Isolation and Quantification of Zika Virus from Multiple Organs in a Mouse

In this video we describe a set of procedures designed to interrogate the impact of a viral infection and the resulting immunological response using a Murine model of Zika infection. This protocol is significant because it allows us to develop a mechanistic understanding of infection and immunity by identifying the extent to which a virus has spread throughout the body, transversing physical barriers and accessing tissues. These techniques allow a detailed understanding of viral pathogenesis and provides the ability to dissect the mechanism of protection provided by vaccines or therapeutics.

Here we demonstrate this method using Zika virus infection of mice, but this method can be applied to a range of viral infections including other flaviviruses like Dengue virus and alphaviruses like Chikungunya virus. Harvesting the organs of the central nervous system can be challenging, so it's important for a person to plan a small experiment, and be prepared to practice the techniques beforehand. This protocol allows for the rapid dissection of viral spread within small animal models with the ability to visualize, capture, and store experimental data.

It is important to visually demonstrate this method to allow the experimenter to understand how to harvest the central nervous system organs. It's also important to see the level of cell confluency and the intensity of the viral foci. To begin, use forceps to remove the pelt of the infected mouse.

Decapitate the head of the mouse with scissors, followed by the removal of the arms and legs. To harvest the brain, use serrated La Grange scissors to cut the skull through the foramen magnum. Peel off the skull with forceps, and scoop out the brain with a spatula.

Using strong blunted scissors, remove the bone surrounding the spine, then cut across the pelvic bone to expose the vertebral foramen at the lumbar level. Then, remove as much muscle as possible surrounding the spinal column. Carefully, place the beveled tip of the needle inside the vertebral foramen, avoiding excessive pressure to prevent the needle trespassing the vertebral body.

Hold strongly to exert pressure on the vertebral body and the needle, and press the syringe plunger to expel the cord into the petri dish. Immediately transfer the spinal cord in the labeled tube, and place in the dry ice bath for further homogenization. For homogenization, add one milliliter of DMEM and homogenize the sample in a BeadBeater.

After ensuring organs are homogenized, centrifuge and aliquot samples on ice and store in the minus 80 degrees Celsius until needed. On the day of the assay, remove the homogenized samples from the minus 80 degrees Celsius freezer, and allow them to thaw. With the samples on ice, dilute Zika virus tenfold into the first well of the dilution plate and use a multichannel pipette to do tenfold serial dilutions down the plate, changing tips each time.

By adding 20 microliters of sample into 180 microliters of growth media, change pipette tips between each dilution. Prepare the focus forming plate by removing the media from the prepared flat bottom 96-well plate covering virocells. To prevent the monolayer from drying out, immediately add 100 microliters of the virus dilution to each well in the viroplate.

Use the same set of tips to add from the lowest to the highest concentration. Rock plate side to side two to four times and incubate at 37 degrees Celsius in 5%carbon dioxide for one to two hours. Warm up 2%methylcellulose to room temperature.

Then, dilute the 2%methylcellulose solution in growth media at a ratio of approximately two to one. After incubation, add 125 microliters of the methylcellulose growth media to each well of the 96-well plates. Incubate again at 37 degrees Celsius in 5%carbon dioxide for 32 to 40 hours.

To fix the virocells, in a biosafety cabinet add 50 microliters of 5%paraformaldehyde over the top of the methylcellulose layer in each well of the 96-well plates. Incubate for 60 minutes at room temperature. Alternatively, cover the plates with Parafilm and place them at four degrees Celsius overnight.

After that, use a pipette to remove overlay and media off cells into a disposable container inside the biosafety cabinet. Wash gently with 150 microliters of PBS per well. Remove PBS from the plates and remove the plate from the biosafety cabinet.

Repeat the PBS wash twice, then add 150 microliters per well one times FFA wash buffer and leave at room temperature for five to 10 minutes to permeabilize the fixed cells. Next, use primary Zika antibody to detect infection. Prepare the primary antibody 4G2 at a concentration of one microgram per milliliter in FFA staining buffer.

Flick FFA wash buffer from the plates, and add 50 microliters of the primary antibody in FFA staining buffer to each well. Then seal the plates with Parafilm, or plastic film, and incubate overnight at four degrees Celsius. Prepare the secondary go anti-mouse HRP labeled antibody at a concentration of one to 5, 000 in FFA staining buffer.

After removing the antibody solution, wash cells three times with 150 microliters FFA wash buffer per well. Remove the wash buffer by flicking into the sink each time. Stain the cells with the secondary antibody in FFA staining buffer at 50 microliters per well and incubate one to two hours at room temperature.

After that, wash cells three times with FFA wash buffer, again, removing the wash buffer by flicking into the sink each time. Add 50 microliters of the TrueBlue Substrate into each well. Wait two to 15 minutes until spots are fully defined with minimal background.

After the spots are visible, wash gently with water using a hand to shield the monolayer from the force of the water running. Tap dry on paper towels. Shortly after, use a dissecting scope to count manually the spots in each well, or use an automated spot counter.

In the ImmunoCapture software, select a dilution with easily distinguished foci between 20 to 200 per well for each sample and calculate titer in focus-forming units per millimeter using the average of duplicate wells. In this experiment, viral burden was shown in the peripheral and central nervous system tissues after Ifnar mice were given Zika virus subcutaneously. The limit of detection was between 100 to 500 FFU per gram, based upon organ.

When performing the focus-forming assay, technical mistakes can result in suboptimal results. Organ toxicity, driven by high concentration of liver, altered the sensitivity of the assay. When the viral titer in the organ was lower than the toxicity, the FFA could not accurately record the viral titers.

Kidney cells illustrated toxicity with high organ concentration but was overcome by the viral titer at low organ concentrations. Vigorous pipetting or washing can remove the monolayer, leading to lost data inaccurate reporting of titer results. Fibers or hairs that are present in lab bench absorbent paper, can contaminate individual wells and cause significant errors if using an automated counting program.

Cell density is another issue, which can dramatically impact the success of a focus-forming assay. Cells at approximately 60%confluency at the start of the assay, compared to cells plated at 90%confluency, showed dramatic difference impacting the focus-forming assay. Planning the size and scope of the experiment so that organ harvest is completed efficiently, is the most important part of the procedure, followed by starting the focus-forming assay with high quality cells.

This procedure has been used to quantify virus for multiple viral families. This protocol can also be adapted to screen for therapeutic compounds and quantify neutralizing antibody titers. These techniques are used to quantitate live virus and appropriate safety steps should be taken according to the guidelines laid out in the BMBL.