All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Organ Isolation

NOTE: The virus is not stable at room temperature (RT) so the number of animals harvested at one time must be planned carefully to preserve viral titers.

1. Infect mice using the chosen dose and route, based on the phenotype required. For this protocol, infect 8-10 week-old male and female type I interferon receptor-deficient (Ifnar1^-/-) C57BL/6 mice.
   1. Anesthetize Ifnar1^-/- mice with a cocktail of Ketamine (90 mg/kg) and Xylazine (10 mg/kg). Test pedal reflex by a firm toe pinch to confirm anesthesia. Administer 1 x 10⁵ focus forming units (FFU) of ZIKV in 50 μL subcutaneously (SC) via the footpad.
2. Prepare all the materials needed for harvest the day before 2 scissors, forceps, and 20 mL of 70% ethanol (EtOH) in a 50 mL conical for disinfection of tools.
   1. Prepare syringes and needles: a 20 mL syringe for perfusion, 23 G needles (approximately 1 per cage), and 1 mL syringe with 25 G needles (1 per 3-5 mice). Prior to weighing tubes, add steel beads (in the hood) to 1.5 mL O-ring screw cap tubes (one for each organ). The tubes need to be appropriate for homogenization.
3. Prepare the day of the harvest the materials needed for the perfusion and organ freezing.
   1. Fill an ice bucket with dry ice and 70% EtOH to make an ice bath. Prepare sterile phosphate buffered saline (PBS) assuming that 25-30 mL per mouse of PBS for each perfusion is needed 5-10 mL for the spinal cord). Obtain anesthesia, a cocktail of Ketamine and Xylazine, and assume 300 μL per mouse. Place all the materials and any additional tubes required for flow cytometry, etc., in a secondary container to transport to the animal facility.
4. Follow all necessary procedures for entry including donning and doffing personal protection equipment during entry and exit into the animal facility.
   CAUTION: All the procedures are to be carried out in a certified biosafety cabinet within a Biosafety level 2 laboratory (BSL-2) facility.
5. Administer anesthesia, 200-500 μL, intraperitoneally depending on the size and weight of the animal. If working alone, administer anesthesia to one animal at a time to avoid killing the animals before organ harvest.
   1. Confirm anesthesia dose by pinching the toe with forceps to evaluate the pedal reflex. Do not continue unless completely nonresponsive. Use pins to secure the mouse to a wax board. At this point, douse the mouse in 70% ethanol to avoid hair contamination in the organ harvest procedure.
6. Terminally bleed by heart puncture.
   1. Using the scissors and forceps, open the animal through the chest cavity to expose the heart. Collect blood via heart puncture (~800 µL), this can be used for multiple assays including clinical chemistry, hematology, flow cytometry to detect antigen-specific responses, and real-time quantitative PCR (qRT-PCR) for the analysis of viral copy number.
   2. For viral RNA analysis by qRT-PCR, collect blood in an EDTA tube if extracting from whole blood or in a microcentrifuge tube if extracting from serum. (For serum, spin the tube at maximum speed in a centrifuge, room temp, for 20 min, and remove serum to a separate tube). Add a linear polyacrylamide as a carrier to the serum sample after finishing. RNA can then be analyzed or stored at -80 °C to further process at a later date.
7. Fill the 20 mL syringe with PBS, at room temperature (or 37 °C), and perfuse mice by inserting the butterfly needle into the left ventricle. Puncture the right atrium to allow blood and PBS to exit. Slowly administer the PBS while checking the color of the liver to confirm that the animal is completely perfused. The liver should change from deep red to pink salmon color.
   1. If preparing the organs for histology, leave the butterfly needle in place and then perfuse with 20 mL of ice-cold 4% paraformaldehyde. If so, it is convenient to have the syringe connected to a 3-way stopcock with both syringes attached to it, turning the valve ON and OFF as one alternates between PBS and PFA.
8. Harvest organs into labeled, weighed tubes. For peripheral organs, follow an established order for harvest: liver, spleen, kidney, and lungs.
   1. Take only one lobe from the liver; it does not matter which one, but for all experiments always try to take the same lobe with the same size piece. Similarly, for kidneys and lungs, take the same kidney and lung. If flow cytometry is also to be completed, any organ can be cut in half. Store the half to be used for cytometry in Roswell Park Memorial Institute medium (RPMI) at room temperature until the harvest is complete.
   2. Immediately after harvest, put each organ in a labeled tube and place it in the dry ice bath. Virus titer reduces over time at room temperature, so the amount of time it takes to harvest organs is extremely important. There must be consistency between mice, so after safety, the next priority is speed.
      NOTE: If harvesting organs for viral titers, it is very helpful to have a second individual harvest the brain at this point, to preserve viral titers.
9. Remove the remaining organs from the mouse body cavity to gain access to the spine and skull.
   1. Remove the mouse from the board and remove the pelt, followed by the removal of the arms and legs.  
      Remove the head of the mouse with decapitation, blunt, or surgical scissors to harvest the brain by cutting the skull with serrated LaGrange scissors through the foramen magnum. Then, peel off the skull with forceps and scoop out the brain with a spatula.
   2. Using strong, blunted scissors, remove the ribs and other bones surrounding the spine. Then, cut across the pelvic bone, exposing the vertebral foramen at the lumbar level. The small tip of the spinal cord should be visible at this point.
   3. Use a 10 mL syringe filled with PBS and an 18 G needle to expel the spinal cord by "flushing" the cord from the lumbar to the cervical spine over a Petri dish.
   4. Carefully, place the beveled tip of the needle inside the vertebral foramen, avoiding excessive pressure to prevent the needle from 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. Immediately transfer the spinal cord to the labeled tube and place it in the dry ice bath.
10. Repeat the procedure with all the animals until the harvest is completed. Place the harvest tools in the 70% ethanol between animals. Focus on consistency and speed. The length of time between each organ harvest should remain consistent so as to not bias viral titer results.
11. When finished, disinfect the biosafety cabinet and all material prior to removing it from the animal facility.
12. Remove each tube from the ice bath and weigh the tubes to determine organ weight. To determine the organs' weight, subtract the weight of the empty tube from the weight of the organ-containing tube.      
    NOTE: At this point, organs can be frozen at -80 °C to further process at a later date or organs can be homogenized immediately before freezing individual aliquots. Freeze-thawing samples multiple times decreases viral titer. Therefore, it is important to do the same procedure for all experiments within a single project.

