at 42°C in a water bath and momentarily returned to ice. Nine hundred and fifty μl of Luria-Bertani (LB) was gently added to the cells and they were incubated with shaking at 37°C for 45 min. The cells were pelleted by centrifugation at 6000 g for 1min and all but 200 μl of the supernatant was removed. The cells were re-suspended in the remaining LB, and they were spread onto 96 mm diameter bacterial LB agar plates containing 100 μg/ml ampicillin and the plates were incubated inverted at 37°C overnight.

Analysis of clones

Next day 30 single colonies were removed using a pipette and washed off into 10 μl LB containing 100 μg/ml ampicillin. These cultures were tested for the presence and orientation of the inserts by PCR. For each culture 3 PCR reactions were set up in a 96-well PCR plate, primer pairs were as follows: FHV_B2_not_Fw/FHV_B2_not_Rv; pMT_forward/FHV_B2_not_Rv; pMT_forward/ FHV_B2_not_Fw. Thermopol DNA polymerase was used (New England Biolabs) and a touchdown PCR protocol. 0.3 μl of each culture was used as a template in a 15 μl reaction volume. Amplified fragments were analyzed by gel electrophoresis. The remainder (~9 ul) of the 5 cultures containing plasmids with B2 inserts of the correct orientation were used to inoculate 10 ml LB cultures and were incubated overnight with shaking at 37°C. The following day plasmids were purified from these cultures (QIAprep Spin Miniprep Kit, Cat No./ID: 27104) and quantified using Qubit dsDNA assay. Two hundred ng of each plasmid were digested with 0.5 μl of NotI in a 10 μl reaction volume for 1 h at 37°C and then analyzed by gel electrophoresis. All plasmids contained the correct sized insert. Three of the 5 plasmids were sequenced using pMT_forward and BGH_reverse primers. Each sequencing reaction contained 200 ng of template plasmid DNA. All 3 plasmids had the correct sequence.

DL2 cell transfection

One day grown DL2 cells were transfected using the plasmids containing the B2 insert. Transfection was performed using Effectene transfection (QIAGEN) reagent according to the manufacturer’s instructions. Transfected cells were incubated in 1.6 ml Schneider medium containing ~1.8x106 DL2 cells seeded in 6-well plastic plates. Growth medium was supplemented with antibiotics and incubated as mentioned above. Second-day post-transfection, the medium was removed and replaced with medium containing 10 μg/ml of puromycin and incubated for 3 days. Each well was monitored every 24 h to check cell conditions. To induce the expression of the B2 protein, 5 mM of CuSO4 was added 24 h before the experiments.

Immunohistochemistry

Shortly before the DL2-B2 cells were used in experiments, immunohistochemistry was performed to test the expression of the vector (S1 Fig). pMT contains the V5 epitope tag allowing its rapid detection with Anti-V5 Antibody. DL2-B2 induced and uninduced cells (CuSO4-free) were cultured under the conditions mentioned before. Briefly, 1 ml of medium was centrifuged at 2000g for 3 min, the supernatant medium was discarded and washed twice with 1% v/v phosphate-buffered saline (PBS). Then, 20 μl of the cells were placed on a slide coated with Poly-L-Lysine for 30 min. Every well was washed 3x with 1% v/v PBS, and 20 μl of 4% w/v paraformaldehyde was added for 30 min, followed by blocking and permeabilization with 1× PBS, 0.01% v/v Triton-X (Sigma), 1% v/v Normal Goat Serum (NGS) for 30 minutes. The cells were labelled with V5 tag mouse monoclonal antibody (Thermofisher) at a dilution of 1:1000 in 1% v/v PBS and incubated overnight at 4°C, then labelled with Alexa Fluor 488 rabbit anti-mouse IgG secondary antibody (Thermofisher) at a dilution of 1:1000 and incubated overnight at 4°C. Finally, cells were washed 3x with 1% PBS and stained with Hoechst for 5 min and washed 3x with PBS. The slide was mounted with 80% glycerol and edges were sealed with nail polisher. The images were taken on a Nikon Eclipse 90i microscope at 20× magnification and processed using the Fiji software.

Cell count and dsRNA after viral infection

To test how permissive the DL2-B2 cells were to viral replication, we counted the number of viable cells after infection with DCV-C. DL2-B2 or DL2 cells (100,000 cell/ml) were grown in a 96-well flat-bottomed microplate. Each well contained 90 μl of Schneider medium, puromycin 10 μg/ml and CuSO4 5 mM. After 24 h, wells were randomly inoculated with 10 μl of DCV at different concentration treatments. All protocols and cell culture incubation were performed at 25°C. Cell concentration was estimated 72-, 144-, and 216-hours post-infection (hpi) by pipetting ~20 μl of cell solution into a disposable hemocytometer (FastRead 102) under the microscope.

