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

The overall goal of this new isolation strategy is to widen the number of new virus isolates in order to study the pan-genome of this new viral family, and to better understand their ecosystems and potential pathogenicity to humans. This method can answer key questions in the virology field, to help virologists world wide interested in prospecting giant viruses and faustoviruses in particular. The main advantage of this technique is that it show new improvement to many stage like the antibiotic and antifungal mixture.

The new starvation medium adapted for the amoeba and the use of floss-cytometry for faster and more sensitive lysis detection. After collecting dirty water samples according to the text protocol, prepare the host cell, V.vermiformis by adding the amoebae to a 75 square centimeter cell culture flask with 30 milliliters of protease peptone yeast extract glucose, or PYG medium and incubate at 28 degrees Celsius for 48 hours. Use counting slides to count the amoebae and when they reach a concentration of five times 10 to the 5th, to 10 to the 6th cells per milliliter, harvest the cells by centrifugation at 720 times g for 10 minutes.

Aspirate the supernatant, and use 30 milliliters of sterile pages amoebal saline or PAS to resuspend the amoebae pellet. After rinsing the pellet a second time, use 30 milliliters of starvation medium, containing an antibiotic and an antifungal to resuspend the cells to approximately 10 to the 6th amoeba per milliliter. Starvation medium enables the amoebae to stay alive without encystment or multiplication.

Next, seed 200 microliters of amoeba in the wells of a 96 well flat bottom plate and incubate at 30 degrees Celsius for 30 minutes. Then, after vortexing the water samples for one minute at room temperature, innoculate 25 microliters of each sample onto the amoebae cells, leaving two wells without sample as negative controls. To avoid evaporation, place the plate containing the primo culture into a bag with a wet compress, and place the humidified chamber with plate into an incubator at 30 degrees Celsius.

Three days after the first innoculation, sub-culture the primo culture plate onto a fresh 96 well plate of amoebae as just demonstrated, without any microscopic observation. Incubate at 30 degrees Celsius for three days. Then, using the subculture, innoculate a third plate as just described, and incubate for two days.

To detect lysis in the final cocultured plate, turn on the cytometer and run a clean and rinse according to the manufacturer’s protocol to reduce the background noise. To carry out a 40 minute completely automated acquisition, launch the high throughput sampler or HTS to automatically load samples from the 96 well plate according to the manufacturer’s protocol. Program the HTS settings as listed here.

That include a sample volume of 150 microliters, a mixing volume of 50 microliters, a mixing speed of 180, the number of mixes as 5, a washing volume between samples of 400 microliters, a flow rate of 2.5 microliters per second, and the number of recorded events as 10, 000. Click on home to allow the HTS to spot or locate the plate. Then assess the negative control wells containing the host amoeba cells first.

In order to gate the amoeba population, and to determine the cell percentage according to their physical characteristics. Be sure to have a homogeneous amoeba population and a second population of lower events corresponding to debris and noise. Next, start the gating by clicking acquire sample.

Then, using the arrow, which defines the gates, localize the amoeba population of interest. After gating at least 10, 000 events, click on run to allow the HTS to automatically run the entire plate. Perform data acquisition and analysis according to the size parameter forward scatter and the structure parameter side scatter.

Refer to the text for additional details. Following licence detection by flow-cytometry, pipette the lysed cocultures to suspend the remaining amoebae. After pipetting 50 microliters of each sample onto slides, directly cytocentrifuge the suspension at 800 times g for 10 minutes.

Next, use pure methanol to fix the slides, then leave the slides to dry at room temperature. After fixation, plunge the slides into ESN solution three times for four seconds each before placing the slides into blue azur for six seconds. Finally, plunge the slides into PBS for 45 seconds.

Leave the slides to dry at room temperature before proceeding to observation. To stain new fixed slides with DAPI, pipette 15 microliters of DAPI into the dry slides. Observe the slides under a light microscope at 1000x magnification in order to check for the presence of virus factories.

To carry out electron microscopy, deposit five microliters of the positive sample onto a glow-discharged grid, and incubate at room temperature for approximately 20 minutes. Carefully dry the grid, then add a small amount of 1%ammonium molybdate and incubate for 10 seconds. After carefully removing the drop, allow the grid to dry for five minutes.

Next, set the electron microscope to 200 kiloelectron volts. Then, place the grid on the holder and introduce the holder into the microscope. Click on the vacuum overview icon and check the vacuum.

Then, close the valves, and set a spot size of five, and a magnification of 25, 000 before beginning observation. After locating a giant virus on the acquisition screen, use the measure tool to measure the virus. Carry out molecular biology and viral genome sequencing according to the text protocol.

This figure shows results from the detection of two faustoviruses compared to the reliable negative controls. The negative control for lysis contained an 86%amoeba population. By contrast, more than 60%of the amoebae were lysed with the highest percentage of debris in the positive samples.

As seen here, Eus and blue azur stains the nuclei of uninfected and infected amoebae cells, and the presence of virus factories inside the infected cells. DAPI staining also confirms the presence of virus factories inside amoebae. In these images, electron microscopy reveals the presence of faustovirus with the typical appearance of megavirals composed of an icosadedral capsid that lacks fibrils and a size of 200 nanometers.

Once mastered and after the coculture, this technique of lysis detection and virus characterization can be done in an hour, if it’s performed properly. While attempting this procedure, it is important to remember to control the amoebal viability and to check any probable contaminations. Following this procedure, other methods like molecular biology targeting giant viruses can be performed in order to answer additional questions like the virus family and or genotype.

After its development, this technique paved the way for researchers in the field of virology to explore this viral family, their specific host, and their potential pathogenicity. After watching this video, you should have a good understanding of how to easily isolate giant viruses by widening the number and types of samples and by choosing the best host cell and the best control conditions. Don’t forget that working with viruses potentially pathogenic for humans can be extremely hazardous, specially with huge samples.

So precautions such as working in an SB2 labs with secure conditions should always be taken while performing this procedure.