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here is our discussion padlet

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Phylogenetic  diversity and biochemical diversity of archaea

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Phylogenetic  diversity and biochemical diversity of archaea

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Hi! We're from Group 6

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we are sevengers!!

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Mind Map

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Isolation of fungi from soil

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Discussing about flowchart on isolation of fungi 😊 (27/2/18)

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Microscopic fungi spread in different types of soils in Western Georgia - ScienceDirect

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200 cultures of microscopic fungi have been isolated from the different types of soils (yellow brown forest, raw carbonate, yellow, red, bog, alluvial, yellow podzolic, mountain meadow and brown forest) of Western Georgia. Mainly, representatives of the following classes Zygomycetes, Ascomycetes and Deuteromycetes are spread in the above-mentioned types of soils.

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each 1g of soil estimates 1× 10^6 of fungus spore. Since we will dilute 8 serial dilution, this will dilute the sample.

http://www.scielo.br/scielo.php?script=sci_arttext&pid=S1517-83822012000100039

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updated with grammar correction and reference

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Updated flowchart of fungi inoculation

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Coral reef ecosystems thrive in tropical oligotrophic oceans because of the relationship between corals and endosymbiotic dinoflagellate algae called Symbiodinium. Symbio- dinium convert sunlight and carbon dioxide into organic carbon and oxygen to fuel coral growth and calcification, creating habitat for these diverse and productive ecosystems.Recently, large-scale coral bleaching events have become pervasive and frequent threatening and endangering coral reefs. Because the coral–algal symbiosis is the biological engine producing the reef, the future of coral reef ecosystems depends on the ecophysiology of the symbiosis. This review examines the photobiology of the coral–algal symbiosis with particular focus on the photophysiological responses and timescales of corals and Symbiodinium.

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isolation of giant virus in liquid medium using automated flow cytometry

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isolation of giant virus in liquid medium using automated flow cytometry

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Centre National de la Recherche Scientifique 7278, Institut National de la Santé et de la Recherche Médicale U1095, Unité de Recherche sur les Maladies Infectieuses et Tropicales Emergentes, UM 63, IRD 198, Facultés de Médecine et de Pharmacie, Aix Marseille Université, Marseille, France

Vascular Research Center of Marseille, Institut National de la Santé et de la Recherche Médicale, Faculté de Pharmacie, UMR-S1076, Aix-Marseille Université, Marseille, France

Institut Hospitalo-Universitaire, Méditerranée Infection, Pôle des Maladies Infectieuses et Tropicales Clinique et Biologique, Fédération de Bactériologie-Hygiène-Virologie, Centre Hospitalo-Universitaire Timone, Assistance Publique - Hôpitaux de Marseille and Aix Marseille Université, Marseille, France

The isolation of giant viruses using amoeba co-culture is tedious and fastidious. Recently, the procedure was successfully associated with a method that detects amoebal lysis on agar plates. However, the procedure remains time-consuming and is limited to protozoa growing on agar. We present here advances for the isolation of giant viruses. A high-throughput automated method based on flow cytometry and fluorescent staining was used to detect the presence of giant viruses in liquid medium. Development was carried out with the Acanthamoeba polyphaga strain widely used in past and current co-culture experiments. The proof of concept was validated with virus suspensions: artificially contaminated samples but also environmental samples from which viruses were previously isolated. After validating the technique, and fortuitously isolating a new Mimivirus, we automated the technique on 96-well plates and tested it on clinical and environmental samples using other protozoa. This allowed us to detect more than 10 strains of previously known species of giant viruses and seven new strains of a new virus lineage. This automated high-throughput method demonstrated significant time saving, and higher sensitivity than older techniques. It thus creates the means to isolate giant viruses at high speed.

Introduction

The discovery of Mimivirus opened a new era in virology ( La Scola et al., 2003). The description and genome analysis of this giant virus revealed the presence of numerous genes of eukaryotic and bacterial origins that suggests new metabolic activities for viruses ( Raoult et al., 2004). Recently, it was proposed that the virus be classified under a new order Megavirales ( Colson et al., 2012). Other members of this order are pathogens of animals and several unicellular eukaryotes. Mimivirus deserved some special attention, not only because it was larger than the largest virus known before but also because it was the first virus having dimensions (particle size and genome complexity) that significantly overlap with those typical of parasitic cellular microorganisms. The recent discovery of Pandoraviruses and Pithovirus revealed the extremely large particle size and genetic contents that viruses could possibly possess ( Philippe et al., 2013; Legendre et al., 2014). The study of giant viruses is of interest to multiple fields, including environmental studies aimed at understanding the ubiquity of these viruses, their diversity, and their roles in evolution and ecosystem changes ( Suttle, 2007; Monier et al., 2008a, b; Claverie et al., 2009). In addition, giant viruses are also of interest in clinical research to study their potential pathogenicity and their impact on human health ( La Scola et al., 2005; Khan et al., 2006; Raoult et al., 2006, 2007; Ghigo et al., 2008; Vincent et al., 2009; Mueller et al., 2013; Saadi et al., 2013). There is a need to discover new strategies capable of isolating giant viruses and intensifying studies on these viruses. Finally, improved knowledge on these giant viruses may also shed some light on the evolution of eucaryotes since Marseilleviridae exhibit histone-like proteins ( Thomas et al., 2011). Considering that giant viruses are pathogens for protozoa ( Abrahão et al., 2014) and cause lysis especially of adherent protists, the amoeba co-culture method became the tool allowing the isolation of these giant viruses ( La Scola et al., 2010; Pagnier et al., 2013; Philippe et al., 2013; Legendre et al., 2014). However, this original technique remains fastidious and time-consuming. A recently proposed system for virus isolation on agar plates called the lysis plaque assay allowed hundreds of samples to be tested in a limited time ( Boughalmi et al., 2012). This technique, however, has a major drawback that it is limited to the use of adherent amoeba and protozoa with limited mobility, particularly Acanthamoeba sp. Furthermore, among the giant viruses, CroV was isolated with a highly motile marine protozoa Cafeteria roenbergensis ( Fischer et al., 2010), demonstrating the need for a high-throughput isolation system in liquid media. To use a wider range of protozoa, and to screen the largest number of samples, we modified our isolation procedures slightly and implemented a new high-throughput automated method to detect the presence of giant viruses infecting protozoa in liquid media by flow-cytometry. Flow cytometry-based methods have been previously described for their applications in studies involving pathogenic free-living amoeba ( Muldrow et al., 1982; Avery et al., 1995) but have not focused on isolation-related studies. We associated enrichment methods with flow cytometry.

Materials and Methods

Microorganisms and Culture Conditions

For the development and validation stages, we used the Acanthamoeba polyphaga ( A. polyphaga) strain Linc AP-1 as a host cell for co-culture. This strain was maintained in a 75-cm ² cell culture flask with 30 ml of peptone-yeast-extract-glucose medium (PYG) at 32°C, as previously described ( Pagnier et al., 2013). After 48 h, the amoebae were harvested and pelleted by centrifugation. The supernatant was removed and the amoebae were re-suspended in 30 ml sterile Page's amoeba saline (PAS). Centrifugation and suspension in PAS were repeated twice. After the last centrifugation, the amoebae were counted on counting slides (kova slides, HYCOR Biomedical, Inc., Garden Grove, CA, USA) and adjusted to 5 × 10 ⁵ amoeba/ml. The original strains of A. polyphaga Mimivirus ( La Scola et al., 2003), Marseillevirus T19 ( Boyer et al., 2009), and Faustovirus ( Reteno et al., 2015) were used in suspension in PAS buffer. The remaining microorganisms used as cellular supports or in the gating strategy are detailed in Table 1. The culture conditions for the eukaryotic organisms were the same as for A. polyphaga, with the exception of the handling of Vermamoeba vermiformis ( V. vermiformis) CD119 strain, and Dictyostelium discoideum (D. discoideum) SG2 stain ATCC 44841, where the amoebae were re-suspended in a newly implemented starvation medium capable of maintaining the amoeba under the best conditions for co-culture, without fast encystment as often observed in PAS. Reteno et al. (2015) used this medium for the isolation of Faustovirus. The starvation medium is composed of Yeast extract 2 gm, Glucose 18 gm, Fe (NH ₄) 2(SO ₄) 2.6 H ₂ O 0.02 gm, PAS 1 L, and filtered on 0.22 μm.

TABLE 1. Microorganisms used as cellular supports or in the gating strategy.

Configuration of the Flow Cytometer to Detect Amoebal Lysis

To configure the flow cytometer, rinsed amoebae suspended in PAS were treated in three ways. First, at a concentration of 5 × 10 ⁵ amoeba/ml, amoebae were mechanically lysed by sonication. Sonication causes the amoeba to burst under the effect of ultrasound delivered by a sterile probe plugged into the falcon tube containing the amoebae. We delivered a 60-Hz frequency for 1 min. The procedure was repeated four times. These mechanically treated amoebae served as our positive control for lysis. We infected the second suspension of amoebae, at the same concentration, with Mimivirus. The third suspension of amoebae remained intact and served as a negative control for lysis. The three specimens were then dispensed to a 24-well plate at 500 μl/well. After repeatedly pipetting to detach the adherent amoebae, a 250-μl fraction of each specimen was fixed in paraformaldehyde (4%) then transferred to a suitable tube for flow cytometric analysis on the BD LSRFortessa (BD Biosciences) at 0, 24, and 48 h after infection. Data acquisition was performed using log scales for instrument scatter parameters, forward scatter (FSC) and side scatter (SSC), respectively associated with size and internal complexity of the event analyzed. Protocol threshold was adjusted on FSC parameter. Acquisition and analysis were performed using "BD FACSDiva Software." The number of collected events was fixed for all specimens. We gated our amoeba population using the density plot representation SSC versus FSC. We performed serial dilutions from 10 ⁶ to 10 ¹ amoebae/ml to calibrate the analyzer, to determine the reliability of the counts and to establish the limit sensitivity of detection of the flow cytometer to choose a lysis threshold that corresponds to the loss of amoebae. All amoebal counts performed by flow cytometry were also performed on kova slides to validate the counts. The quantification was also verified using counting beads (true count or cytocount "DakoCytomation," a suspension of concentration-calibrated fluorescent microspheres). This provides the absolute count of the amoebal population using the following equation: (number of cells counted/number of Cytocount TM beads counted) × Cytocount TM concentration (1100 beads/μl) × dilution factor ( Khan et al., 2010). We also used the Sytox nucleic acid stain for flow cytometry (Molecular Probes, Life Technologies, USA) to determine the viability of our host cells. All experiments were performed in triplicate.

Validation of the Technique Using Samples Containing Giant Viruses

By blind culture, without microscopic observation, we tested several samples of water and soil. Some of the samples were artificially contaminated with giant virus suspensions using Mimivirus and Marseillevirus (10 μl of virus at a concentration of 10 ⁵ virus/ml were used to contaminate 3 ml of the sample). We contaminated 5 of 20 samples before proceeding to co-culture then for flow cytometry detection.

In a second step, we tested 80 environmental samples, including 78 samples, previously tested negative by co-culture with A. polyphaga and the two samples from which the Terra1 and Terra2 Mimivirus strains had previously been isolated ( La Scola et al., 2010). The processing steps for isolating these Terra1 and Terra2 strains have previously been described ( Pagnier et al., 2013; Yoosuf et al., 2014). For our technique, we conducted three blind co-cultures as an enrichment step; primo-culture, sub-culture, and final culture without any microscopic observation. Data acquisition on the flow cytometer was performed 24 and 48 h following the final enrichment step. The number of collected events was fixed for all samples. Uninfected amoebae were used as a negative control. They were analyzed first and used to gate the population of amoebae according to the parameters defined above. The percentage of live and dead cells was evaluated. Moreover, the presence and percentage of debris characterizing amoebal lysis was also evaluated in the FSC SSC plot. All data from LSR Fortessa were analyzed using kaluza software (Beckman Coulter). An arbitrary threshold for amoebal loss of more than 50% after 24-48 h was used to define the possible presence of a lytic agent.