To quantify dsRNA in DL2-B2 and DL2 cells, cells were infected with DCV or FHV and 48 hpi, cells were transferred into a 96-well microplate provided by the In-Cell ELISA kit, and the dsRNA was detected and analyzed as mentioned above.

Drosophila species and infection of flies

Five species that belong to the subgenus Sophopora were used in the study. Flies were maintained in glass vials (~ 28.5 × 95 mm) with standard cornmeal (1200 ml water, 13 g agar, 105 g dextrose, 105 g maize, 23 g yeast, 35 ml Nipagin 10% w/v), and incubated at 25°C and ~70% humidity.

The infection experiments used LJV isolate GBM-15052019-4-305 that was isolated from Gialousa, Cyprus, and NFV isolate GBM-09102019-1-393 that was isolated from Cambridge, UK. All virus isolates were cultured in DL2-B2 cells. To produce aliquots of virus for the experiments, 2 ml of cell culture was added to a conical tube (Falcon) and briefly spun (1000g for 2 min) to remove cell’s debris, then the supernatant was aliquoted (10 μl) into sterilized 0.2 ml PCR tubes and stored at −80°C. The viral concentration of the isolates was estimated using the TCID50 ml–1 method [69]. To do this, DL2-B2 cells (100,000 cell/ml) were grown in a 96-well flat-bottomed plate containing 190 μl of standard Schneider medium supplemented as mentioned above. Then, 10 μl of the viral solution was inoculated into a well, and serially diluted 1:10 14-fold. The cytopathic effect of the cells was monitored daily for 10 days. The aliquot stocks were thawed immediately before each pricking assay on ice and diluted in sterile Ringer’s solution to standardise the concentration of the virus isolates to 1×105 TCID50 ml–1.

For the experiment measuring viral titres over a time-course after feeding to D. suzukii, additional NFV was produced as follows. 200 flies from DGRP lines 852 and 177 were anesthetized on a CO2 pad and pricked on the dorsolateral thorax with NFV (isolate GBM-09102019-1-393, concentration 1×103 TCID50 ml–1). Four days post infection, 100 flies were placed into microcentrifuge tubes and homogenized in 1 mL of sterile Ringer’s solution. The tubes were spun and the supernatant filtered using a PVDF filter (Ultrafree-MC, 0.22 μm).

Viral infection

Flies were anesthetized on a CO2 pad and then pricked on the dorsolateral thorax under a stereomicroscope using a needle (Austerlitz Insect Pin) dipped into a Ringer’s solution containing 1×105 TCID50 ml–1 viral titre. To avoid cross-contamination between viruses, different CO2 pads and needles were used for each virus. These utensils were kept in independent plastic bags and cleaned with Virkon (5% w/v) and ethanol (70% v/v) frequently. After infection, infected flies were kept in independent trays for each virus treatment to avoid cross-contamination. Unless otherwise mentioned, flies were transferred every 3 days to new vials with fresh food cornmeal and incubated at 25°C over the course of the experiments.

Oral infection

To infect orally the flies, a cohort of virgin adult females 3–5 days old was transferred into an empty vial without food containing a damp towel paper for 24 h at 25°C. The next day, the starved females were transferred into a vial containing 300 μl of yeast paste (25% w/v yeast powder, 5% vegetable red dye v/v, and 1×105 TCID50 final concentration of one of the viruses) and incubated at 25°C. Another cohort of females was transferred into vials with Ringer’s solution (25% w/v yeast powder and 5% red dye v/v) as a control. To confirm that the flies ingested the virus solution, the next day flies’ gut was checked under a stereoscopic. Flies with red-stained intestines were selected for the experiment and transferred into new vials with fresh cornmeal food.

