Zika Virus Infection of Cultured Human Fetal Brain Neural Stem Cells for Immunocytochemical Analysis

Human neural stem cells were originally derived from discarded human fetal cortexes in the first trimester¹². All protocol procedures adhere to the University of Texas Medical Branch ethics guidelines concerning the use of human tissue samples, and the cell lines were approved by the Institutional Biosafety Committee.

1. Stock medium preparation and stem cell recovery

1. Prepare culture medium stock (DFHGPS) by combining the reagents in steps 1.1.1.-1.1.5.
   1. Add 210 mL of Dulbecco's Modified Eagle Medium with high glucose and L-glutamine (D). Store it at 4 ˚C.
   2. Add 70 mL of Ham's F12 Nutrient mix with L-glutamine supplement (F). Store it at 4 ˚C.
   3. Add 4.2 mL of 15 mM HEPES Buffer (H) and store it at room temperature.
   4. Add 4.2 mL of 10% D-glucose Solution (G) and store it at room temperature.
   5. Add 2.88 mL of penicillin/streptomycin (PS) solution.. Aliquot the stock solution into 3 mL aliquots and store the aliquots at -20 °C until needed.
      NOTE: The final concentration of penicillin in the medium will be 100 units/mL, and the concentration of streptomycin will be 100 µg/mL. This medium will be referred to as DFHGPS for the remainder of the protocol and should be stored at 4 °C for use within 1-2 weeks. If needed, DFHGPS may be scaled up proportionally.
2. The day before cells are recovered, coat a T75 flask with 5 mL of conditioned medium, obtained from previous cultures of the hNSC cell line, and leave in a 37 °C incubator with 8.5% carbon dioxide (CO₂) overnight.
   NOTE: If currently culturing hNSCs, save the medium that cells have been growing in during a medium change. This is conditioned medium as it has been "conditioned" by the cells. Conditioned medium can be saved in a screw cap tube and stored at 4 °C for approximately 7 days. Contact the corresponding author to receive hNSCs. Conditioned medium is preferred but not absolutely required, particularly when first starting the culture of hNSCs.
3. The day of recovery, warm 20 mL of DFHGPS in a 50 mL screw cap tube in a 37 °C water bath for at least 10 min and keep it in the water bath until ready to use.
4. In a separate 15 mL screw cap tube, warm 10 mL of DFHGPS in a 37 °C water bath for at least 10 min and keep it in the water bath until ready to use in step 1.13.
5. Retrieve a container of ice and all medium components (Table 1). Thaw all medium components on the ice and leave them on the ice while preparing for the medium change.
6. Obtain 5 mL of conditioned medium stock (stored at 4 °C) and keep it in a 37 °C water bath until ready to use in step 1.14. See note after step 1.2 if there is no conditioned medium or current human fetal neural stem cell culture.
7. Retrieve the 50 mL screw cap tube from step 1.3 and place it in the biosafety cabinet (BSC). Transfer 10 mL of DFHGPS into a new 15 mL screw cap tube, and then place the 50 mL tube containing the remaining 10 mL of DFHGPS medium back in the 37 °C water bath.
8. Obtain a cryo-vial of hNSCs stored in liquid nitrogen. See note after step 1.2 if there is no current human fetal neural stem cell culture.
   NOTE: Human neural stem cells were originally derived from discarded human fetal brains¹² and maintained in culture without genetic modifications¹⁰. 5 × 10⁶ cells should be thawed and plated on a T75 flask. Each cryo-vial should contain 5 × 10⁶ cells; therefore, one vial per flask is needed.
9. Thaw a vial of cells in 37 °C water bath by inverting the vial every 10 s for approximately 1 min or until ice is melted and the contents of the vial are completely liquid. Do not submerge the whole vial under water to avoid potential contamination.
10. In the BSC, use a P1000 micro-pipette to aspirate the thawed cell solution and add it drop-wise to the 15 mL tube from step 1.7, containing 10 mL of DFHGPS. To add the thawed cell solution drop-wise, slowly press down on the plunger so that only a drop or two is released at once. Meanwhile, swirl the 15 mL tube to allow a quick mixture.
11. Centrifuge the cell suspension at 200 x g for 5 min at room temperature. Discard the supernatant, making sure to retain the cell pellet.
12. During step 1.11, retrieve the 50 mL tube containing the remaining 10 mL from step 1.7. After the centrifugation, resuspend the cell pellet in the 10 mL DFHGPS and centrifuge again at 200 x g for 5 min at room temperature.
13. During the final spin, prepare the new growth medium by following Table 1 to add growth factors to the 10 mL of DFHGPS medium from step 1.4 (Table 1) in BSC. Leave the new medium containing growth factors in the BSC until ready to use.
14. Retrieve the T75 flask from step 1.2 and discard the conditioned medium coating the flask by aspiration. Then add the 5 mL of conditioned media from step 1.6. to the flask using a serological pipette. Be careful not to scratch the bottom of the flask.
15. Retrieve the tube of cells from the centrifuge and discard the supernatant by aspirating it, leaving the cell pellet intact.
16. Resuspend cell pellet in 10 mL of new media from step 1.13 by using a 5 mL serological pipette and pipetting up and down several times. Add the suspension to the coated T75 flask from step 1.14. Use the remaining 5 mL to rinse the tube that contained the cell pellet, and then add those 5 mL to the flask as well.
17. Rock the T75 flask back and forth 3 times to ensure the cells are evenly distributed across the flask. Make sure the cap of the flask is loose to allow air flow. Under a light microscope observe the cells, make notes, and take images if possible.
    NOTE: The cells should look translucent and spherical. Make a note if there are cells that look dark, opaque, or have asymmetrical boarders as this can indicate cell death. Also watch for any fungal or bacterial contamination, such as the media becoming yellow in color.
18. Place the flask of cells in a 37 °C incubator with 8.5% CO₂.
    NOTE: Use 8.5% instead of 5.0% CO₂ to maintain pH of this serum-free culture media in the range of 7.1-7.4.