Gating of Microorganisms Using Fluorescence Staining

We used viruses of known sizes from our collection. We also used amoeba-associated chlamydiae, which can be amoebal pathogens ( Greub et al., 2003). We often find chlamydiae in culture because of their resistance to the antibiotic mixtures used. Viruses and chlamydiae were analyzed on LSR fortessa. To set up Side Scatter and a 520-nm fluorescence channel on the flow cytometer, we used Megamix+ SSC beads (Biocytex, Marseille, France). This blend of size and fluorescence-calibrated beads (0.16, 0.22, 0.24, and 0.5 μm) is specially designed to evaluate and standardize Side Scatter parameters for sub-micron biological events. Virus and bacteria were gated and characterized using their original SSC/SYBR profile. Acquisition was performed using a fluorescent threshold to limit Side Scatter background noise. Sybr green was used to label DNA (SYBR Green I nucleic acid gel stain; Molecular Probes, Life Technologies, USA). The final concentration of SYBR Green I was a 10⁻⁴ dilution of the commercial stock solution. For an amoebal suspension at a density of 1 × 10 ⁶, we added 5 μl of the DiD cell labeling solution and mixed thoroughly by gentle pipetting. The cells were then incubated for 1 h at 30°C, washed three times by centrifugation and re-suspended in PAS (the purpose was to eliminate amoebal debris from the gating). We used Mimivirus, Marseillevirus, Faustovirus, and a parachlamydia-like bacterium (an intra-amoebal chlamydia isolated in our laboratory). Data for this chlamydia, that we proposed to name Candidatus Rubidus. massiliensis ( C. R. massiliensis), were submitted to the EMBL database and were assigned Bio-projects number PRJEB6078; the accession numbers for the genome at EMBL are CCSC01000001-CCSC01000005) and the corresponding paper has been also submitted. A threshold on SYBR green parameter to limit background noise from side scatter was applied to detect all three potential viruses or/and bacteria in the samples. We then prepared mixes of different viral populations at various concentrations. To avoid coincidence of viral particles (i.e., two or more particles being simultaneously within the sensing zone), the samples were diluted such that the event rate was between 100 and 1000 viruses s⁻¹. We tested 30 blinded mixes of viral suspensions in order to validate the proof of concept. After optimizing the gating strategy, we screened all of the previously tested real environmental samples that showed host cell lysis and contained mixture of viruses. Ten μl of the culture supernatant after host lysis detection on the sample was diluted to 1/10 ⁴ in PAS, then stained with SYBR green at 30°C over night. We then performed gating to distinguish between a single virus, a viral mixture and bacteria. The results of the gating were confirmed by targeted PCR of the different giant viruses ( Ngounga et al., 2013).

Automation and Adaptation of the Method to Other Protozoa

We adapted our method to all protozoa enumerated in Table 1. All tests performed during the developmental stage with A. polyphaga, were carried out on each protozoon. We also tested protozoan survival in the antibiotic and antifungal mixture before the gating stage and the use of the protozoa as cell hosts in culture. Initially, all experiments were performed in 24-well plates. Because our purpose was high-throughput detection, we then adapted our technique to 96-well plates. We optimized the concentration and volume of amoebae in these plates (150 μl of amoeba suspension at a concentration of 5 × 10 ⁵ amoeba/ml for Acanthamoeba castellanii ( A. castellanii), A. polyphaga, Poterioochromonas malhamensis ( P. malhamensis), Tetrahymena hyperangularis ( T. hyperangularis), and 150 μl of amoebal suspension at a concentration of 10 ⁶ amoeba/ml for V. vermiformis, and D. discoideum), and used 100 μl of each sample. Following optimization in 96-well micro-plates, we integrated a high-throughput sampler (HTS) from BD Biosciences with the LSRFortessa analyzer. The HTS loads samples from the 96-well plates and performs fully automated acquisition within 40 min. Under high pressure and after several mixes, the HTS is easily capable of re-suspending the adherent amoebae in the plates. The above experiments were performed in triplicate on these 96-well plates to confirm the reliability of the technique when optimized for micro-plates. The same principle of automation was applied to the gating strategy. We adapted the same experiment principle performed in tubes to 96-well micro-plates.

Automated Sample Screening

For sample testing, we used our high-throughput method to launch high-speed screening for giant viruses in environmental and clinical samples. There were 735 bronchoalveolar lavages (BALs) from the intensive care unit of the La Timone Marseille healthcare center, France. We collected 250 samples of sewage (80 from Marseille-France, 50 from Dakar-Senegal, 70 from Tripoli Lebanon, and 50 from Guyana DOM France), 1025 seawater and soil samples (400 seawater and 50 soil samples from Marseille France, 400 seawater samples from La Ciotat, Bandol, France, 160 seawater and 15 soil samples from Brittany France). All samples were tested on four protozoa ( A. polyphaga, V. vermiformis, T. hyperangularis, and P. malhamensis). The enrichment step used antibiotic and antifungal mixtures containing 10 μg/ml vancomycin (Mylan, Saint-Priest, France), 10 μg/ml imipenem, 20 μg/ml ciprofloxacin (Panpharma, Z.I. du Clairay, France), and either a fungicide [30 μg/ml thiabendazole (TBZ)] or an antifungal (30 μg/ml Fungizone, Bristol Myers Squibb, Rueil-Malmaison, France), as previously described ( Pagnier et al., 2013). After host cell lysis detection, and preliminary gating characterization, lytic agents were then identified using MALDI-TOF MS for bacteria ( Seng et al., 2010, 2013) and a specific primer-probe system for giant viruses as previously described ( Pagnier et al., 2013).

Ethics Statement

LBA samples were received from an anonymized patient file for the research of emerging pathogens (IFR48- No 13-031).

Statistics

We used the R software package ( R Core Team, 2013), a language and environment for statistical computing and analysis version 3.0. Spearman correlations were used to look for a link between kova slides counts and flow cytometry counts. A p-value of < 0.05 was considered statistically significant. Experiments were performed in triplicate.

Results

Detection of Amoebal Lysis by Flow Cytometry

For flow cytometry configuration, we observed a significant decrease (90%) in the percentage of amoebae and an increase in the number of debris following mechanical lysis and infection with Mimivirus at 24 and 48 h compared to the percentage of control amoebae. The counts on kova slides and by flow cytometry are correlated (Spearman r = 0.97, p-value < 0.001). The detection limit of the FACS was of 10 ² amoebae/ml, which is sufficient for detecting the arbitrary threshold of an amoebal loss greater than 50% after 24 h.

Technique Concept Validation

During the artificial contamination tests, we managed to detect all five artificially contaminated samples among the 20 tested samples. At 0 h, the negative control used as a reference population contained 86% of amoebae. The same percentage was equally observed for the other samples at this time point. It is important to note the physiologic increase in sytox positive dead cells for both control and tested samples at 24 h post-culture. At 48 h we still had our negative control with no substantial changes. By contrast, the positive control showed an almost complete loss of gated amoeba with the highest percentage of debris. These results were obtained by applying size and granularity parameters (FSC and SSC channels). The sytox nucleic acid stain was used to confirm and validate our gatings (Figure 1). For all real environmental samples the results obtained by our new high-speed method were identical to those obtained previously by the traditional co-culture method ( Pagnier et al., 2013). We detected the two positive samples among the 80 tested samples. PCR for giant viruses, targeting pol B DNA, was performed to identify the sequences corresponding to Terra1 and Terra2 viruses ( La Scola et al., 2010). In addition, a single sample, previously identified as negative in co-culture, appeared positive by cytometry. For that sample, additional investigations, such as staining (Gram, Giminez, Hemacolor, negative staining) and a PCR system targeting giant viruses ( Ngounga et al., 2013) were performed on this sample, and demonstrated a newly isolated Mimivirus of genotype C ( Colson et al., 2012), submitted under bioproject PRJNA278018.

FIGURE 1. Detection of potential amoebal infection. A living amoebal population was used as a negative control with Sytox staining. Three gates were designed in the FSC-SSC plot, potentially corresponding to live cells, dead cells and debris. At 0 h the negative control and sample showed the same gated populations (A,B). After 24 h of culture, a physiologic increase in dead cells was observed for both the negative control and test samples (C,D). The appearance of a large population of debris confirmed a potential infection after 48 h of culture and a larger part of dead cells in the tested samples than in the negative control showing no substantial changes in the gated populations ( E,F representing the negative control panel after 48 h and, G,H representing the sample containing Mimivirus after 48 h).

Results of the Gating Strategy

Single virus/bacterium analysis allows SSC/DNA content profile to be determined. After setting up the instrument using Megamix and SSC beads (Figure 2A), we found that Chlamydia gave an SSC signal close to the 0.55 μm Megamix and a high SYBR labeling, as expected (Figure 2B). Mimivirus with SSC beads showed intermediate SYBR labeling between 0.24 and 0.55 μm (Figure 2C). Marseillevirus with SSC showed the lowest SYBR signal of the four species at 0.16 μm bead SSC signal (Figure 2D). Finally, Faustovirus showed minimal SSC (below the 0.16 μm bead) but a higher DNA content than Marseillevirus (Figure 2E). A linear correlation was found between green fluorescence intensity and genome size for giant viruses ( r ² = 0.98). We confirmed our findings of the presence of a viral mixture of Mimivirus and Marseillevirus in natural environmental samples by this gating strategy (Figure 2F). All of the samples that were previously tested in culture and that showed host cell lysis were screened by the gating technique. We were able to detect all of the viral or bacterial populations in the samples that were amplified by the enrichment step. All of the results were confirmed by a giant virus-specific PCR system ( Ngounga et al., 2013).

FIGURE 2. Cytometric analysis of amoebal pathogens: giant viruses and chlamydiae. (A) Using Megamix+ SSC beads (0.16-, 0.2-, 0.24-, and 0.5-μm fluorescent beads), LSR fortessa was easily set up for SSC, and a 520 nm Fluorescent channel (for SYBR DNA virus labeling) was used for virus range analysis. Fluorescence threshold was set to limit background noise of the instrument. Each microorganism was first analyzed alone to clearly define virus gates [(B) chlamydia, (C) Mimivirus, (D) Marseillevirus, and (E) Faustovirus]. Thanks to the wide resolution of LSR fortessa in SSC and the clear DNA content between the different biological events, four different gates were created. Due to the high concentration of virus in the supernatant after amoebal burst, a virus was considered present if its concentration was more than 10 ³ virus/μl (counting beads). (F) Viral mixture of Mimivirus and Marseillevirus, detected in the same sample.

Results of the Method Automation and Sample Screening

After adapting our technique to all of the protozoa types used as host cells, the automation allowed us to test 96 samples in less than 40 min. Of the 735 BAL samples tested on A. polyphaga, V. vermiformis, T. hyperangularis, and P. malhamensis, three led to A. polyphaga lysis detection. We identified three bacteria ( Pseudomonas aeruginosa, Acinetobacter baumannii, and Stenotrophomonas maltophilia), but no giant viruses. Among the sewage samples carried out on A. polyphaga, 10 led to detection of amoebal lysis. The gating strategy revealed the presence of viral mixtures in many samples. PCR performed on positive wells confirmed the gating strategy results and revealed the simultaneous presence of a Mimivirus of genotype A, a Mimivirus of genotype C and a Marseillevirus in one sample from Marseille. A second sample contained a Mimivirus of genotype A from Lebanon, and the eight remaining samples contained a mix of Mimivirus genotypes A and C, 3 from Dakar and 5 from Marseille. For the same sewage samples inoculated on V. vermiformis, five were positive and we isolated a new giant virus specific for Vermamoeba, named Faustovirus E12 ( Reteno et al., 2015). Seven other viruses belonging to the same family were also isolated from the different sewage samples (3 from Marseille, 3 from Dakar, and 1 from Lebanon; the genomic data corresponding to these isolates were submitted to the EMBL database and were assigned Bio-projects numbers respectively: E24: PRJNA279158, E23: PRJNA279157, E9: PRJNA279166, D3: PRJNA279161, D5: PRJNA279159, D6: PRJNA279164, Liban: PRJNA279165). No lysis was detected on the other tested protozoa. The results of this screening are summarized in Table 2.

TABLE 2. Isolates from environmental and clinical samples represented in collection area and with the different used host cells.

Discussion

The study of giant viruses is a newly emerging field in virology. Metagenomics data have suggested the ubiquity of giant viruses not only in the environment ( Claverie, 2004; Monier et al., 2008a, b; Claverie et al., 2009) but also in humans ( Colson et al., 2013). We must therefore search for these viruses at large scales. This high-throughput sample screening was almost impossible to conduct with our usual techniques of co-culture used by almost all the scientific committee nowadays, especially for non-adherent or motile protozoa. The method we developed in this paper provides the necessary characteristics for high-throughput analyses by allowing host cell lysis detection in micro-assays, followed by a gating strategy capable of preliminary characterization of the infectious agent responsible for host cell lysis. The method was developed in three stages. The first step included the calibration and configuration procedures performed with ( A. polyphaga) to detect amoebal lysis. This was followed by the validation stage, then by the automation that simplified the enrichment step. We then adapted this technique to other types of protozoa, which we think that may harvest other giant viruses, to finally test hundreds of environmental and clinical samples for the presence of viruses.

Detecting the loss of the amoeba-gated population represents a signal for the presence of a pathogenic agent responsible for the cytopathogenic effect and amoebal lysis. It is thus important to accurately determine the percentage of each population using the best possible gating. These percentages can vary, so it is important to always have a negative control containing the host cell under the most favorable culture conditions, which will be used as the reference population for the rest of the samples. The Sytox nucleic acid stain can always help us evaluate the viability of our amoebae.

