A Rapid Strategy for the Isolation of New Faustoviruses from Environmental Samples Using Vermamoeba vermiformis

1. Sample Collection

1. Collect 70 samples from different environments and regions. In this case, use the following: 5 dirty water samples from the village of Saint Pierre de Meyzoargues (France), 15 samples from the lake in Parc Borély in Marseille (France); 15 sea water samples with sediment from the rocky inlets at Samena in Marseille (France), 25 river water samples from the Alps (France), and finally, 10 samples from sewage in La Ciotat (France).
2. Vortex samples for homogenization before inoculating, for one minute at room temperature without any treatment.

2. Isolation Procedure

1. Host Preparation for Blind Culture Procedures and Sample Inoculation
   1. Use V. vermiformis (strain CDC19) as a cell support for the co-culture.
   2. Maintain the amoebae at 28 °C, in a 75 cm² cell culture flask with 30 ml of protease-peptone-yeast extract-glucose medium (PYG). Please check PYG composition in Table 1.
      Note: After 48 hr, amoebae are quantified by a normal count using counting slides (e.g., Kovaslides).
   3. Harvest cells at a concentration of 5 x 10⁵ to 10⁶ cell/ml, and pellet amoebae by centrifugation at 720 x g for 10 min.
   4. Remove the supernatant by aspiration, and re-suspend the amoebae pellet in 30 ml of sterile Page's amoebal saline (PAS) Table 1. Repeat this rinsing procedure.
   5. Re-suspend amoebae in 30 ml of starvation medium with an antibiotic and antifungal mixture at a concentration of approximately 10⁶ amoeba/ml. The starvation medium is a survival medium for the amoeba, enabling it to stay alive without encystment or multiplication. Check composition in Table 1.
      Note: The antimicrobial agent mix contained 10 µg/ml vancomycin, 10 µg/ml imipenem, 20 µg/ml ciprofloxacin, 20 µg/ml doxycycline, and 20 µg/ml voriconazole. This mix served to reduce the bacterial and fungal contamination that can decrease or alter viral multiplication.
   6. Directly inoculate 25 µl of each sample onto the 200 µl amoeba monolayer in a 96-well plate flat bottom favoring the amoeba adherence and incubate at 30 °C in a humid environment (a humid environment consists of placing a wet compress inside the bag containing the plate in order to avoid evaporation). This is the primo-culture. Leave two wells of amoebae without any sample inoculation; this acts as the negative control. Incubate this primo-culture for three days.
   7. Three days after the first inoculation, sub-culture the primo-culture plate on fresh amoebae as described above without any microscopic observation. Incubate the new plate for three days under the same conditions as the primo-culture.
   8. Perform the final culture after these three days of incubation the same way as for the primo- and sub-culture. Incubate the final culture for two days. Then proceed to detection.
2. Detection of Lysis by Automated Flow Cytometry
   Note: Blind culture coupled with flow cytometry differs from the previous standard system of isolation. It is more sensitive, precise and automated. For detection purposes, we applied a new technique developed to detect lysis at the highest speed and without microscopic observation. This technique is advantageous for screening samples and has better sensitivity than older techniques.^7-9
   1. Use the 96-well plate of the final co-culture to detect lysis.
   2. Turn on the cytometer and run a clean and rinse cycle in order to obtain lower background noise according to manufacturer's protocol.
   3. Launch the High Throughput Sampler (HTS) combined with flow cytometer to automatically load samples from the 96-well plate and perform a completely automated acquisition within 40 min according to manufacturer's protocol.
   4. Set up the HTS as follows: sample volume 150 µl, mixing volume 50 µl, mixing speed 180, number of mixes 5, washing volume between each sample 400 µl, flow rate 2.5 µl/sec, number of recorded events 10,000.
   5. Assess the negative control wells of the final co-culture micro-plate first in order to gate the amoeba population and to determine the percentage of these cell hosts according to their physical characteristics. Be sure to have a homogeneous amoeba population and a second population of a lower events corresponding to debris and noise.
      Note: This gating procedure distinguishes subcellular debris and clumps from single cells by size, estimated by forward scatter. It is therefore important to accurately determine the percentage of each population using the best possible gating procedure. These percentages may vary, especially if many types of protozoa are used, so it is important always to have a negative control, which can be used as the reference population for the rest of the samples.
   6. Start the gating by clicking acquire sample.
   7. Record the number of events not less than 10,000 events.
   8. Localize the amoeba population of interest using the arrow. This is how to define the gates.
   9. After gating, let the HTS spot or locate the plate by clicking 'Home', then run the entire plate automatically. Perform data acquisition and analysis according to the size and structure parameters (respectively, FSC (forward scatter) and SSC (side scatter)).
      Note: Detecting the loss of the amoeba-gated population signals the presence of a pathogenic agent responsible for amoeba lysis. Protozoa lysis associated with virus infection is detected with an arbitrarily designed threshold of 50% lysis of the amoeba population gated by the analyzer and compared to the same non-infected population used as a reliable negative control.