2. hNSC Medium Change

NOTE: Change medium every 3-4 days .

1. Turn on the BSC and clean thoroughly with 70% ethanol. Observe cells under a light microscope.
   NOTE: Make notes on the appearance of the cells and sphere sizes. Three days after recovery or passage, cells will cluster together and begin to form non-adherent neurospheres.
   Caution: If cells adhere to the bottom of the flask, the culture will need to be discarded as cell adherence will increase premature differentiation and cells will be unable to maintain a stem state. If there are too many cells in the flask, the cells may form large spheres (greater than 2 mm in diameter) in which case, cells in the center of the sphere will begin to differentiate due to lack of access to growth factors in the medium. If spheres larger than 1 mm in diameter are observed, cells can be passaged (steps 3.1-3.22).
2. Follow steps 1.4 and 1.5, and prepare 10 mL of fresh medium (see step 1.13).
3. Tilt flask to the side, allowing spheres to sink to the bottom of the flask.
4. Remove 5 mL of conditioned media from the top of the pooled medium using a 5 mL serological pipette, taking care not to aspirate any neurospheres. Place the 5 mL of conditioned media into a clean 15 mL tube.
5. Remove an additional 5 mL of conditioned media from the flask, again carefully avoiding the spheres, and place the 5 mL into the same 15mL tube.
6. Add 10 mL of new media from step 2.2 to the remaining 5 mL of conditioned media and cells in the flask. Set the flask down flat and observe cells under a light microscope.
7. If it appears cells have been aspirated during collection of conditioned medium, spin the conditioned medium at 100 x g for 5 min at room temperature. Re-suspend any cells that accumulate at the bottom in 1 mL of conditioned medium, and then add them to the culture flask.
8. Store the 10 mL of unused conditioned media from steps 2.5/2.6 at 4 ˚C in a 15 mL screw cap tube.
9. Repeat steps 1.17/1.18.

3. hNSC Passage

NOTE: hNSCs should be passage every 9-11 days if cells grow normally, as population doubling time is approximately 3 days¹³.