Following the successful results from the developmental stage, the proof of concept was validated by detecting not only known viruses from both artificially contaminated samples and environmental samples but also a new virus, previously undetected by the traditional methods of co-culture with Acanthamoeba ( Pagnier et al., 2013). The isolation of this later strain suggests a higher sensitivity of our new technique. This method allowed the rapid isolation of more than 10 new strains of giant viruses. No giant virus was found in the BAL samples, clearly confirming the rare direct evidence for detecting giant viruses in BAL, at least using A. polyphaga as a support for the culture. We did, however, detect three bacterial strains. These bacteria were isolated from intensive care unit patients because they were resistant to the antibiotic mixtures used during the different stages of the co-culture. The discovery of a new host-specific giant virus ( Reteno et al., 2015) is good proof that other protozoa species should be tested as hosts cells capable of harvesting giant viruses. We also succeeded in isolating 7 new isolates of this newly discovered Faustovirus.

Cytometric analysis has several advantages over current isolation methods. Using this system, it is possible to rapidly quantify amoebae and identify the presence of pathogenic microorganisms, such as giant viruses, in any type of sample.

A major benefit of this high-throughput method is its automation. This automated method largely reduces the time required to screen 1000s of samples for giant viruses, especially as data acquisition on the flow cytometer does not exceed half an hour. The cytometer is able to detect lysis at an early stage, which is better than microscopic observation. Other advantages of flow cytometry over routine microscopic observation include a high degree of statistical precision due to the large numbers measured, the high-speed gating and identification of the amoebal status, along with the elimination of subjectivity.

The production of amoebae in culture was one of the limiting factors of the old techniques ( Boughalmi et al., 2012; Pagnier et al., 2013) especially as they remain time consuming and not amenable to automation. In this respect, the enrichment steps using micro plates greatly reduced culture preparation time. For example, a 96-well plate of samples requires four times fewer amoebae than the routine culture performed in 12- or 24-well plates. All of these advantages made our method a compatible tool for high-throughput platforms. Although the Acanthamoeba species allowed the isolation of giant viruses, we know that other protozoa are capable of growing similar viruses, where Cafeteria roenbergensis, a marine biflagellate was isolated with its associated virus CroV ( Garza and Suttle, 1995; Fischer et al., 2010). It is the same case for PgV-16T ( Phaeocystis globosa virus) that infects a nanoplanktonic marine alga ( Santini et al., 2013). The isolation of the latter two viruses was the consequence of their associated protozoan isolation and was not from direct research using co-culture. Because the protozoan world is vast, the diversity of giant viruses is most likely vast as well, and with the previous culture techniques, our research and knowledge was limited. Indeed, several electron microscopy photos suggest that some relatively large viruses showing similar morphology to the NCLDV family are present within numerous other eukaryotic microorganisms, including the parasites Giardia and Blastocystis ( Stenzel and Boreham, 1997).

The method we designed is readily amenable to other protozoa, especially those capable of growing only in liquid media, such as ciliates and flagellates. Thus, it is adapted to other types of protozoa and allows us to enhance the number of host cells for the screening of diverse environmental or human samples. The limitation of the method comes from the detection limit of the cytometer where we cannot go below 10 ² cells/ml. The method still requires a culture enrichment step that needs the host species to be cultivated in sufficient number, which is a limiting factor with species such as Giardia and Blastocystis. We are aware that it will be hard to cover all types of protozoa and we will try to improve the detection sensibility in order to be able to detect lysis on fastidious and slowly growing host cells.

The gating strategy will detect the nature of the lytic agent (virus, bacteria, or mixture in the same sample) without losing time, especially compared to standard staining techniques such as Gram, hemacolor, dapi, etc. We developed and applied the gating strategy using SYBR Green 1 inspired from previous works ( Marie et al., 1999; Brussaard et al., 2000; Brussaard, 2004). Based on the different fluorescence profiles that we have already characterized for each gated microorganism, we can easily discriminate between all known giant viruses that we could isolate after detection of lysis. The gating strategy will limit us to only gated microorganisms, so it would be interesting to enlarge the gating data by discovering more giant viruses. Many samples contained mixtures of viruses. For this, we believe that the adaptation of the method to a real FACS sorting technique will be the best strategy for better preliminary characterization of new isolates and especially for separating viral mixtures in order to allow a better comprehensive pangenomic study of these virus families and their ecosystems. The integration of cell sorting to purify cell populations enables more facilities to isolate, produce and clone giant viruses, while offering the possibility of studying the infectious stage before amoeba burst to possibly identify non-lytic viruses or endosymbionts and this by focusing on the cell host nuclear changes and viability biomarkers.

Conflict of Interest Statement

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

References

Greub, G., La Scola, B., and Raoult, D. (2003). Parachlamydia Acanthamoeba is endosymbiotic or lytic for Acanthamoeba polyphaga depending on the incubation temperature. Ann. N. Y. Acad. Sci. 990, 628-634. doi: 10.1111/j.1749-6632.2003

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# data[].attributes.attachment_link.title
High-Throughput Isolation of Giant Viruses in Liquid Medium Using Automated Flow Cytometry and Fluorescence Staining

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The isolation of giant viruses using amoeba co-culture is tedious and fastidious. Recently, the procedure was successfully associated with a method that detects amoebal lysis on agar plates. However, the procedure remains time-consuming and is limited to protozoa growing on agar. We present here advances for the isolation of giant viruses.

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Here are some identification for protozoa (Toxoplasma gondii

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000 units/ml penicillin and 10 mg/ml streptomycin in 0.9% normal saline sterile filtered

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L-Glutamine-Penicillin-Streptomycin Solution w/ 200mM L-Glutamine, 10,000 units/ml Penicillin and 10 mg/ml Streptomycin in 0.9% normal saline Sterile filtered | Animal Cell Culture | Cell Culture Reagents | Antibiotic Solutions | L-Glutamine-Penicillin-Streptomycin Solution

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Buy and get information for L-Glutamine-Penicillin-Streptomycin Solution w/ 200mM L-Glutamine, 10,000 units/ml Penicillin and 10 mg/ml Streptomycin in 0.9% normal saline Sterile filtered, A007, Animal Cell Culture, Cell Culture Reagents, Antibiotic Solutions, L-Glutamine-Penicillin-Streptomycin Solution

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Toxoplasma gondii

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Isolation of Hendra virus from pteropid bats: a natural reservoir of Hendra virus

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Isolation of Hendra virus from pteropid bats: a natural reservoir of Hendra virus

# data[].attributes.attachment_link.title
Microbiology Society Journals | Isolation of Hendra virus from pteropid bats: a natural reservoir of Hendra virus

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Since it was first described in Australia in 1994, Hendra virus (HeV) has caused two outbreaks of fatal disease in horses and humans, and an isolated fatal horse case. Our preliminary studies revealed a high prevalence of neutralizing antibodies to HeV in bats of the genus Pteropus, but it was unclear whether this was due to infection with HeV or a related virus.

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Isolation and characterization of Pseudomonas fluorescens bacteria

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Isolation of fungi from clinical specimens

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Isolation of fungi from clinical specimens

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https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3536260/

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Molecular identification and control of somepathogenicFusarium species isolated from maize in Egypt

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Molecular identification and control of somepathogenicFusarium species isolated from maize in Egypt

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Molecular phylogenetic identification of endophytic fungi isolated from three Artemisia species 

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Molecular phylogenetic identification of endophytic fungi isolated from three Artemisia species 

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Abstract

Escherichia coli O157:H7 (EcO157), an agent of life threatening hemolytic-uremic syndrome, resides in ruminants and is released in feces at numbers as high as 10 million cells/gram. EcO157 could survive in manure for as long as 21 months, but we observed a 90% decrease in cells of an outbreak strain of EcO157 within half a day in wastewater from dairy lagoons. Although chemical, environmental and biological factors may be responsible for this decrease, we observed an 11-fold increase in native protozoa when wastewater was re-inoculated with 2×10 ⁷ cells of EcO157/mL. These protozoa engulfed the green fluorescent protein labeled EcO157 within 2 hours after inoculation, but expelled vacuoles filled with live EcO157 cells within 3 days into surrounding wastewater, whereas other protozoa retained the EcO157-filled vacuoles for 7 days. EcO157 was not detected by confocal microscopy either inside or outside protozoa after 7 days. Mixed cultures of protozoa enriched from wastewater consumed EcO157 preferentially as compared to native aerobic bacteria, but failed to eliminate them when EcO157 cells declined to 10 ⁴/mL. We isolated three protozoa from mixed cultures and typed them by 18S sequencing as Vorticella microstoma, Platyophyra sp. and Colpoda aspera. While all three protozoa internalized EcO157, only Platyophyra and Colpoda acted as predators. Similar to mixed cultures, these protozoa failed to eliminate EcO157 from PBS containing no other supplemental nutrients or prey. However, spiking PBS with cereal grass medium as nutrients induced predation of EcO157 by Platyophyra sp. after 3 days or enhanced predation by Colpoda after 5 days. Therefore, attempts to enrich protozoa to decrease EcO157 from dairy lagoons, may correspond to an increase in protozoa similar to Vorticella and possibly facilitate transport of bacterial pathogens to food crops grown in proximity.

Introduction

Produce was linked to 23% of 117,136 foodborne illnesses associated with outbreaks occurred during 1998 and 2007 in the United States [1]. Escherichia coli O157:H7 (EcO157) infections were responsible for 5% of the 684 produce associated outbreaks during the same period and 19% of all EcO157 associated outbreaks were due to consumption of contaminated produce [1]. In addition, major outbreaks associated with produce occurring in the US and in other parts of the world indicate that contamination has occurred in many cases in the field (pre-harvest), so it is critical to identify sources of pathogens in the environment and interventions for minimizing them [2]. Since pathogen contamination on 'ready to eat' produce cannot be removed easily [3], prevention of pre-harvest contamination is critical. Pre-harvest contamination emphasizes the importance of identifying animal reservoirs and the biological and environmental factors that regulate the proliferation of pathogens during their transport from ranches or dairies or wildlife to produce grown in proximity to these locations.

Although highly prevalent in some cattle herds in the US [4], the shedding of EcO157 is transient but elevated during summer and fall months [5]. Even if fecal shedding of pathogen is heavy during summer months, it has been reported in some studies that the pathogen disappears rapidly as the temperature increases during summer [6], [7]. However, we reported previously the lack of detection of the EcO157 in manure and wastewater collected from dairies in the central valley of California between 2001 to 2003 [6]. In contrast, EcO157 was isolated from watersheds, cattle herds, and wild pigs on the central coast of California throughout spring and summer months of 2004-2006 [8]. These results indicate a dynamic incidence that depends on spatial and temporal factors and possibly agricultural practices.

Pathogenic EcO157 often escapes detection because of its low numbers or, possibly, because it is concealed in alternate hosts such as flies, birds, dogs, cats, rodents, horses [9] and protozoa [10], [11]. Protozoa have gained notoriety as the 'Trojan horses' [12] of bacterial pathogens. Salmonella enterica serotype Typhimurium phage type DT104 [13], Legionella pneumophila[14] and Mycobacterium avium[15] passaged through protozoa have been reported to become hypervirulent; and coliforms (including non-pathogenic E. coli) and other pathogenic bacteria demonstrated increased resistance to chlorine disinfection [16]. It has been shown recently that Shiga toxin encoding prophage augmented the fitness of EcO157 in the presence of grazing protozoa, Tetrahymena pyriformis[17], thus providing a potential environmental selection pressure for bacteria to retain prophage encoding Shiga toxin. Although, EcO157 was known to survive and proliferate in co-culture with protozoa [12], [18], a predatory interaction could not be established between rumen ciliate protozoa and Shiga-toxigenic E. coli[19].

Protozoa constitute half of the microbial biomass in rumen and are essential in providing nutrients to the host through a symbiotic relationship with other rumen inhabitants [20]. Ruminants are primary reservoirs of EcO157 and may sometimes shed as many as 10 ⁷ per gram of feces [21], but there are no reports of evidence for the consumption of EcO157 by rumen protozoa. However, protozoan predation was linked indirectly to the decrease of EcO157 in well waters [22] and wastewater from a dairy [23]. Direct evidence for predation was reported recently for enteropathogenic E. coli in co-culture with Acanthamoeba polyphaga[24]. However, none of these studies have addressed EcO157 and protozoa in dairy wastewater even though both predator and prey in feces end up in lagoons. Since, EcO157 survived for 21 months in manure [25], but declined rapidly in wastewater from lagoons [6], we monitored protozoa in wastewater from dairy lagoons to determine if they are responsible for the decrease in EcO157. In addition, protozoa were isolated, identified by 18S rRNA sequences and evaluated further for their ability to consume and internalize EcO157. We used an outbreak strain of EcO157 marked with green fluorescent protein (GFP) reporter to confirm internalization of EcO157 in natural protozoa and to monitor the influence of nutrients and resident bacteria on protozoan predation in wastewater.