Virus quantification

The total RNA of the homogenized flies was extracted using the chloroform isopropanol method following the manufacturer’s protocol (Life Technologies). One μl of RNA per sample was reverse-transcribed with Promega GoScript reverse transcriptase using random hexamer primers, and then diluted 1:10 with nuclease-free water. qRT–PCR was performed on an Applied Biosystems StepOnePlus system using Sensifast Hi-Rox Sybr kit (Bioline) with 1 μl of complementary DNA (cDNA) per sample was used to quantify the viral load using specific primers for LJV LaJolla1_foward (5’-CGGACCAGAGTGTAGCCAAG-3), and LaJolla1_reverse (5’-AGTGCCATCCAYCGATTTGT-3’), and NFV NewfieldVirus_2_forward (5’-TTGATGATGTCGCCACGAGA-3’), NewfieldVirus_2_reverse (5’-CATTCGCCGAGACCTCCATC-3’). The fly gene RPL32 was used to normalize the expression using primers RpL32_forward (5’-TGCTAAGCTGTCGCACAAATGG-3’) and RpL32_reverse (5’- TGCGCTTGTTCGATCCGTAAC-3’) [58]. The qRT-qPCR was performed with the following PCR cycle: 95°C for 2min followed by 40 cycles of 95°C for 5 secs followed by 60°C for 30 secs. Two technical replicates per qRT-PCR reaction were carried out per sample with both the viral and reference genes. To estimate the relative viral load, the formula ΔCt = CtRPL32–Cttarget_virus was used, where Ct is the mean Ct value of the technical replicates performed on each target sequence. For ANV, qRT–PCR was performed on QuantStudio 5 (Applied Biosystems) using specific primers for ANV AmericanNodavirus1_foward (5’-ATTGGTATGGGGCACAAGGA-3’), and AmericanNodavirus1_reverse (5’-TGACACAACTTTCTTGCCGG-3’) designed using ANV sequence [27].

Viral titre across different Drosophila species

To evaluate the effects of the viruses across different fruit flies, 5 Drosophila species were infected with LJV and NFV. For each species, two male and 2 female flies were transferred into 6 vials (n = 12 vials total), containing standard diet and incubated at 25°C—flies were discarded after 2 days. After 2 weeks, 10 adult males (3–5 days old) for each species were transferred into a vial with fresh cornmeal food. Then, 6 vials per species treatment were pricked with either LJV or NFV. Three vials were pricked with Ringer’s solution as an uninfected control. Four-days dpi, all flies per vial were anaesthetized using CO2 and transferred into Eppendorf tube (2 ml) containing beads. Every tube was chilled on ice for 10–15 min and 250 μl of TRIzol Reagent (Invitrogen) was added. Immediately, tubes were homogenised using a Qiagen TissueLyser II and stored at −80°C. For each tube, the total RNA was extracted, and the relative virus concentration was estimated as mentioned above.

Lifespan

To assess the effect of the viruses on lifespan of flies, different Drosophila species were infected with either LJV or NFV. For each species, 12 vials with flies (2 males and 2 females) were set up and the parental flies discarded as in the experiment above. After 2 weeks, 20–25 adult males (3–5 days old) per species were transferred into a vial with fresh cornmeal food. Four vials per species were pricked with either LJV, NFV or Ringer’s solution (control). The number of dead flies per vial was monitored daily until the last fly was dead. Mortality on day 1 was attributed to the damage induced by the needle, and the data was discarded from the analysis.

Fecundity

To investigate the effect of the viruses on fecundity after viral infection, different female fly species were infected with LJV and NFV. For each species, vials with flies (2 males and 2 females) were set up and the parents were discarded as mentioned above. One virgin female and 2 males (3–5 days old) per species were transferred into a vial with cornmeal food with live yeast sprinkled. Females were pricked with either NFV, LJV, or Ringer’s solution (control). Males were not infected. To quantify the number of eggs produced, two females and 2 males were transferred into new vials without yeast with fresh food and allowed to lay eggs for 24h. The number of eggs was quantified under a stereomicroscope at 3-time points, between 4–5, 9–10, and 14–15 dpi. Because D. sechellia fertility is reduced in standard Drosophila diet, it was not used in this experiment.