1. If expanding the NSCs into a new flask, make sure all new flasks are coated as in step 1.2.
2. Clean the BSC as in step 2.1, and conduct steps 1.4 and 1.5.
3. Prepare medium as in step 1.13. Each T75 flask requires 10 mL of medium. For multiple T75 flasks, multiply 10 mL of media by number of flasks.
4. Prepare either 1 or 3 mL of Dulbecco's phosphate-buffered saline (dPBS) and glucose and place in a 37 °C water bath to warm for 10 min (for trypsin solution) according to the volumes in Table 2. Do not add trypsin or DNase at this time. DNase may be required to break down any DNA released from the dead cells due to mechanical or chemical cell dissociation.
   NOTE: For one T75 flask passaged after 9-10 days of growth there will be approximately 30 × 10⁶ cells, if 5 × 10⁶ were originally seeded on the flask. Typically, 1 mL of dPBS solution is used for 10 × 10⁶ cells. Therefore, 3 mL of dPBS solution is needed for a T75 flask passaged after 9-10 days.
5. Place clean small weigh boat and hemocytometer in hood. Clean with 70% ethanol and allow to air dry in hood for approximately 5 min.
   NOTE: The weigh boat will be used in steps 3.17.1.
6. Retrieve the flask of cells that will be passaged from the incubator and bring into the BSC. Tilt flask gently allowing cells to sink to the bottom of the pooled media. There is about 15 mL of media in the flask.
7. Remove about 10 mL of media in 5 mL portions (i.e. 5 mL + 5 mL). Place this 10 mL of media into a clean 15 mL tube labeled "CM" for "conditioned medium". Be careful not to aspirate any of the cells settled in the remaining 5 mL of media in the flask; these cells and 5 mL of media will be subjected to step 3.8.
   1. Take 3 mL of conditioned media from the "CM" tube (from step 3.7) and place into a 15 mL tube labeled "trypsin inhibitor solution". This will be used in step 3.12. After removal of the 3 mL, there is 7 mL of conditioned media left in the "CM" tube. Set the "CM" tube aside until step 3.9.
8. Remove the 5 mL of media and cells remaining in the T75 flask, and place in a clean 15 mL tube labeled "Cells".
9. Take the "CM" tube containing 7 mL of conditioned media (from step 3.7.1) and rinse the bottom of the T75 flask twice with 3.5 mL portions of conditioned media, using a 5 mL serological pipette. After each rinse, place the 3.5 mL portions into the "Cells" tube. The purpose of this step is to remove all neurospheres (cells) from the T75 flask.
10. Centrifuge the "Cells" tube at 100 x g for 5 min at room temperature. While cells are spinning, obtain dPBS/glucose solution from 37 °C water bath and add the appropriate amount of trypsin and DNase according to Table 2.
11. After the "Cells" tube has stopped spinning, remove all the conditioned medium supernatant and transfer to the "CM" tube.
12. Take the tube labeled "trypsin inhibitor solution" from step 3.7.1. and add the volume of trypsin inhibitor indicated in Table 3. Use the same volume of trypsin inhibitor solution as was used for the trypsin solution.
13. Place trypsin inhibitor solution in the 37 °C water bath until needed.
14. Add trypsin solution to the cell pellet in the 15 mL tube and pipette up and down approximately 5-10 times with a 5 mL pipette. Close the cap of the tube and incubate in a 37 °C water bath for 5 min. After 5 min, retrieve the cells and pipette up and down 5-10 times. Then incubate in the 37 °C water bath for an additional 10-15 min.
15. In the last 5 min, move the trypsin inhibitor solution to the BSC.
16. Retrieve the cells after step 2.14 is complete and pipette the cells up and down until spheres are completely dissociated (20-30 times). Then immediately add the trypsin inhibitor solution and pipette up and down 10 times. This solution of dissociated cells and trypsin inhibitor will be referred to as the "cell suspension".
17. Count the number of cells in the cell suspension by following steps.
    1. Pipette 15 µL of Trypan Blue pre-mixture (containing 5 µL of 0.4% Trypan Blue and 10 µL of dPBS) onto a small weigh boat, and then add 5 µL of the cell suspension. Pipette up and down 3-5 times to mix. Then, pipette 10 µL of the Trypan Blue/cell suspension onto either side of a hemocytometer covered with a glass coverslip.
    2. Observe the cells under a light microscope and count the total number of cells in each of the four corner grids. Add these numbers together.
18. Repeat counting for the other side and average the two sides together.
19. Multiply this number by 10,000 to give the total number of cells per mL. Multiply this number by the total number of milliliters to calculate the total number of cells in the cell suspension. Calculate how much volume of cell suspension will be needed to seed 5 million cells per T75 flask.
    NOTE: Typically, after 9-10 days, a T75 will have 25-30 × 10⁶ cells. Therefore, if passaging all cells into new flasks, a total of 5-6 flasks is needed.
20. Add the volume of cell suspension calculated in step 3.19 to each T75 flask to obtain 5 × 10⁶cells per flask. If the volume is less than 5 mL, add additional condition medium to make up to a total of 5 mL. Then add 10 mL of new DFHGFPS media containing growth factors (Table 1) to the flask.
21. Repeat steps 1.17 and 1.18 and ensure the flask is labeled with the type of cells, the dates, the number of cells in the flask, and the passage number (this is the number of times the cell has been passaged). Then move to Steps 4-9.
22. If needed, seed extra cells into one T75 flask at the density up to 30 × 10⁶cells per flask overnight, and freeze the next day according to Step 4.