Materials and Methods

Enumeration of protozoa, EcO157, aerobic and coliform bacteria

Protozoa from 10-fold serial dilutions of wastewater samples were determined using the most-probable number (MPN) method [26]. Triplicate 50-µL portions of each dilution in optically clear 96-well polystyrene plates (Corning, Inc., Corning, NY) were monitored for protozoa using 40 to 200× magnification of an inverted microscope (Leica DM IL, Leica Microsystems, Wetzlar, Germany). Minimum detectability by the MPN method was 3.6 protozoan cells per 50 µL samples.

The populations of rifampin and nalidixic acid resistant EcO157 were monitored by plating 100-µL portions of serial dilutions of wastewater in 0.01M PBS on LB agar supplemented with 100 µg/mL rifampin, and 50 µg/mL each of nalidixic acid and cycloheximide (Sigma-Aldrich, St. Louis, MO). Cycloheximide was used to control fungal growth on enumeration media. The populations of GFP-EcO157 strain MM123 were monitored by plating the serial dilutions on LB agar supplemented with 100 µg/mL rifampin, and 50 µg/mL each of cycloheximide and kanamycin. The plated media were incubated overnight at 37°C and counted, except that the fluorescent colonies of MM123 were counted on a UV Transilluminator (Fotodyne, Hartland, WI).

One-mL portions of serial dilutions of wastewater were plated on Petrifilm count plates (3M Microbiology Products, St Paul, MN) to enumerate aerobic and coliform bacteria [6].

Protozoan response in wastewater to repeat inoculation of EcO157

Native protozoan populations were monitored in wastewater supplemented with or without EcO157 strain MM123 as alternate prey. MM123 is a spontaneous rifampin-resistant (100 µg/mL) mutant of GFP-labeled strain RM2315 (original designation SEA13B88; provided by S. Wegant, FDA-Bothell, WA), which has been described previously [6], [27]. The wild type strain was the cause of a major outbreak associated with non-pasteurized apple juice [28]. Wastewater was collected from an aerated manure lagoon from a medium-sized dairy (ca. 800 milking head) in central California (Oakdale, CA) and acclimated overnight at room temperature prior to use [23]. Wastewater supernatant, similar to dairy wastewater with solids removed and reused to wash free-stall lanes on the diary, was used to accommodate enumeration of motile protozoa by the MPN method. Bottom feeding amoebae settling with manure solids were avoided. Tests were conducted in triplicate 250-mL Erlenmeyer flasks containing 100 mL of wastewater samples inoculated with two different levels of MM123: 1.1±0.2×10 ⁵ and 8.6±0.7×10 ⁷ CFU/mL. Overnight growth of EcO157 in LB broth supplemented with 100 µg/mL of rifampin and 50 µg/mL of kanamycin was centrifuged and resuspended in 0.01M PBS was used as a food source for protozoa. The flasks were incubated at 25°C on a gyratory shaker operated at low speed (50 rpm) to simulate circulating aeration in the lagoons, and samples were monitored at various intervals for GFP-EcO157, protozoa and aerobic bacteria. The microcosms containing the high inoculum were re-inoculated with 1.7±0.2×10 ⁷ CFU/mL after 7 days.

Confirmation by confocal microscopy of protozoan feeding of EcO157

Wastewater samples from the experiment described above were examined at various intervals to determine the diversity of native protozoa that engulf GFP-EcO157. Five-hundred-microliter samples of wastewater containing 0.3% Polyox ( c. 4,000,000 mol wt, Sigma-Aldrich) were stained with 40 mM propidium iodide and 10 mM SYTO 59 (Invitrogen, Carlsbad, CA) for 1 h to stain dead and live bacteria, respectively, then visualized by scanning on a Leica TCS-NT confocal microscope. Polyox enabled visualization by slowing down the protozoan movement. The internalization of EcO157 was determined by scanning through the total depth of protozoa as 5 to 10 µm optical sections. The sections were compiled using the Leica TCS NT Software and the fluorescence by GFP, propidium iodide, and SYTO 59 were assigned green, red and blue colors, respectively, for compiled images.

The diversity of protozoa in 10-fold concentrated wastewater samples was also determined by phase contrast microscopy using a Leica DMR microscope after staining the samples for 1 h with 1 µL each of neutral red (100 µg/mL; Sigma-Aldrich) and acridine orange (1 µg/mL; Invitrogen), and diluting samples with an equal portion of glycerol to decrease protozoan motility.

Consumption of EcO157 by a mixed culture of natural protozoa from wastewater

Consumption of EcO157 strain MM123 by protozoa was determined using natural protozoa enriched from wastewater. One liter of wastewater was filtered through a 250 µm sieve, centrifuged (300×g, 10 min) and the pellet containing protozoa was transferred in 10 mL of 0.01M PBS to 250 mL of Sonneborn medium (Solution 1 of ATCC medium 802, https://www.atcc.org/Attachments/4018.pdf, Accessed 6 October 2010) supplemented with 5% of 0.2-µm-filter-sterilized wastewater (hereafter referred to as cereal grass-wastewater medium [CGWM]). A 7-day old growth of protozoa from CGWM was concentrated at 300×g and suspended in 0.01M PBS as the inoculum to test predation. Tests were conducted in triplicate 250-mL Erlenmeyer flasks containing 50-mL of CGWM inoculated with 2×10 ⁷ cells of EcO157 and 8×10 ³ cells of enriched protozoa per mL (2,500 bacteria/protozoa). CGWM inoculated with only MM123 was used as a no-predation control. The populations of GFP-MM123, protozoa and aerobic bacteria were monitored at various intervals.

Isolation, purification and 18S rRNA characterization of protozoan monocultures

Protozoan enrichments described above were transferred sequentially 3 times at 7-day intervals to 250-mL portions of CGWM and incubated at 25°C on a gyratory shaker operated at 50 rpm. After the third transfer, 100-µL portions of ten-fold serial dilutions of the growth was transferred to optically clear 96-well polystyrene plates (Corning) with each well containing 100-µL cereal grass medium without the supplemental wastewater (CGM) and incubated at 25°C. Purity of protozoan cultures in 96-well plates was checked microscopically at periodic intervals and purified further by transferring 10-fold serial dilutions at 7-day intervals to fresh 100-µL of CGM until they were confirmed pure by microscopy.

Protozoa were grown in 100 mL-portions of CGM, concentrated by centrifugation at 300×g for 10 min and the cell pellet was used to extract DNA used for the amplification and sequencing of 18S rRNA as described in Method S1.

Consumption of EcO157 by monocultures of protozoa from wastewater

Consumption of EcO157 strain MM123 was determined using three cultures of protozoa isolated from wastewater. Protozoan growth from 250-mL CGM supplemented with 50 µg/mL of tetracycline was centrifuged (300×g, 10 min), the pellet was washed twice in 100 mL of 0.01M PBS to remove bacterial carry-over, and re-suspended in 100 mL PBS as the inoculum to test predation. Tests were conducted in triplicate 250-mL Erlenmeyer flasks containing 50 mL of PBS inoculated with 1.1×10 ⁷ cells/mL of EcO157 and 1.1×10 ³ cells/mL of Vorticella microstoma or 1.5×10 ² cells/mL of Platyophyra sp or 2.1×10 ³ cells/mL of Colpoda aspera. These treatments provide ∼10,000, 70,000 and 5,000 cells of EcO157 as food for each one of the Vorticella, Platyophyra and Colpoda cells, respectively. The survival of EcO157 in the absence of predation was monitored in PBS inoculated with only EcO157. As protozoa appeared to slow down in PBS, the treatments were supplemented with 5% CGM on day 3 for Vorticella and Platyophyra and on day 6 for Colpoda. The populations of GFP-EcO157, protozoa and aerobic bacteria were monitored at various intervals.

Supplementation of wastewater to increase EcO157 and native bacteria in wastewater and stimulate predation

In an attempt to increase the numbers of native protozoa that consume EcO157, the wastewater was supplemented with or without 10% (w/v) LB broth and inoculated with two different concentrations (7.3±0.7×10 ⁶ and 6.6±0.9×10 ⁸ CFU/mL) of EcO157 strain MM149 as a supplementary food source (850 and 77,000 cells of EcO157 per protozoan, respectively). MM149 was isolated from dairy manure in northwestern Oregon and was selected for rifampin (110 µg/mL) and nalidixic acid (50 µg/mL) resistance to aid in discriminating the colonies from the bacteria in wastewater during enumerations [23]. Overnight growth of MM149 in LB broth supplemented with 100 µg/mL of rifampin and 50 µg/mL of nalidixic acid, centrifuged and resuspended in 0.01M PBS was used as food source for protozoa. The treatments were in triplicate and the populations of EcO157, protozoa, aerobic and coliform bacteria were monitored for 8 days.

Data analysis

Repeated-measures 2-way ANOVA (Prism 4.0; GraphPad Software, Inc., San Diego, CA) was used to compare differences in growth of protozoa, EcO157, coliforms, and aerobic bacteria in wastewaters. All comparisons were in triplicate. Bonferroni post hoc test was used to determine the earliest interval at which the decline of EcO157 or growth of protozoa differed significantly ( P<0.05) between various treatments.

Results

Protozoa in wastewater are stimulated by sequential inoculation of EcO157

These studies and microscopy experiments were facilitated with a GFP-labeled strain of EcO157 (MM123). After an initial 1-day lag, EcO157 populations declined rapidly with a decrease of 90% populations within 0.5 day for both inoculum levels, and also after the second re-inoculation to samples treated with the higher inoculum of EcO157 (). MPN counts indicated that each protozoan was provided initially with 14 or 11,200 cells of EcO157 for grazing and we anticipated that the higher level would support protozoan growth. Although no increase in protozoa was observed after the first treatment of 9×10 ⁷ cells of EcO157/mL (), a repeat inoculation at 7 days resulted in an 11-fold increase in protozoa at 9 days (an increase from 3.2×10 ⁴ to 3.5×10 ⁵ protozoa/mL; P<0.001) after the start of the experiment. EcO157 cells decreased from 2.1×10 ⁷ to 1.5×10 ⁴ (a 99.9% decrease) during the same period and resulted in a decline in ratio of EcO157 to protozoa from 122∶1 to 0.04∶1. The spike in aerobic bacterial counts ( P<0.001) at zero and 7 days was a result of enumerating the inocula as aerobic bacteria. At the low inoculum level of 10 ⁵ CFU EcO157/mL, the populations of protozoa and aerobic bacteria remained essentially the same as in uninoculated wastewater (not shown).

Internalization of EcO157 in wastewater protozoa

GFP-EcO157 (MM123) cells engulfed in a variety of predatory protozoa were observed by confocal laser scanning microscopy of wastewater samples. GFP-EcO157 cells were sparsely distributed in the surrounding wastewater at the lower inoculum level (10 ⁵ CFU/mL) and could not be detected inside protozoa ( Figure S1). However, at the higher inoculum level, the predominant protozoa that consumed EcO157 measured between 10 to 100 µm in diameter and appear to be ciliated. We captured numerous images of motile verticellid-like organisms ( Figure 2A to 2C ) that ingested the GFP-labeled cells within 2 h () after inoculation and remained internalized even after 7 days (). EcO157 cells abundant before day 3 () appear to be nearly absent from wastewater after 4 days ( Figure S2), but remained clustered inside some protozoa at day 7 (). Live bacteria in vesicles stained with Syto-59 (colored pseudo blue) were GFP-EcO157 (inset, ). Ten samples evaluated at 7 days confirmed the absence of GFP-labeled cells in the surrounding wastewater.

Internalization of EcO157 was confirmed from compiled optical sections illustrating that GFP-EcO157 cells were inside a protozoan (). However, it should be noted that GFP-EcO157 cells were observed also attached to the surfaces of protozoa ( Figure S3). We also observed the release of a vesicle containing GFP-labeled cells () from a vorticellid () packed with EcO157 cells within 3 days after grazing. Cross-sectional confocal laser scans of protozoa revealed that the food vacuoles were densely packed with GFP ( Figure 3A and 3B , GFP-panel). Although we observed an occasional flagellate that consumed EcO157 ( Figure S2), not every type of protozoa in wastewater consumed EcO157, even at the higher inoculum level. Some appeared refractile and the cyst-like structures did not consume any EcO157 throughout the incubations. We did not find any amoebae by either confocal or phase-contrast microscopy.

Preferential consumption of EcO157 by protozoa enriched from wastewater

Protozoa from wastewater were enriched in nutrient media and tested for their ability to consume GFP-EcO157 strain MM123 relative to other aerobic bacteria enriched from wastewater. Potential inhibitory compounds from wastewater [23] were minimized by supplementing CGM with only 5% of filter-sterilized wastewater. However, wastewater supplementation was required to prevent the rapid die-off of enriched protozoa from CGWM.