2. Organ Homogenization

1. Prepare 3 labeled, 1.5 mL snap-capped tubes for each organ to be homogenized.
2. If samples are not being homogenized immediately after harvest, remove tubes from -80 °C. The samples do not need to be thawed to homogenize.
3. Put the samples on ice to keep them cold to minimize viral titer loss. Then, add 1 mL of cold DMEM containing 5% FBS to each organ-containing tube.
4. Immediately beat tubes in a bead beater according to the manufacturer's instructions. Homogenize all organs with steel beads in a bead-homogenizer instrument. Check to ensure each organ has been completely homogenized.
5. Spin down organ debris in a microfuge at 12,000 x g for 5 min in a microfuge that has been chilled to 8 °C. Then return the tubes to the ice bucket. In the biosafety cabinet, aliquot samples into tubes for necessary assays. Then return the tubes to the ice bucket.
6. For focus forming assay, aliquot 500 µL into a labeled tube. Then place the tubes in the rack on an ice bucket. Aliquot 50 µL into a tube for RNA for fluorogenic quantitative RT-PCR (qRT-PCR) to measure viral genome copy number.
7. Isolate total RNA from the organs of the infected animals using a commercial RNA isolation kit. Determine flavivirus viral RNA using the primer-probe sets specific for ZIKV, which recognizes unique sequences in each flavivirus genome. Determine viral copy number using a copy control plasmid containing a defined positive single-stranded RNA generated in vitro using T7 polymerase containing the ZIKV target sequences.
8. Aliquot the remaining sample, which is approximately 300 µL, into the third tube and store it at -80 °C if needed.  
   NOTE: If samples were not previously frozen, freeze at -80 °C. If samples had been previously frozen, continue onto the focus forming assay and/or RNA isolation before stopping.

3. Zika Virus Focus Forming Assay

NOTE: It is important to include a no-virus control and a positive control. The positive control is a dilution series of a virus stock with a known concentration. Not all controls need to be on the same plate, but as the assay becomes larger than 5 plates, more controls should be added and spread out among plates. Take care not to scratch the monolayer with either the pipet tips or by vigorous washing. Multiple organs can be titered on the same day or different days. However, an individual organ should not be titered over multiple days because different assay conditions can impact viral titer. It is strongly recommended to run an individual organ on a single day.