4. hNSC Freezing and Storage

1. The day after passaging, retrieve the flask from the incubator containing the cells for freezing (from step 3.19 and 3.22). Tilt the flask and remove all medium and cells from the flask and place in a 15 mL tube. Then centrifuge the tube at 216 x g for 5 min.
2. During the centrifugation process, prepare freezing medium (Table 4). For every 5 × 10⁶ cells prepare 1 mL of freezing medium. The number of cells was determined in step 3.19 when determining how many cells to put in each flask. This number will not have changed overnight.
3. Prepare cryo-preserve vials with labels of cell name, passage number, cell number, initials and date. After centrifugation, remove supernatant by aspiration and re-suspend cell pellet in freezing medium so that there are 5 × 10⁶ cells/mL. Using a 5 mL pipette, aliquot 1 mL per vial and seal tightly (5 × 10⁶ cells per vial).
4. Place vials in a cryo-preserve container with 250 mL of isopropanol (max usage of 5 times per replacement of isopropanol). Store in -80 °C freezer overnight, and then transfer to a liquid nitrogen tank storage system.

5. Plating adherent hNSC for validation

1. The day before passaging (steps 3.1-3.22), clean the BSC as in step 2.2, and obtain a 24-well plate and sterile glass 12 mm coverslips.
2. Using forceps, place a single coverslip on the bottom of each well of the plate.
3. Coat the wells containing cover slips with 0.01% poly-D-lysine (PDL) in sterile water and incubate for 1 h at 37 ˚C. For a 24-well plate, use 250 μL of PDL per well.
4. Remove the PDL from the wells, and then coat with 1 µg/cm² laminin/dPBS (250 μL per well). Incubate the plate overnight at 37 °C. If not needed the next day, seal the plate with parafilm by wrapping the parafilm around the edges of the plate and store at 4 °C.
   NOTE: Plates coated with laminin can be stored up to 2 weeks if sealed with parafilm as long as cover slips do not dry.
5. Passage cells as described in steps 3.1-3.22, and then determine the number of cells to seed into the 24-well plate with coated coverslips.
6. Remove any excess laminin solution from wells through aspiration, and rinse once with 0.5 mL of dPBS. Then seed cells into the wells at a density of 0.6-1 x 10⁵ cells/cm² per well. Use any remaining cells according to steps 3.16-3.21.
7. Atvarious times during the 24-well plate culture, fix and stain the cells for various stem cell markers following Step 9.

6. Priming to obtain GABA and glutamate neurons

1. Passage the cells as described in 3.1-3.22, and then determine the number of cells to seed into a 24-well plate as in step 5.6.
2. The day before priming, prepare the cover slips and coat plate as described in steps 5.1-5.4.
3. On priming day, retrieve the flasks containing 2-3-day cells. Place in a 15 mL tube and pellet the cells by centrifuging at 100 x g for 5 min.
4. Prepare the epidermal growth factor/leukemia inhibitory factor/laminin (ELL) priming medium according to Table 5.
5. Resuspend cells with ELL medium.
6. Seed suspended cells into a 24-well plate (follow step 5.6).

7. Priming to obtain motor neurons

1. Follow steps 6.1-6.4.
2. Prepare the basic fibroblast growth factor/Heparin/Laminin (FHL) priming medium according to Table 6.
3. Follow step 5.6.