The number of EcO157 declined rapidly during the first 4 days in the presence of protozoa (), whereas they grew 5-fold on day 1 (an increase from 2×10 ⁷ to 1×10 ⁸ cells/mL) in the absence of protozoa and maintained at an elevated level of 5×10 ⁷ CFU/mL after 4 days. The number of aerobic bacteria remained stable during the first 3 days (3×10 ⁷ at day 3), whereas the cells of EcO157 were consumed preferentially (1×10 ⁵ at day 3). After 3 days, the protozoa switched to grazing aerobic bacteria after 99.6% of EcO157 cells were consumed. The ratio of aerobic bacteria to protozoa on the 3 rd day was 189,000∶1, whereas the EcO157 cells to protozoan ratio declined to 630∶1. However, protozoan numbers also declined within a day, but maintained at 79±3 cells/mL after the initial decline.

Predation of EcO157 by monocultures of protozoa isolated from wastewater

Protozoan diversity was observed to decrease from 8 to 10 morphological types to 5 types during the transition from natural wastewater to CGM supplemented with 5% filtered wastewater. Purification by serial dilutions and culturing in CGM resulted in the isolation of three ciliates characterized by 18S rRNA sequencing as Vorticella microstoma (GenBank accession # DQ868347), Platyophyra sp. ( EU039905) and Colpoda aspera ( EU039892). All the sequences obtained from 5 clones selected for each protozoan matched 100% with the GenBank sequences.

With GFP-EcO157 strain MM123 as the sole inoculated food source in PBS, Platyophyra and Colpoda consumed EcO157 cells in preference to the aerobic bacteria ( Figure 5B and 5C ) carried-over from wastewater. Addition of 50 µg/mL tetracycline to CGM used to prepare the protozoan inoculum did not eliminate any aerobic bacteria.

Although protozoan numbers maintained steadily throughout the incubations, Vorticella did not consume any EcO157 even after supplementation of 5% CGM (). However, supplementation of the Platyophyra and Colpoda samples resulted in significant consumption of EcO157 cells ( Figures 5B and C ). An initial 1.3-log decline of EcO157 cells was observed with Colpoda, followed by an additional smaller decline, but supplementation of 5% CGM on day 6 resulted in a further approximately 1-log decline of EcO157 at day 14 (). The number of Colpoda declined slightly also along with the decline of EcO157 populations after 8 days.

Although all three protozoa consumed GFP-EcO157 cells ( Figure 5A to 5C ), within 3 hours after inoculation of both EcO157 and protozoa to PBS, the majority of Vorticella, which appeared to be cysts with contractile stalks, did not ingest any GFP-EcO157 cells. Decreasing motility was observed during the 13-day incubation period, however, all motile vorticellids (, inset) consumed EcO157. Cross-sectional scans obtained for all three protozoa experiments confirmed that GFP-EcO157 were internalized ().

Aerobic bacterial numbers remained relatively stable throughout the incubation of EcO157 with Vorticella in PBS, but appeared to be consumed initially along with EcO157 cells by Colpoda. In contrast, Platyophyra began to consume more aerobic bacteria and EcO157 after a supplementation of CGM at day 3 followed by an increase in the aerobic bacteria after 6 days and a significant decrease in EcO157 ().

Nutrient addition to stimulate grazing of EcO157 by native protozoa

Protozoa isolated from wastewater required supplemental nutrients to aid in the consumption of EcO157 strain MM123 (). Similarly, supplementing wastewater with 10% LB broth () also increased significantly the growth of EcO157 strain MM149 and coliform bacteria after 2 days ( P<0.001). Addition of LB resulted also in a 3-fold increase in protozoan numbers (1.6×10 ⁵ with LB vs 5×10 ⁴ with out) after a 3-day incubation ( P<0.05). However, there was approximately a 4-log and 2-log higher decrease in EcO157 and coliforms at day 3, respectively, compared to the experiments with LB, indicating a corresponding increased fitness in E. coli and coliform bacteria in an environment with adequate nutrients (). No increase in protozoan growth was observed at a low inoculum of 7×10 ⁶ CFU/mL of EcO157. However, in both instances of supplementation of LB broth in the presence or absence of high inocula, the protozoan numbers increased significantly within 2 to 3 days, then declined to background levels corresponding to native aerobic bacterial counts. Both EcO157 and native coliform bacteria declined to undetectable levels after 8 days regardless of the presence or absence of LB. The spike in native aerobic bacteria after 2 days resulted from enumerating the increased growth of EcO157 as aerobic bacteria ().

Discussion

We have shown conclusively that protozoa in a dairy lagoon wastewater microcosm graze and grow on EcO157 and decrease significantly populations of EcO157. The protozoa that consumed EcO157 in wastewater were isolated for further characterization of their identities and individual activities with EcO157.

We observed by confocal microscopy a rapid intake of GFP-EcO157 within 2 h by protozoa, and the subsequent release of food vacuoles filled with live cells of EcO157 within 3 days after ingestion (). This result is similar to a previous report of the expulsion of food vacuoles containing EcO157 within 24 h after initiation of feeding by Glaucoma sp. isolated from store-bought lettuce [10]. Furthermore, we observed internalized GFP-EcO157 cells in some protozoa after 7 days, whereas GFP-EcO157 cells were absent from wastewater within 5 days after inoculation (). This differs from an earlier finding of growth of EcO157 during 35 days in the presence of axenically grown A. polyphaga or its lysates [18]. Differences in results with different protozoa, pathogen strains and conditions emphasize the importance of further work to determine whether protozoa are a serious reservoir for some pathogens and can protect them from destruction by environmental stresses and, especially, whether hypervirulent pathogen phenotypes are selected and/or they replicate inside protozoa and their vacuoles for subsequent ejection into the agricultural environment.

We demonstrated incremental growth of native protozoa with sufficient populations of EcO157. This would be consistent with our previous study with E. coli ONT:H32 (MM158) inoculated four times sequentially into wastewater from dairy lagoons over a 45-day period [6]. In the current study, predation of EcO157 (MM123) was correlated with the incremental growth of protozoa above the level noted for growth obtained on native bacteria alone (). Furthermore, an 11-fold increase in protozoan numbers was observed after they had consumed 99.9% of the second inoculum of 2×10 ⁷ EcO157 CFU/mL. Rates of consumption of bacteria by protozoa of 9 to 266 per hour for a flagellate and 200 to 5,000 per hour for a ciliate have been reported previously [29]. This corresponds to a protozoan population of 10 ⁴ consuming from 10 ⁶ to 10 ⁹ bacteria per day and, therefore, is consistent with protozoan increases after the second inoculation due to acclimation of protozoa to the EcO157 strain tested in this study. These results are in agreement with a recent observation of increased growth of protozoa in co-culture with EcO157, S. enterica and Listeria monocytogenes[10]. Since the protozoan numbers increased after only one re-inoculation in this study, we speculate that the significant decrease of 90% of E. coli ONT:H32 cells from 9.7 to 0.9 days after the 4 th re-inoculation [6] resulted from enhanced consumption of E. coli by specific protozoa [30]. This suggests adaptation of the protozoa to a specific bacterial strain present in high numbers compared to other natural strains present in the wastewater. Potential adaptation of predator biology of protozoa to bacteria prevalent in an environment, especially bacterial pathogens, would be an intriguing area for further study.

To distinguish protozoan predation from other biotic and abiotic factors, we confirmed that protozoa enriched from wastewater and grown in CGWM were responsible for a significant decrease in the population of outbreak strain of EcO157 (MM123). Furthermore, EcO157 grew in the absence of protozoa and maintained at ∼8 log CFU/mL. Direct evidence for predation of EcO157 in wastewater has not been reported, however, extended survival has been reported for EcO157 in filtered well water [22], in autoclaved wastewater from dairy lagoons [23] and cycloheximide-treated cattle trough water [31].

Protozoa enriched from wastewater consumed EcO157 preferentially () in comparison to the background aerobic bacterial populations in CGWM. Native bacteria were consumed as alternate prey when the preferred food source, EcO157, declined to a threshold level [32] of 10 ⁵ cells/mL after a 3-day incubation. If the predation is merely density dependent [33], aerobic bacteria would have been preferentially consumed during the first 3 days. Although protozoa are known to consume gram negative bacteria preferentially over gram positives [34], [35], this is the first report that protozoa from dairy lagoons consume EcO157 preferentially over natural microflora.

We have isolated and identified V. microstoma, Platyophyra sp. and C. aspera by culturing serial dilutions of the protozoan enrichments in CGWM. Apparently these protozoa in wastewater from dairy lagoons did not originate from the ruminants [36], [37], as suspected. It is likely that mixing of enrichments on a shaker might have eliminated the anaerobic ciliates Entodinium and Epidinium that are most prevalent in the cow rumen [37]. Distinct feeding patterns were detected for each protozoan in consuming EcO157 in PBS (). Initially, C. aspera consumed simultaneously both EcO157 and aerobic bacteria carried over from wastewater, in contrast to Platyophyra sp. requiring a supplemental addition of 5% CGM as nutrients to initiate consumption of EcO157 and aerobic bacteria. However, V. microstoma, a common inhabitant of activated sludge systems [38], ingested (, inset), but failed to digest EcO157 during the 13 days of incubation. Neither the cysts of Vorticella () nor any other cysts observed in wastewater samples contained GFP-EcO157. We detected motile vorticellids with vesicles filled with GFP-EcO157 after 7 days in wastewater () that was devoid of free EcO157 cells. It is possible that Shiga toxins [17] provided a selective advantage in resisting the digestion by Vorticella, in contrast to Platyophyra and C. aspera unaffected by Shiga toxin in consumption of EcO157. However, both Platyophyra and Colpoda failed to eliminate EcO157, possibly due to the exhaustion of supplemental nutrients required to stimulate grazing or for reaching a threshold level for consumption by protozoan cells, as observed for the efficient consumption of non-pathogenic E. coli by T. pyriformis[32].

The ability of EcO157 to be ingested and survive for 13-days in co-culture with Vorticella, Platyophyra and Colpoda support an earlier report of a protective role for protozoa during the transport of pathogenic bacteria to environments conducive for re-growth [11]. Indeed, a recent study reported that EcO157 released from expelled food vacuoles grew in spinach extracts [10] that represent potential nutrients present on leaf surfaces as a result of tissue damage or cut edges during or prior to harvest of produce [39]. Such a role for protozoa in sequestering bacterial pathogens in vacuoles resulting in enhanced survival has been reported also for S. enterica[10], [11], S. typhimurium[40], M. avium[15], L. pneumophila[41], [42], Staphylococcus aureus (MRSA) [43] and others [44]. If coexistence of pathogenic E. coli and predatory protozoa as observed in nutrient and alternate prey rich wastewater supplemented with CGM occurs in nature, the protozoa could be an effective refuge for bacteria in general, and foodborne pathogens in particular, thus serving as an environmental reservoir, as has been reported for coexisting M. avium and A. polyphaga[45]. Since motile cells of V. microstoma ingested EcO157 without digesting them (), they are candidates for "storing" and/or transporting pathogenic EcO157 in dairy environments.

In contrast to the failure of protozoan grazing at a certain critical density of prey discussed above, populations of EcO157 ( and ) inoculated into wastewater directly were eliminated without a noticeable threshold for the consumption of EcO157. This is predictable considering we observed 90% decline rates for several strains of EcO157 within 1 to 10 days in wastewater from different dairy lagoons in microcosms; EcO157 cells were eliminated in less than 4 weeks [6]. Since protozoa isolated from wastewater failed to eliminate EcO157 ( and ), we suspect other components of wastewater may be responsible for decreases, such as lytic phages [46], bdellovibrios [47], [48] and inhibitory chemicals [23], emphasizing the complex microbial ecology that may affect bacterial pathogens in the environment and food production systems.

Since supplemental nutrients aid in decreasing the populations of EcO157 by Platyophyra and Colpoda () and other protozoa in wastewater (), attempts to augment beneficial protozoa in dairy lagoons could be a novel approach for on-farm control of zoonotic pathogens in wastewater before the water is distributed to other locations (feed crops, orchards, watersheds, etc.) or contaminates crops grown in proximity. Protozoa in feces and entering wastewater are affected by animal diet. High-grain diets [49], [50] often result in the decrease of protozoan cells and diversity of species. In addition, EcO157 declined rapidly in manure from cattle fed with 100% straw as compared with digestible silage supplemented with soy and maize concentrates [51]. Although, cattle diet might increase the beneficial protozoa that consume EcO157 in dairy lagoons, it could also result in stimulating protozoa that may protect and transport EcO157 to crops grown nearby to dairies.

To our knowledge, this is a first report of characterization of protozoa native to dairy lagoons that are capable of consuming but not eliminating the populations of EcO157. These protozoa are not reported to be common inhabitants of dairy animals, but they were capable of grazing on EcO157. Grazing of EcO157 by two of the three isolates of protozoa resulted in decreases in the populations of EcO157. Since all three protozoa failed to eliminate EcO157, but retained them in food vacuoles, a potential exists for EcO157 in protozoan vacuoles to re-colonize cattle in free-stall dairies that use recycled water commonly for washing lanes. This would result in a complex ecology and possible evolution of EcO157 or other zoonotic pathogens in protozoa in different wastewater conditions (nutrients, fecal load) and different cows. This complex ecology could involve specific interactions between protozoa and pathogens, elimination or protection of pathogens in protozoa, and protozoa adaptations specific to predominant bacteria in the agricultural environment. Further studies will be required to determine the reproducibility of these complex interactions, the detection of other factors and, potentially, the identification of novel interventions beneficial for food safety.