Disection of ovaries and immunofluorescence microscopy

Immunofluorescence microscopy was used to assess the morphological effects of viral infection on D. melanogaster ovaries. Approximately twenty females were infected as described above with either Ringer’s solution (control), LJV or NFV. After recovery they were placed on vials with fresh cornmeal food supplemented with dried yeast, together with an equivalent number of uninfected males, to stimulate oogenesis. Flies were flipped on fresh vials every day for 10 days. At 10 dpi female flies were anesthetized on ice and ovaries were processed for immunostaining. Briefly, specimens were dissected in ice cold PBS. Ovaries were fixed with formaline (SIGMA 252549) diluted 1/10 in PBS for 20 minutes at room temperature. After fixative was removed, tissue was rinsed and permeabilized in Triton X100 0.1% v/v in PBS, followed by blocking in BBS buffer (PBS, Triton X100 0.1%, NaCl 0.25M, Bovine Serum Albumin 0.3% w/v, Sodium Azide 0.02% w/v). The ovaries were labelled with: mouse monoclonal antibody J2 (Scicons) to detect dsRNA, diluted 1:400, followed by Goat Anti-Mouse IgG H&L (Alexa Fluor® 488) secondary antibody (Abcam 150113) diluted 1:1000; Phalloidin-iFluor 594 reagent (Abcam 176757) to detect Actin filaments, diluted 1:2000; and DAPI (4′,6-diamidino-2-phenylindole) (SIGMA D9542) to detect DNA, 1 ug/mL. After staining, ovaries were placed in mounting medium (PBS, Glycerol 50%, n-Propyl Gallate 0.5 w/v), mounted on slides and imaged at Cambridge Advanced Imaging Center (CAIC) using a Leica SP8 confocal microscope. Images were processed using Leica Application Suite X and Fiji / ImageJ software.

NFV viral oral infection of D. suzukii

To evaluate the fertility of D. suzukii females after oral viral infection over time, D. suzukii female flies were infected with NFV. This experiment was performed as the abovementioned fecundity assay with the exception that females were infected orally. Virgin females (3–5 days old) were orally infected with NFV or Ringer’s solution (control) and transferred into new vials with fresh food and 2 males 3–5 days old. The number of eggs produced was quantified at 4-time points, between 4–5, 9–10, 14–15, and 19–20 dpi.

To investigate whether NFV can establish an oral viral infection on D. suzukii flies, female flies were infected with either NFV or a control solution. Adult females 3–5 days old were orally infected with NFV or Ringer’s solution as the abovementioned oral infection protocol. Flies were transferred into new vials with fresh food and ten days after oral infection, the total RNA of single female flies was extracted, and the relative virus concentration was estimated as mentioned above.

NFV-induced fertility in D. suzuki was determined when allowed to oviposit on blueberry fruits. Female flies were orally inoculated with either NFV or a control solution, as mentioned above. The females were kept in vials with fresh cornmeal food for 4 days. Then, 5 infected females and 5 uninfected males were placed in a vial containing 2 organic blueberry fruits and a damp towel paper to maintain humidity. After 10 days, the flies were discarded and the number of flies that emerged from each vial was measured for 15 days.

Statistical analysis

We conducted all statistical analyses in R [70]. All graphs were generated with the ‘ggplot2’ package [71]. Results from ELISAs, cell numbers and qPCR were analysed using a Type II ANOVA. P values from post-hoc comparisons were adjusted by the Tukey method using the ‘lsmeans’ package [72]. To analyse mortality, a Cox’s proportional hazard mixed-effect model was fitted in the ‘coxme’ package [73], including the fixed effects of Drosophila species, virus treatment (LJV, NFV, or control) and their interaction, and vial of flies as a random effect. Fecundity was analysed Generalized Linear Model with a negative binomial distribution, including zero inflation where appropriate (as assessed by model AIC scores). This was fitted using the ‘glmmTMB’ package [74], including the fixed effects of Drosophila species, virus, and timepoint as fixed effects (together with interactions). To analyse the number of flies hatched between blueberry fruits infected and uninfected with D. suzukii, a t test was used. A Type III Wald test was used to estimate significance of effects within models using the ‘Anova()’ function in the ‘car’ package [75].

Supporting information

S1 Table. Virus detected by qPCR in 19 samples showing high cytopathic effect and high dsRNA concentration.

https://doi.org/10.1371/journal.ppat.1010883.s001

(PDF)

S2 Table. Accession and Isolate Numbers of Virus Sequences.

https://doi.org/10.1371/journal.ppat.1010883.s002

(PDF)

S1 Fig. Immunofluorescent images showing the expression of the plasmid.

(a) DL2-B2 cells overexpressing V5 tagged B2 fusion protein and incubated in the presence of CuSO4 (5 mM) or (b) uninduced (CuSO4-free) for 24 h. Cells were stained with V5 tag mouse monoclonal antibody (1:1000) and labeled with Alexa Fluor 488 rabbit anti-mouse IgG secondary antibody (1,1000). Nuclei (blue) was stained using Hoechst.

https://doi.org/10.1371/journal.ppat.1010883.s003

(PDF)

Acknowledgments

Chrysoula Pitsouli kindly allowed us to use her laboratory in Cyprus. Ben Longdon and Luis Teixeira supplied fly stocks.

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