3. Characterization of the New Isolates after Lysis Detection

1. Preliminary Staining to Reveal the Presence of Giant Viruses
   1. After lysis detection by flow cytometry, pipette the lysed co-cultures to suspend the remaining amoebae. Then directly cytocentrifuge 50 µl of the suspension at 800 x g for 10 min.
   2. After centrifugation, fix the slides with a pure methanol solution. Stain the slides using a commercial rapid staining of Eosin/blue Azur and DAPI staining and observe under a light microscope at 1,000X magnification in order to check for the presence of virus factories.
      1. For rapid staining, plunge the slides into the first eosin solution (4 sec repeated three times) then pass directly to plunge slides into the second fixing solution containing blue Azur for 6 sec, and finally rinse the slides for 45 sec with a phosphate buffer solution pH 7.2. Leave the slides to dry at room temperature, then proceed to observation.
      2. For DAPI staining, deposit 15 µl of the DAPI commercial stock solution onto the fixed dry slide. Protect from light then proceed to observation.
2. Electron Microscopy
   1. After the first identification on the slides, assess the presence of the giant virus through electron microscopy observation of supernatant cultures.
   2. Deposit 5 µl of the positive sample onto the glow-discharged grid. Wait for approximately 20 min at room temperature.
   3. Dry the grid carefully and deposit on it a small drop of 1% ammonium molybdate for 10 sec. Leave the grid to dry for 5 min.
   4. Proceed to electron microscopy at 200 keV.
   5. Place the grid on the holder; introduce the holder into the microscope. Check the vacuum overview by clicking on the corresponding icon. Close valves and begin observation at a spot size 5, and magnification of x 25,000.
   6. Measure the giant virus using ImageJ or directly on the microscope using the measure tool of the microscope located on the acquisition screen.
      Note: Classify the sample into the Faustovirus group with a capsid size of approximately 200 nm.
3. Biologie moléculaire
   Note: Following this identification, molecular biology tests were crucial for the confirmation in order to distinguish Faustovirus from Marseillevirus, which has the same capsid size and appearance under electron microscopy, although they do not have the same host specificity. The Marseillevirus cannot grow on Vermamoeba, the Faustovirus is specific to Vermamoeba and all attempts to grow it on other amoeba such as Acanthamoeba have failed to date. The design and application of the specific primer/probe systems listed in the works of Reteno et al. enabled a more precise preliminary classification of the newly isolated viruses through amplification and sequencing of the viral genes.
   1. Extract DNA from the positive culture samples using an automated extraction system according to the manufacturer's protocol.
   2. Perform Real-time PCR using a thermocycler as described in Reteno et al.¹⁹
   3. Sequence using the same primers that were used for amplification. Use the primers targeting DNA-directed RNA polymerase subunit 1 gene: Fstv_S2F 5'-CCA GGA CAT GAT GGT CAC ATA G-3' (forward) and Fstv_S2R 5'-TTG CAC CTC CGC AGT TAA A-3' (reverse) with the Fstv_S2P FAM-TATGCTCCAATGGCCTTCAACGACA-TAMRA probe.

4. Virus Production, Purification and Genome Sequencing

1. Subculture the single virus amoeba culture for end-point dilution cloning.
2. For the production: prepare 20 flasks containing 30 ml of PYG, 10 ml of Vermamoeba vermiformis and 5 ml of the isolated virus already transferred from wells to small flasks.
3. Incubate at 30 °C until complete lysis of the amoeba. Observe every day by inverted optical microscopy until all amoeba are lysed.
4. After complete lysis, pool all flasks into one recipient. Filter with a 0.45 µm filter to eliminate debris.
5. Start the purification by ultracentrifugation at 89,800 x g for 75 min. Make sure to calibrate the weight of the tubes before starting centrifugation.
6. After centrifugation, remove the supernatant from each tube by aspiration and re-suspend the pellet in PAS with the same volume used for calibration, before repeating centrifugation.
7. As above, remove the supernatant and re-suspend all the pellets in 1 ml of phosphate buffered saline (PBS).
8. Purify the virus that is produced, using 25% sucrose (27.5 g of sucrose in 100 ml of PBS, filtered on a 0.22 µm filter).
   1. Use 8 ml of sucrose and 2 ml of the viral suspension. Centrifuge at 89,800 x g for 1 hr 15 min. Re-suspend the pellet in 1 ml of PBS. Store at -80 °C.