Our study provides new information about the risks associated with run-off from dairies and ranches [8] that could contaminate potentially nearby crops and orchards and possibly yield commodities that could become a major public health issue.

Supporting Information

Method S1

Characterization of monocultures of protozoa by 18S rRNA sequencing

(DOC)

Figure S1

Confocal image of a protozoan taken 2 h after treating the wastewater with 1×10 ⁵ CFU of GFP-EcO157/mL. The compiled image was obtained with transmitted light, propidium iodide and GFP. Protozoa did not contain any internalized GFP-EcO157 cells.

(TIF)

Figure S2

Image of a flagellate taken 4 days after inoculating the wastewater with 9×10 ⁷ CFU of GFP-EcO157/mL. The flagellate ingested and retained live cells of EcO157 within food vacuoles although the surrounding wastewater contained little or no GFP-EcO157.

(TIF)

Figure S3

A protozoan with GFP-labeled cells attached to the surface. Confocal image was taken 2 h after inoculating the wastewater with 9×10 ⁷ CFU of GFP-EcO157/mL.

(TIF)

Acknowledgments

We thank Anna Bates, Robert Hnasko and Michael Cooley for guidance on confocal microscopy, and Anna Engelbrektson for help on statistics.

Footnotes

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

Funding: The work was funded by the U. S. Department of Agriculture, Agricultural Research Service under Current Research Information System projects 5325-32000-005 and 5325-42000-044. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

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# data[].attributes.attachment_link.title
Identification of Protozoa in Dairy Lagoon Wastewater that Consume Escherichia coli O157:H7 Preferentially

# data[].attributes.attachment_link.description
Escherichia coli O157:H7 (EcO157), an agent of life threatening hemolytic-uremic syndrome, resides in ruminants and is released in feces at numbers as high as 10 million cells/gram. EcO157 could survive in manure for as long as 21 months, but we observed ...

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MOLECULAR IDENTIFICATION OF PATHOGENIC FUNGI

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MOLECULAR IDENTIFICATION OF PATHOGENIC FUNGI

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Isolation of Anaerobic cellulose-decomposing bacteria from soil

# data[].attributes.subject
Isolation of Anaerobic cellulose-decomposing bacteria from soil

# data[].attributes.attachment_link.title
Microbiology Society Journals | The Isolation of Anaerobic Cellulose-decomposing Bacteria from Soil

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SUMMARY: A strictly anaerobic mesophilic species of cellulose-decomposing bacteria was isolated from soil by a new technique. Cultures were made in screw-capped bottles with media containing finely divided cellulose. The method and apparatus used for filling these bottles with reduced media and nitrogen is described.

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MOLECULAR IDENTIFICATION OF FUNGI FROM RED SEA

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MOLECULAR IDENTIFICATION OF FUNGI FROM RED SEA

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https://ac.els-cdn.com/S1815385216300384/1-s2.0-S1815385216300384-main.pdf?_tid=b82d605a-f20f-446d-aa1a-6414037b9833&acdnat=1519975869_4cb011fa9d705c58af5d7a0c81307b40

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Detection and characterization of

Shigella species isolated

from food and human stool samples in Nabeul, Tunisia, by

molecular methods and culture techniques

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Detection and characterization of Shigella species isolated from food and human stool samples in Nabeul, Tunisia, by molecular methods and culture techniques - Mokhtari - 2012 - Journal of Applied Microbiology - Wiley Online Library

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Rapid identification of fungi by sequencing the ITS1 and ITS2 regions using an automated capillary electrophoresis system

# data[].attributes.subject
Rapid identification of fungi by sequencing the ITS1 and ITS2 regions using an automated capillary electrophoresis system

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https://www.tandfonline.com/doi/abs/10.1080/13693780310001600435

# data[].attributes.headline
Identification of fungi from dairy products by means of 18S rRNA analysis

# data[].attributes.subject
Identification of fungi from dairy products by means of 18S rRNA analysis

# data[].attributes.attachment_link.title
Identification of fungi from dairy products by means of 18S rRNA analysis - ScienceDirect

# data[].attributes.attachment_link.description
The role of fungi as cause of spoilage of dairy products, such as cheese and yoghurt, has been clearly demonstrated. Despite of this, there is still a lack in rapid methods for the identification of food-associated fungi.

# data[].attributes.headline
Identification of fungi in the Gaeumannomyces-Phialophora complex by RFLPs of PCR-amplified ribosomal DNAs

# data[].attributes.subject
Identification of fungi in the Gaeumannomyces-Phialophora complex by RFLPs of PCR-amplified ribosomal DNAs

# data[].attributes.attachment_link.title
Identification of fungi in the Gaeumannomyces-Phialophora complex by RFLPs of PCR-amplified ribosomal DNAs - ScienceDirect

# data[].attributes.attachment_link.description
The polymerase chain reaction (PCR) was used to amplify ribosomal internal transcribed spacer and 5·8S DNA from isolates of Gaeumannomyces graminis, Phialophora graminicola and other root-infecting fungi, isolated mostly from cereals and grasses. Different restriction enzymes were used to digest these amplified rDNAs to find polymorphisms useful in identification.

# data[].attributes.attachment_link.title
Microscopic and Molecular Studies of the Diversity of Free-Living Protozoa in Meat-Cutting Plants

# data[].attributes.attachment_link.description
In order to investigate the role of environmental free-living protozoa in the contamination of food by food-borne pathogens, an inventory of these unicellular eukaryotes in food-processing environments is a necessary first step. Although the bacterial flora of meat-processing environments has been described, to our knowledge, no studies of the environmental protozoan communities in these environments have been performed previously.

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http://jwh.iwaponline.com/content/ppiwajwh/13/4/953.full.pdf

# data[].attributes.headline
identification and characterization of RNA viruses

# data[].attributes.subject
identification and characterization of RNA viruses

# data[].attributes.attachment_link.readable_text
Guillaume Carissimo,

Karin Eiglmeier,

Julie Reveillaud,

Inge Holm,

Mawlouth Diallo,

Diawo Diallo,

Amélie Vantaux,

Saorin Kim,

Didier Ménard,

Sovannaroth Siv

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Abstract

Mosquitoes of the Anopheles gambiae complex display strong preference for human bloodmeals and are major malaria vectors in Africa. However, their interaction with viruses or role in arbovirus transmission during epidemics has been little examined, with the exception of O'nyong-nyong virus, closely related to Chikungunya virus. Deep-sequencing has revealed different RNA viruses in natural insect viromes, but none have been previously described in the Anopheles gambiae species complex. Here, we describe two novel insect RNA viruses, a Dicistrovirus and a Cypovirus, found in laboratory colonies of An. gambiae taxa using small-RNA deep sequencing. Sequence analysis was done with Metavisitor, an open-source bioinformatic pipeline for virus discovery and de novo genome assembly. Wild-collected Anopheles from Senegal and Cambodia were positive for the Dicistrovirus and Cypovirus, displaying high sequence identity to the laboratory-derived virus. Thus, the Dicistrovirus ( Anopheles C virus, AnCV) and Cypovirus ( Anopheles Cypovirus, AnCPV) are components of the natural virome of at least some anopheline species. Their possible influence on mosquito immunity or transmission of other pathogens is unknown. These natural viruses could be developed as models for the study of Anopheles-RNA virus interactions in low security laboratory settings, in an analogous manner to the use of rodent malaria parasites for studies of mosquito anti-parasite immunity.

Citation: Carissimo G, Eiglmeier K, Reveillaud J, Holm I, Diallo M, Diallo D, et al. (2016) Identification and Characterization of Two Novel RNA Viruses from Anopheles gambiae Species Complex Mosquitoes. PLoS ONE 11(5): e0153881. https://doi.org/10.1371/journal.pone.0153881

Editor: Eric Jan, University of British Columbia, CANADA

Received: December 20, 2015; Accepted: April 5, 2016; Published: May 3, 2016

Copyright: © 2016 Carissimo 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: All sequence files are available from the EBI European Nucleotide Archive database ( http://www.ebi.ac.uk/ena/) under study accession number ERP012577, and sample accession numbers: ERX1180340, ERX1180341, ERX1180342, ERX1180343, ERX1220645, ERX1220646 and ERS977505). All assembled sequences are available from NCBI (accession numbers: KU169878, KU169879 and KU169880). Study Galaxy workflows are available at: ( https://mississippi.snv.jussieu.fr/u/drosofff/p/anopheles-viruses).

Funding: This work received financial support to KDV from the European Research Council ( http://erc.europa.eu/funding-and-grants), Support for frontier research, Advanced Grant #323173 AnoPath; European Commission ( http://ec.europa.eu/research/fp7/index_en.cfm), FP7 Infrastructures #228421 Infravec; and French Laboratoire d'Excellence ( http://www.enseignementsuprecherche.gouv.fr/cid55551/investissements-avenir-projets-labex-par-regiondomaine.html), and "Integrative Biology of Emerging Infectious Diseases" #ANR-10-LABX-62-IBEID. This work received financial support to CA from the Agence Nationale de la Recherche grant ANR-13-BSV2-0007 " Plastisipi ".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

Insect specific RNA viruses from various families (i.e. Flaviviridae, Togaviridae, Bunyaviridae, Rhabdoviridae, Mesoniviridae) and the taxon Negevirus have been described in Anopheles mosquitoes [ 1- 5]. These viruses have been discovered by isolation from cell cultures, by RT-PCR and manual sequencing targeting regions of known viruses, or using deep sequencing on field caught insect samples [ 6, 7]. The siRNA pathway of mosquitoes is involved in the interaction and processing of the viral double strand RNA (dsRNA) intermediates produced by RNA viruses [ 8- 14]. Deep sequencing of small RNA and bioinformatics has been used to reconstruct active novel viruses in plants, Drosophila or mosquitoes, using detection of viral-derived small interfering RNAs (viRNAs) as a criterion for active replication [ 7, 15, 16]. Small RNA deep sequencing should allow sensitive detection and discovery of viruses that produce dsRNA intermediates. Alternately, viral discovery by inoculation of extracts onto cell lines is not limited to detection of dsRNA intermediates, but can be biased due to differential efficiency of viral replication across cell lineages [ 17- 19].

Among Anopheles mosquitoes, the most mature reference genome sequence and genomic tools have been developed for the An. gambiae complex, particularly the sister taxa An. gambiae and An. coluzzii, and these are currently the most powerful Anopheles models. Within the African malaria vectors of the An. gambiae complex, to our knowledge no natural RNA viruses have yet been described, with the exception of O'nyong-nyong virus (ONNV), a pathogenic arbovirus transmitted to humans [ 20], which is a close relative of Chikungunya virus transmitted by Aedes mosquitoes [ 21, 22]. Consequently, research on viral response and antiviral immunity of Anopheles has been limited to ONNV infections [ 14, 23- 27], but research using ONNV requires sophisticated biosecurity conditions. Identification of Anopheles RNA viruses that could be used as low-biosecurity model systems would facilitate studies of Anopheles antiviral immunity and mosquito-virus interactions.

In the current study, we used small RNA deep-sequence datasets to reconstruct more than 90% of a novel Dicistrovirus, and several segments of a previously unknown Cypovirus. We verified their infection prevalence within the Anopheles colony where they were discovered. Oral infection experiments were used to evaluate the species-specificity of these newly discovered viral strains and their ability to be transmitted to the progeny. Finally, we detected both viruses in wild Anopheles mosquitoes captured in Cambodia and in Senegal, confirming that they are components of the natural virome.

The results indicate that a laboratory Anopheles colony, widely used to study vector interactions with different pathogens, carries previously undetected natural insect viral infections that could potentially influence the results of such studies. These novel insect viruses also could become valuable tools to study Anopheles antiviral immunity, as well as the underlying mechanisms of vector interactions with RNA viruses, without the requirement for high biosecurity infrastructure or procedures.

Results

Four datasets of small RNA sequence generated from pooled An. coluzzii (Ngousso colony) mosquito midguts were analyzed using the Metavisitor pipeline [ 28], a suite of open source bioinformatic tools designed to simplify virus diagnostic, discovery and genome reconstruction on a Galaxy framework [ 29]. Two novel viruses were detected in the mosquitoes, a Cypovirus (genus: Cypovirus, sub-family: Spinareovirinae, family: Reoviridae), and a Dicistrovirus (genus: Cripavirus, family: Dicistroviridae, order: Picornavirales).