1. Prior to the day of the assay, prepare the cells and reagents needed for the focus-forming assay.
   1. Prepare growth media containing 500 mL of DMEM with 5 mL of HEPES and 25 mL of FBS. Have Vero-World Health Organization (WHO) cells growing in growth media at 37 °C, 5% CO₂ prior to the start of the assay. The Vero cells should not be a high passage or ever grown over 100% confluency prior to the start of the assay.
   2. Prepare 500 mL of a 2% methylcellulose solution by autoclaving a 1 L glass media bottle with 10 g of methylcellulose, and a large stir bar, and a separate 1 L glass media bottle with 500 mL of H₂O. If the autoclaved water has cooled reheat in the microwave until the bottle is hot to the touch, but not boiling.
   3. Gently pour warm/hot water into a bottle of methylcellulose while in the tissue culture hood. Partially cap the bottle and stir on the hotplate until methylcellulose is in solution (1-4 h). Aliquot the 2% methylcellulose solution into a sterile 50 mL conical tube. 2% Methylcellulose can be stored at 4 °C until needed.
   4. Prepare a 5% Paraformaldehyde solution in PBS for fixing the plates and stored it at 4 °C until needed. Prepare a 1x focus forming assay wash buffer by adding 0.05% Triton X-100 to PBS and stored at RT. Prepare a 1x FFA staining buffer by adding 1 mg/mL saponin to PBS and stored at 4 °C until needed. These can be prepared one- to two weeks in advance.
2. Calculate the number of flat-bottom 96 well plates needed for the assay such that each organ is plated in triplicate. Include enough additional wells for positive and negative controls.
   1. Grow up enough Vero cells in growth media to complete the assay designed. Trypsinize the Vero cells and count them resuspending them at 1.5 x 10⁵ cells per mL in growth media. Plate Vero cells in the 96 well flat-bottom plates at 3.0 x 10⁴ Vero-WHO cells/well in growth media by adding 200 µL per well.
   2. Incubate plates at 37°C at 5% CO₂ overnight, make sure the plates are level in the incubator so the cells are equally distributed within the well.
      NOTE: Each laboratory grows Vero cells slightly differently; the target is a 90-95% confluent monolayer in each well on the day of the assay for Zika virus.
3. Dilute Zika virus samples on the day of the assay. If the organ samples had been previously homogenized, remove samples from the -80 °C freezer and allow them to thaw before placing the samples on ice for the assay.
   1. On the ice, prepare a round bottom 96 well plate by adding 180 µL of cold growth media to rows B through H leaving row A empty. Add 150 µL of each homogenized organ sample to row A of the round bottom plate.
   2. Prepare serial 10-fold dilutions of each sample, using a multi-channel pipette. Dilute samples in a round bottom 96 well plate by adding 20 µL of sample into 180 µL of growth media, changing pipette tips between each dilution.
4. Prepare the focus forming plate by removing the media from the flat-bottom 96 well plate covering Vero cells. Do this immediately before adding the virus samples to prevent the monolayer from drying out.
   1. Add 100 µL of the virus dilution to each well in the Vero plate. Add a sample using the same set of tips by going from the lowest to the highest concentration. Rock plates side-to-side 2-4 times being careful not to swirl.
   2. Incubate at 37 °C, 5% CO₂ for 1-2 h. Make sure the plates are level within the incubator. During the incubation warm up 2% methylcellulose to RT.
   3. Dilute 2% methylcellulose solution in growth media. The dilution should be at a ratio of approximately 2:1 of 2% methylcellulose to growth media. Keep it at room temp until it is time to use it. Add 1-2 drops/well (set the pipet to 125 µL) of the methylcellulose: growth media to each well of the 96-well plate.
   4. Incubate 32-40 h at 37 °C, 5% CO₂. As everyone's Vero cells grow slightly differently, and factors including cell confluency and strain of the Zika virus can change the incubation time.
5. Fix the Vero cells using the prepared 5% paraformaldehyde solution.
   1. Add 50 µL of 5% paraformaldehyde to each well over the top of the methylcellulose layer in the biosafety cabinet. Incubate for 60 min at RT. The fixing can go overnight at 4 °C but cover the plate with parafilm to reduce evaporation.
   2. Dump overlay and media off cells into a disposal container inside the biosafety cabinet. Wash gently with PBS, 150 µL/well. Remove PBS from the plates and remove the plate from the BSL-2.
   3. Repeat the PBS wash 2x adding 150 µL/well. Then remove the PBS. Add 150 µL/well 1x FFA wash buffer and let sit for 5-10 min at RT to permeabilize the fixed cells.
6. Detect infection using primary Zika antibody. Prepare the primary antibody 4G2 (D1-4G2-4-15) at a concentration of 1 μg/mL in FFA staining buffer. Prepare enough antibodies for the whole assay.
   NOTE: Stay away from lab diapers or other high-lint absorbent material as the fibers will negatively impact the imaging of the foci.
   1. Remove the FFA wash buffer from the plates. Add 50 µL/well of the primary antibody in the FFA staining buffer. Seal the plates with parafilm and incubate overnight at 4 °C on a rocking platform. The assay can be done with an incubation for 2 h at RT.
7. Visualize the infection with the addition of a secondary HRP-conjugated antibody. Prepare the secondary Goat anti-mouse HRP-labeled antibody at a concentration of 1:5,000 in the FFA staining buffer. Prepare enough antibodies for the whole assay.
   1. Wash cells 3x with FFA wash buffer, removing the wash buffer by flicking it into the sink each time. Stain the cells with the secondary antibody in FFA staining buffer at 50 µL/well. Incubate 1-2 h at RT.
   2. Wash cells 3x with FFA wash buffer, removing the wash buffer by flicking it into the sink each time. Add 50 µL/well of the Trueblue Substrate.
   3. Watch the plates carefully, waiting for 2-15 min until spots are fully defined and 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 (NOT DIAPERS) and image as soon as possible.
   4. Spots may be counted manually or using an automated spot counter. If counted manually, a dissecting scope can be used to aid in visualization.
   5. For each sample, select a dilution with easily distinguished foci (e.g., 20 to 200 per well) and calculate titer in focus-forming units per mL (FFU/ml), using the average of duplicate wells: FFU/mL = (mean foci/well) × (dilution factor) ÷ (mL inoculum).