Enumeration of virus particles in aquatic or sediment samples by epifluorescence microscopy
Prepare reagents.
Collect and prepare sample according to sample type.
Prepare slide labels with critical information such as the date, sample location, and volume filtered to keep track of the samples once they have been filtered and stained.
For every set of four samples to be stained, use a permanent pen to mark the bottom of a plastic Petri plate into four labeled sections.
For each filter that is to be prepared, add a 78- µL drop of 0.02-µm filtered dH2O on each section of the marked Petri plates.
Thaw a 40-µL vial of the stock SYBR solution and add 2 µL stock solution to each drop (78 µL) of sterile dH2O or buffer.
Mix the stain by gently pipetting up and down.
Place the Petri plates in the dark so that the stain is not bleached.
Prepare the antifade solution in a clean, sterilized 2-mL microcentrifuge tube.
Dilute the 10% (wt/vol) stock of p-phenylenediamine 1:100 using glycerol/PBS as the diluent.
Keep the solution on ice and protect it from light.
For each filter that is to be prepared, add a 80-µL drop of thawed Yo-Pro working solution on each section of the marked Petri plates.
Place a 9-cm-diameter filter paper soaked with 3 mL aqueous NaCl solution (0.3% wt/vol) in the lid of the Petri plates to prevent evaporation of the stain.
Connect a filtration unit for 25-mm filters to a vacuum source, ensuring the vacuum is ≤13 kPa.
Place a 0.45-µm nitrocellulose backing filter on each filter support, and overlay it with a thin layer of dH2O.
Carefully pick up a 0.02-µm Anodisc filter by its plastic ring and lay it over the wet backing filter, with the plastic ring facing upward.
If the sample has been preserved and frozen as described, thaw it in a 37°C water bath.
For a sample that has just been collected, fix it with 0.5% glutaraldehyde for 15–30 min at 4°C before preparing slides.
In addition, prepare duplicate control samples by fixing 1 mL of the 0.02-µm filtered water that was used to dilute the SYBR stain.
Because divalent cations interfere with the binding of the stain, seawater samples should be diluted to <7 psu with 0.02-µm filtered dH2O before filtration.
It is a good idea to make test slides (including a control with no sample added) to be sure an appropriate volume is filtered, that the procedure is working, and that the filters and reagents do not have viruses on or in them (some batches of Anodiscs have been covered with bacteria and viruses).
For most lake and coastal seawater samples, which have viral abundances of ~107 mL–1, 0.8–1.0 mL sample is added to the surface of the Anodisc filter while the vacuum is off.
Turn on the vacuum and suck the sample through the filter.
For oligotrophic or very deep ocean samples, it may be necessary to filter 4 mL or more.
Filter towers must be cleaned between samples.
Rinse the towers with 0.02-µm filtered dH2O followed by ethanol.
Dry with lint-free paper (e.g., Kimwipe).
Once the sample is filtered, remove the Anodisc with the vacuum still on.
Allow the filter to air-dry (typically a minute or less), until the surface is visibly dry.
Place the Anodisc, sample side up, on a drop of stain in the Petri dish.
Allow the filter to stain for 15 min in the dark.
Allow the filter to stain for 48 h in the dark, at room temperature.
Add a drop of dH2O on the backing filter, lay the stained Anodisc on top, and use the vacuum to remove any remaining fluid.
Some samples (e.g., sediments, vent fluid, and humic waters) may require the filters to be rinsed to reduce background fluorescence.
If so, while the vacuum is still on and the filter is damp, rinse the filter twice with 1 mL of 0.02-µm filtered dH2O.
Remove the Anodisc while the vacuum is on.
Place the Anodisc, sample-side up, on a 9-cm filter paper or Kimwipe in the dark, and allow the filter to dry until it appears opaque.
Place 12–15 µL antifade solution on a labeled glass slide and lay the dry Anodisc on top.
Add ~20 µL antifade on top of the Anodisc and cover with a coverslip.
Place 12–15 µL spectrophotometric-grade glycerol on a labeled glass slide and lay the dry Anodisc on top.
Add ~20 µL glycerol on top of the Anodisc and cover with a coverslip.
Remove any air bubbles that are trapped under the coverslip by gently pressing on the surface.
The slides can be counted immediately or stored frozen at –20°C for at least 4 months with no decrease in estimates of viral abundance.
Count the viruses at 1000× magnification using a 100× oilemersion objective.
Begin by checking the test filters to ensure that the reagents or filters were not contaminated and the filtered volumes were appropriate.
Check each slide before counting to make sure that the filter is evenly stained and that the viruses are on a single plane of focus and not suspended in the mounting medium and are evenly distributed across the filter.
Using the ocular reticule, select an appropriate number of grid squares so that there are 10–100 stained viruses in each field.
Estimate the abundance of viruses by counting at least 20 random fields.
Keep a tally of the number of particles in each field so that the variation in abundance of particles among fields can be determined.
For each sample, record the number of particles counted in each field, the number of fields counted, the area of the field, and the volume of sample filtered.
The abundance of viruses mL–1 (Vt) in the sample = Vc ÷ Fc × At ÷ Af ÷ S, where Vc = total number of viruses counted, Fc = total number of fields counted, At = surface area of the filter (µm2) (see note below), Af = area of each field (µm2), and S = volume of sample filtered (mL).
The total number of particles counted will determine the size of the 95% confidence intervals on the estimates of viral abundance.
By assuming a Poisson distribution, the 95% confidence intervals can be estimated using the following equations (Suttle 1993):