Detection of a Cypovirus

We detected a transcript (accession number KU169880) of 892 nt aligning to segment 10 of Culex restuans Cypovirus (CrCPV)[ 30] and to Uranotaenia sapphirina Cypovirus (UsCPV)[ 31] with 91% and 83% identity, respectively. Multiple alignments of the nucleic sequences with Clustal O [ 32] indicated that it was not a chimeric segment of the two known viruses. This segment possesses an intact 237 aa ORF that codes for a complete polyhedrin protein with 96% and 90% amino acid identity to the polyhedrin of CrCPV and UsCPV, respectively. Cypovirus genomes are comprised of 10 to 11 segments of double strand RNA, each coding for a different viral protein. The polyhedrin, protein, coded by segment 10, is highly conserved among Cypoviruses.

A second contig of 3889 nt was reconstructed (accession number KU169879) containing an ORF coding for a 1246 aa protein that has 34% identity with the RNA dependent RNA polymerase (RdRp) of Bombyx mori Cypovirus 1 (ACX54961.1) over 98% of its sequence. Realignment of the original small RNA reads to the contig shows a dicing profile with the read length peaking at 21nt in all four datasets ( Fig 1). This is the characteristic product of viral replication intermediates processed by the siRNA pathway [ 33], indicating that the reconstructed sequences belong to an actively replicating virus infecting the mosquitoes.

The four small RNA datasets were independently aligned to the sequence of Anopheles Cypovirus segment 1 (accession number KU169879) with bowtie allowing no mismatches. From the bowtie output, lattice maps and read size histograms were generated representing the positions and size of the small RNAs that align to the viral genomic segment.

Phylogenetic analysis of the predicted polymerase and polyhedrin peptide sequences were performed using the available sequences for the different Cypovirus species ( Fig 2). Neighbor-joining trees show that reconstructed RdRp and polyhedrin sequences are related but distinct from known Cypoviruses. The CPVs have been classified into 16 types by the International Committee on Taxonomy of Viruses [ 34], or 21 types based on the electrophoretic migration patterns of their dsRNA genome segments [ 35]. Therefore, it is not surprising that the phylogeny of these viruses inferred from the polymerase sequence, and to a lesser extent from the polyhedrin gene, does not reflect the virus types classification. We conclude that the reconstructed sequences are evidence of a previously unknown virus, here named Anopheles Cypovirus (AnCPV). We hypothesize that AnCPV is a segmented (10 to 11 segments) double stranded RNA virus of the genus Cypovirus, sub-family Spinareovirinae, family Reoviridae. Based on the phylogenetic tree on the polyhedrin gene, this virus could be a novel member of the Cypovirus type 17 [ 35].

Light microscopy of An. coluzzii larval midguts displayed gross pathological signs of Cypovirus infection, detectable as localized clusters of tightly packed inclusion bodies in the midgut epithelium that appear visually as iridescent blue-white features [ 31] (white arrows, Fig 3). RT-PCR primers were designed based on the sequence of segments 1 and 10 and successfully used to amplify fragments from viral RNA templates in the total RNA used for the original small RNA library preparation. Sanger sequencing of the amplicons showed no variation from the bioinformatically reconstructed genomic sequences ( S1 Zipped Archive).

Signs of Cypoviral infection are white-blue iridescent areas as indicated by the white arrows in each panel. Each panel corresponds to a different mosquito larva.

Detection of a Dicistrovirus

Other contigs reconstructed from the same small RNA datasets displayed significant similarity using blastx to Drosophila C virus (DCV) and Cricket paralysis virus (CrPV) from the Distroviridae family. Using additional long paired-end RNAseq datasets (accession number ERS977505), we were able to reconstruct a 8919 nt viral genomic sequence that completely includes these contigs and contains two ORFs (accession number KU169878). The first ORF (5' ORF) codes for an 1802 aa (nucleotides 391 to 5799 in the reconstructed genome) polyprotein with 48% blastp identity to DCV nonstructural polyprotein. The second ORF (3' ORF) codes for a predicted 852 aa protein (nucleotides 6067 to 8625 on the reconstructed genome) and displays 54% blastp identity to DCV structural polyprotein. This dicistronic organization of the reconstructed genome is typical of Dicistroviridae with an ORF1 that encodes for non-structural proteins and an ORF2 that encodes for virion proteins (reviewed in [ 36]). Dicistroviridae are related to the Picornaviridae, which encode the same proteins in a monocistronic manner as one polyprotein, and to the Comoviridae, which encode the two ORFs but on two monocistronic RNAs.

Realignment of the small RNA reads on the reconstructed genome revealed, as for AnCPV, a dicing profile of virus replication intermediates, yielding virus-derived siRNAs with a read length peaking at 21 nt in all four datasets ( Fig 4). These results are evidence for a previously unknown, actively replicating virus.

The four small RNA datasets were independently aligned to the sequence of the Anopheles C virus (AnCV) genome (accession number KU169878) with bowtie allowing no mismatches. From the bowtie output, lattice maps and read size histograms were generated representing the positions and size of the small RNAs that align to the viral genome.

Phylogenetic tree reconstruction of non-structural proteins from Dicistroviridae shows that the discovered virus is related to known Dicistoviruses and groups with the Cripavirus genus ( Fig 5). Similar to the detection of AnCPV, RT-PCR primers, designed on the reconstructed AnCV sequence, were also successfully used to generate amplicons from the total RNA used for small RNA library preparation. Sanger sequencing of the amplified region showed no variation from the bioinformatically reconstructed genomic sequence ( S1 Zipped Archive). Because a paralysis phenotype was not observed in the source mosquito colony, we inferred that the virus is most likely a C-type Dicistrovirus rather than a paralysis virus and named it Anopheles C virus (AnCV).

We investigated whether, in addition to the two ORFs at similar positions found in other Dicistroviruses, AnCV also possessed an Internal Ribosome Entry Site (IRES) of the same class and family as observed in other Dicistroviruses in order to translate the second ORF. Searching the RNA family database (Rfam) [ 37], we identified a 200nt region (position 5795 to 5994) of AnCV as a potential IRES, with a p-value of 9.4e-33 for the RF0458 CrPV IRES family ( S1 File). Multiple alignment of the IRES sequences of the RF0458 family and AnCV ( S2 File) showed RNA secondary structure conservation ( Fig 6A) and covariance of the nucleotides within the structured pairs ( Fig 6B). These results indicate that this region of the AnCV genome, situated between the two ORFs, is structurally conserved and has a high probability of being a true functional IRES, validating the result of the Rfam search.

Cypovirus and Dicistrovirus biological evidence

Cloned virus fragments were used to synthesize RNA probes for hybridization to a Northern blot of the total RNA used for small RNA sequencing ( Fig 7). The probes with sequence homology to the AnCV genome and the segment 1 (RdRp) of AnCPV revealed the presence of intact genomic viral RNA in the total RNA from mosquito pools for both viruses. This result also rules out RNA degradation as a source of the small RNAs detected by deep sequencing, and strengthens the interpretation of the dicing patterns (Figs 1 and 4) as a product of RNA from actively replicating virus.

In order to assess the infection prevalence and pattern in the An. coluzzii colony, we examined individual mosquitoes and individual mosquito organs by RT-PCR using virus-specific primers ( Table 1). The results indicated that adult mosquito midguts, carcasses and salivary glands were susceptible to infection by both AnCPV and AnCV. Dissected tissues were washed extensively in PBS to minimize contamination, and negative controls were included.

Mosquitoes displayed variable intensity of the RT-PCR amplification products on agarose gels. Although not a quantitative assay, this observation suggests that individual mosquitoes harbored different levels of infection. Mosquitos were infected by one, both or neither of the two viruses ( S1 Table). Some mosquitoes were PCR-positive for AnCPV in the carcass but negative for the midgut, suggesting that infection in those mosquitoes might have been resolved in the midgut or that transmission might have been vertical. The reason for differences in the AnCV prevalence between batch 1 and 2 could be explained by multiple factors such as age of mosquitoes since emergence, or blood feeding status. Detailed studies will be needed to clarify the tropism and temporal dynamics of infection in mosquitoes.

Mosquito host range and transmission

Other colonies of An. coluzzii (Fd03, Fd05, Fd09) and An. gambiae (Mbita) tested by RT-PCR were consistently infected with both viruses, but the significance of the observation is unclear because they were all maintained in the same insectaries as the Ngousso An. coluzzii colony at Institut Pasteur Paris. Samples tested from a colony of An. dirus (Pursat) maintained at Institut Pasteur Cambodia were also positive for both viruses, but the history of other insectaries where the colony was previously maintained is unknown (although it has not been kept in the Paris facility).

Interestingly, we did not detect either virus in a colony of An. stephensi (SDA500) maintained in the same Institut Pasteur Paris insectary as the An. coluzzii Ngousso colony, nor in two Aedes aegypti isofemale lines. Since oral transmission is the natural route of DCV infection [ 38] as well as of the two known mosquito Cypoviruses [ 30, 31], we tested whether the novel viruses are transmissible orally to these uninfected mosquito species. We reared eggs from the virus-negative An. stephensi and the two Ae. aegypti lines and divided first instar larvae of each into two trays. One tray of larvae was supplemented with homogenized An. coluzzii Ngousso fourth instar larvae, carrying AnCV and AnCPV viruses, while the other tray of larvae was fed normally and maintained in the same insectary as a control. An. stephensi and Ae. aegypti fourth instar larvae and adult females were then collected and assayed for AnCV and AnCPV as pools or individually. Larvae were washed in multiple changes of clean water and starved over several hours before processing in order to minimize the possibility of external contamination or midgut content contamination. This per os exposure did not infect the Ae. aegypti mosquitoes, but successfully infected An. stephensi mosquitoes with both AnCV and AnCPV at a prevalence of 59% and 100% respectively, while mosquitoes issued from the control trays remained negative ( Table 2).

To assess whether the viruses can be transmitted vertically, An. stephensi adult females developed from larvae experimentally infected as above (the F0 generation) were allowed to blood feed and lay eggs (F1 generation). Both F1 larvae as well as resulting adult females displayed AnCPV and AnCV infections ( Table 3). These results suggest that both of the viruses can be transmitted vertically to the progeny by oviposition. An. stephensi mosquitoes maintained until the F4 generation after virus exposure displayed persistent infection with AnCPV, but AnCV prevalence decreased each generation and was absent in F4 adults.

Virus presence in wild Anopheles mosquitoes

We captured wild Anopheles spp. mosquitoes in villages in Senegal and Cambodia, and tested individual mosquito RNAs for the viruses using specific RT-PCR. Bands of the correct predicted sizes were detected in some of the samples: 494nt for AnCV and 524nt for AnCPV ( Fig 8). Samples were processed along with negative controls in order to control for cross-contamination. Individual wild mosquitoes were positive for one, both, or neither virus.

One wild Anopheles spp. from Senegal was positive for AnCV, and six wild Anopheles spp. from Senegal and Cambodia were positive for AnCPV. The resulting amplicons were sequenced in both orientations. In addition, AnCV amplicons were detected in two An. dirus mosquitoes from the Cambodia insectary colony, and were also sequenced. Alignment of these sequences with the reconstructed consensus sequences showed high sequence homology with a maximum of three SNPs for AnCPV and one SNP for AnCV in the analyzed region ( S3 and S4 Figs). The results indicate that AnCV is present in the natural virome of wild Anopheles mosquitoes from at least Africa, and AnCPV is present in the natural virome of wild Anopheles mosquitoes from Africa and Asia.

Discussion

By deep sequencing of small RNAs we have identified two novel insect viruses, a Cypovirus and a Dicistrovirus, in a laboratory colony of An. coluzzii mosquitoes. Reconstructing viral genomes from viral-derived small RNAs enriches for detection of actively replicating viruses. We present biological evidence for the presence of these viruses as a persistent infection in colonies of three species of Anopheles ( An. coluzzii, An. gambiae, An. dirus) and their exclusion from a colony of one other species ( An. stephensi). Because of the possibility of horizontal transmission in the insectary, data from colonies are not informative for infection in nature, but it does at least indicate the species range of permissive hosts. Importantly, the viruses were present in wild Anopheles mosquitoes captured in Senegal and Cambodia, thus establishing them as components of the natural virome.

We identified Anopheles Cypovirus (AnCPV), which contains a genomic segment coding for the polyhedrin protein characteristic of this viral family and a genomic segment coding for an RNA-dependent RNA polymerase that shows conservation with the genomes of known Cypoviruses. This is, to our knowledge, the first reported member of the Cypoviridae family infecting Anopheles mosquitoes, complementing previous studies identifying Cypoviruses in other mosquito genera [ 30, 31]. We also identified an Anopheles C virus (AnCV) that displays genetic architecture consistent with other members of the family Dicistroviridae, with two open reading frames, 5' non-structural polyprotein and 3' structural polyprotein, as well as conserved RNA secondary structure and sequence of the central IRES. This virus was further confirmed to belong to the Dicistroviridae family based on the amino-acid sequence conservation of the AnCV RNA-dependent RNA polymerase with its orthologs of the Drosophila C virus (DCV) and Cricket paralysis virus (CrPV).

The novel Dicistrovirus AnCV can infect some Anopheles but not Aedes mosquitoes, and appears not to induce high mortality nor detectable paralysis phenotypes in mosquitoes infected per os. These data suggest that infection phenotype and range of this virus resemble that of DCV and not CrPV. We cannot exclude that in our model, the virus and the laboratory colony could have co-evolved to minimize pathogenic symptoms. The virus-negative An. stephensi colony, when infected with AnCV per os, also did not display detectable pathogenic symptoms. This scenario could be tested with isolated viruses by infection experiments using different concentrations of virus to study symptoms and pathogenicity. Isolated and culturable viruses would also allow studies of virion morphology, mechanisms of entry, host range, obtaining complete genomic sequences, and generating infectious clones. The main mosquito organs were found to be positive for both viruses. Development of a quantitative real-time PCR assay would be useful in order to measure genome copies in different organs.

DCV, first isolated in 1972 [ 39], and CrPV are models for the study of insect antiviral immunity. These viruses as well as other members of the family are ecologically and agriculturally relevant [ 40]. The discovery of a virus from this family in laboratory and wild anopheline mosquitoes is an important first step to understanding the natural viral population in these major human malaria vectors, and potentially establishing a viral infection model in those mosquitoes.

It will be interesting to determine whether AnCV has similar phenotypes for Anopheles mosquitoes as the model insect virus DCV has for Drosophila. For example, DCV leads to a strong immune response and to fly death within several days when intra-thoracically injected [ 41], but produces only mild effects after infection by the oral route, with transcriptional induction of only 11 genes [ 42]. Although the small transcriptional response may suggest a co-evolved state, DCV has also been associated with population crashes and shorter longevity of some laboratory colonies, even in persistently infected fly stocks [ 40].

The first infectious clone of a Dicistrovirus, CrPV [ 43], was recently generated, which suggests that an AnCV infectious clone as a tool is within reach. This would open an array of possibilities to study RNA virus antiviral mechanisms in the malaria vector mosquito under low-biosecurity conditions. The discovery of the first RNA viruses in the An. gambiae species complex is an initial step that will facilitate work on mosquito-virus interactions and anti-viral immunity, including the influence of the microbiota as well as potential reciprocal interactions with other pathogens such as malaria parasites.

Materials and Methods

Mosquito colonies

Deep sequencing of small RNAs used mosquitoes from the An. coluzzii Ngousso colony previously established from Yaoundé, Cameroon [ 44], and formerly described as An. gambiae M molecular form. Other colonies used and raised in the Institut Pasteur Paris insectaries were An. gambiae Mbita from Kenya (obtained from the ICIPE Field Station, Mbita Point, Kenya), Fd03 initiated from Mali, Fd05 and Fd09 from Burkina Faso, and An. stephensi colony SDA500. In addition, two Aedes aegypti isofemale lines from Thailand were used [ 45]. All mosquitoes were reared under standard conditions at 26°C and 80% relative humidity, with a 12 h light/dark cycle as previously described [ 44]. Finally, mosquito samples from the An. dirus Pursat colony reared in the insectaries of Institut Pasteur Cambodia, and never grown at Institut Pasteur Paris, were also used.

Wild mosquitoes

Wild Anopheles mosquitoes were used for testing of virus presence in nature. Mosquitoes were captured in Ratanakiri and Kampong Chnang, Cambodia and Kedougou, Senegal were visually identified as Anopheles spp., and immediately transferred into RNAlater at 4°C, and then stored at -80°C. Permission to collect mosquitoes was obtained by Institut Pasteur Cambodia from authorities of Ratanakiri and Kampong Chnang, and by Institut Pasteur Dakar from authorities of Kedougou. RNA was extracted from individual mosquitoes using the Nucleospin miRNA kit (Macherey-Nagel) following the supplied protocol. RT-PCR assays for AnCV and AnCPV amplicons were performed as described below (using 45 PCR cycles), with blank reactions and negative control RNA samples. Amplicons from positive samples were Sanger sequenced and aligned with virus sequences assembled from deep sequencing of An. coluzzii Ngousso mosquito RNA. SNPs were verified using the trace files.

RNA preparation and deep sequencing

Pools of 30 An. coluzzii Ngousso colony mosquito midguts were homogenized in TRIzol (Ambion) and total RNA was extracted following the supplied protocol. Small-RNA libraries were prepared from the small RNA fraction using TruSeq Small RNA Library reagents (Illumina). Libraries were sequenced on an Illumina Hiseq 2500 in a multiplexed 51 +7 bases single read using a TruSeq SR cluster kit v3 cBot HS, and a TruSeq SBS kit v3 HS 50 cycles (Illumina). Primary analysis of the sequences was performed with Casava software (v1.8 Illumina). Library preparation and sequencing was performed by ARK Genomics at the Roslin Institute (University of Edinburgh). Data are available in the EBI European Nucleotide Archive ( http://www.ebi.ac.uk/ena/) under study accession number ERP012577 and sample accession numbers: ERX1180340, ERX1180341, ERX1180342 and ERX1180343.

Long paired-end RNA libraries were prepared and sequenced by the Institut Pasteur Transcriptomics and Epigenomics core facility using standard long paired-end rapid run protocol on an Illumina Hiseq 2500. Primary analysis of the sequences was performed with Casava software (v1.8 Illumina). Long paired-end sequencing data are available in the EBI European Nucleotide Archive ( http://www.ebi.ac.uk/ena/) under accession numbers: ERX1220645 and ERX1220646.

Bioinformatic virus discovery

Bioinformatic analysis was performed with the Galaxy [ 29, 46, 47] Mississippi server instance hosted by the ARTbio facility using the Metavisitor workflows (v1.0) [ 28]. Briefly, small RNA sequencing reads were clipped of the adapter sequence, converted to fasta format and concatenated. Then reads were converted to a weighted fasta format to reduce computing load. Reads were then depleted by successive bowtie alignments on the genomes of the known organisms present in the dataset. Unaligned unique reads were then used for the subsequent contig assembly. Velvet and Oases were used to reconstruct longer contigs with k-mers ranging from 15 to 29 [ 48, 49]. The NCBI BLAST tool suite was used to search a BLAST database created from the nucleotide sequences deposited in the NCBI repository with the reconstructed virus contigs, for related homologies. To reconstruct a more complete AnCV genome, an additional dataset from long paired-end sequencing was used (ERS977505). Similarly, Metavisitor was used but with longer k-mers (ranging from 25 to 69). These analyses are accessible in the Mississippi Galaxy server using the link to a Galaxy page https://mississippi.snv.jussieu.fr/u/drosofff/p/anopheles-viruses.

IRES structure analysis

Using the structure of the IRES RF0458 family as a scaffold and the multiple alignment between the seeds of this IRES family and the potential IRES sequence of AnCV ( S2 File, we used webserver based versions of TRANSAT to plot the most probable conserved helix and secondary structures of the IRES [ 50]( Fig 6A) ( http://www.e-rna.org/transat), and R-chie ( http://www.e-rna.org/r-chie) in "single covariance" mode to plot the covariance between the nucleotides in the structure [ 51]( Fig 6B). Both programs were used with the defaults parameters and with the structure and multiple alignments given in S2 File as inputs.

RT-PCR assays for detection of virus presence

RNA from individual mosquitoes, individual organs, saliva or pools of either was extracted using either TRIzol (Ambion), NucleoSpin RNA II or NucleoSpin 96 RNA kits (Macherey-Nagel) according to the supplied protocol. RT-PCR was performed using Superscript One-Step RT-PCR kit (Invitrogen), primer ( S2 Table) annealing was performed at 56°C. For the sequencing of PCR amplicons from wild or colony mosquitoes, amplicons were purified using the QIAquick PCR Purification kit following manufacturer instructions. Sanger sequencing was performed using T7 primers by SEGENIC Eurofins, Cochin Institute, Paris.

Northern blotting

RNA migration and Northern blotting used standard procedures. Northern blot sense and antisense probes were synthesized by SP6 transcription of the PCR fragments incorporated in the PGEM-t easy vector. Multiple vector clones were verified by classical Sanger sequencing and no variation from the in silico reconstructed contigs was observed. RNA probe hybridization to the Northern blot was performed at 68°C.

Phylogenetic analysis

Basic multiple alignment of the polyhedrin segment sequences was performed using Clustal O (v1.2.1) and sequence accession numbers DQ212785.1 and AY876384.1. Basic amino acid comparison for the polyhedrin protein of AnCPV (segment 10, KU169880) was performed against ABA61432.1 and AAW80667.1 using Serial Cloner (v2.6.1, http://serialbasics.free.fr).

MUSCLE multiple alignment and evolutionary analyses were conducted in MEGA (Tamura et al., 2011) with 1000 bootstrap replicates. Because non-conserved regions could possibly influence the phylogenetic tree reconstructions, gaps were removed using the 'Complete Deletion' option in MEGA. Phylogenetic reconstructions using neighbor joining (NJ), maximum parsimony (MP) or maximum likelihood (ML) showed the same topology for each dataset. Trees constructed with partial deletion are also included ( S2 Fig). EMBL-EBI Transeq and Sixpack online tools ( http://www.ebi.ac.uk/Tools/st/) were used to translate nucleic acid sequences to the corresponding peptide sequences.

The multiple alignments for Fig 2 and S1 Fig are based on the amino acid sequences given in S3 and S4 Files.

Accession numbers for the RdRp alignment: YP_009111328.1 ( Dendrolimus punctatus cypovirus 22), NP_149147.1 ( Lymantria dispar cypovirus 1), ABB51571.1 ( Heliothis armigera cypovirus 14), AF291683_3 ( Trichoplusia ni cytoplasmic polyhedrosis virus 15), DQ192251.1_1 ( Operophtera brumata cypovirus 19), AJC97790.1 ( Thaumetopoea pityocampa cypovirus 5), AHJ14791.1 ( Inachis io cypovirus 2), ACA53380.1 ( Choristoneura occidentalis cypovirus 16), ADH10220.1 ( Antheraea mylitta cypovirus 4).

Accession numbers for the polyhedrin alignment: AAA50751.1 ( Heliothis armigera cypovirus 8), AJC97797.1 ( Thaumetopoea pityocampa cypovirus 5), NP_066381.1 ( Trichoplusia ni cypovirus 15), ABB17224.1 ( Operophtera brumata cypovirus 19), AAY96324.1 ( Heliothis assulta cypovirus 14), ACX54964.1 ( Bombyx mori cypovirus 1), ABB17220.1 ( Operophtera brumata cypovirus 18), AAW80667.1 ( Uranotaenia sapphirina cypovirus 17), YP_009002596.1 ( Inachis io cypovirus 2), ABW87644.1 ( Choristoneura occidentalis cypovirus 16), ABH85367.1 ( Simulium ubiquitum cypovirus 20), AAP44510.1 ( Antheraea proylei cypovirus 4), AIY60608.1 ( Dendrolimus punctatus cypovirus 22).

The amino acid sequences used for the multiple alignments for Fig 5 and S2 Fig are given in S4 Fig. Accession numbers for the amino acid sequences used: ALS55295 ( Anopheles C virus), NP_647481.1 (Cricket Paralysis virus), NP_044945.1 ( Drosophila C virus), AF486073_1 (Acute bee paralysis virus), YP_008888535.1 ( Formica exsecta virus 1), YP_164440.1 ( Solenopsis invicta virus 1), NP_851403.1 (Kashmir bee virus), ABY57949.1 (Israeli acute paralysis virus), NP_620555.1 ( Plautia stali intestine virus), AGF84786.1 (Aphid lethal paralysis virus), AAT81157.2 (Taura syndrome virus), NP_046155.1 ( Rhopalosiphum padi virus), YP_006666503.1 ( Macrobrachium rosenbergii Taihu virus), YP_004063985.1 (Mud crab dicistrovirus), AGW80519.1 ( Himetobi P virus), NP_620564.1 (Black queen cell virus).

Supporting Information

S2 Fig. AnCV non-structural protein phylogeny.

Neighbor-joining tree of non-structural polyprotein sequences of Dicistrovirus. Multiple protein alignment was done using the partial deletion option in MEGA. NJ bootstrap values above 0.75 are indicated above branches. The two genera of the Dicistroviridae family are indicated on the tree.
https://doi.org/10.1371/journal.pone.0153881.s002
(PDF)

Acknowledgments

We acknowledge Emilie Pondeville for aid with larvae midgut dissection, Sabine Thiberge for salivary gland dissection, Sebastian Lequime and Louis Lambrechts for Aedes aegypti eggs and cDNAs, and Ferdinand M. Nanfack for RT-PCR detection of virus.

Author Contributions

Conceived and designed the experiments: GC KE CA KDV. Performed the experiments: GC IH KE MD DD AV SK DM SS E. Belda E. Bischoff CA. Analyzed the data: GC E. Belda E. Bischoff CA JR. Wrote the paper: GC KE CA KDV.

# data[].attributes.attachment_link.title
Identification and Characterization of Two Novel RNA Viruses from Anopheles gambiae Species Complex Mosquitoes

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