8. Zika Virus Infection

NOTE: ZIKV is highly sensitive to temperature, so it is imperative that ZIKV stock are stored at -80 °C and freeze/thawing cycles are avoided. Keep working aliquots on ice.

1. Passage cells as described in steps 3.1-3.22. 3-4 million cells in total are needed to seed a 24-well plate, therefore when passaging, put the number of cells needed for infection and seeding in a flask when passaging.
2. If seeding cells into a 24-well plate for staining, prepare the coverslips and plate according to steps 5.1-5.4.
3. After passaging, place 1 mL of cell suspension from step 8.2 into a 1.5 mL microcentrifuge tube, and centrifuge at 200 x g at room temperature for 5 min. Then remove the supernatant by aspiration.
4. Resuspend pellet from step 8.3 in ZIKV stock to acquire a multiplicity of infection (MOI) of either 1 to 10. The volume of ZIKV solution should not exceed 0.5 mL. For mock treatment (control cells), resuspend cells in the same volume and type of medium used in ZIKV stock.
   NOTE: ZIKV stock preparation and infection are detailed in a previous publication¹¹. Mock infections are also conducted with medium.
5. Incubate above cell/ZIKV mixture at 37 °C for 1 h. Invert the tube 2-3 times, and then centrifuge mixture for 5 min at 216 x g at room temperature to obtain a cell pellet. Resuspend pellet in dPBS and centrifuge again for 5 min at 216 x g at room temperature.
6. Remove the supernatant by aspiration, resuspend cells with appropriate medium, and load infected cells into flask or plate needed for the experiment. If seeding the cells into a 24-well plate resuspend the pellet in 12 mL of appropriate medium, and then add 500 μL of suspension to each well.
   NOTE: At this point either maintain the cells in growth media to observe effects of ZIKV on stem cells, or go to steps 6 or 7 to study the effect of ZIKV on the differentiation process.

9. Staining procedure

1. To fix the cells, remove medium, rinse once with 0.5 mL per well (24-well plate) of ice cold phosphate-buffered saline (PBS).
2. Remove PBS, and cover cells with 0.5 mL of 4% paraformaldehyde in PBS and leave at room temperature for 20-30 min.
3. Remove the paraformaldehyde and rinse cells three times with 0.5 mL of PBS, leaving the third PBS rinse on for 10 min at room temperature.
4. At this point either leave the PBS on overnight and store the plate at 4 ˚C, or begin staining after the 10 min rinse.
5. After the 10 min PBS rinse, block the cells for 45-60 min in a stationary position with a 0.5 mL solution of 5% normal goat serum (NGS), 0.3% bovine serum albumin (BSA), 0.25% Triton X-100 in Tris-buffered saline (TBS). For example, to make 1 mL of blocking solution use: 10 µL of 25% Triton X-100, 50 µL of 100% NGS, and 100 µL of 3% BSA mixed in 840 µL of TBS.
6. During the blocking step, prepare the primary antibody solution, making sure all reagents are kept on ice. For validating hNSCs, proteins such as Nestin can be targeted using the appropriate antibody against them. To validate ZIKV infection, use an antibody against ZIKV coat protein¹¹. For differentiation, use markers such as class III β-tubulin to identify neuronal populations while glial fibrillary acidic protein (GFAP) can be used to identify astrocytes. The concentrations of primary antibodies are empirically determined depending on vendor and lot. We have used most antibodies from 1:100 to 1:2000 dilution.
   1. Centrifuge the primary antibody at 12,000 x g for 2 min at 4 °C, and then make a dilution; e.g. to make a 1:1000 dilution of antibody, add 7.2 µL of desired antibody into 7.2 mL of 0.25% Triton X-100 in TBS.
7. Add approximately 300 µL of antibody solution to each well of a 24-well plate, and incubate the cells with primary antibody in a stationary position for 2 h at room temperature or at 4 ˚C overnight in a stationary position.
8. After the incubation, wash cells with 0.5 mL of TBS three times at room temperature for 10 min each in a stationary position.
9. Centrifuge the secondary antibody at 12,000 x g for 2 min at 4 °C, then dilute in 0.25% Triton X-100/TBS solution. Make enough antibody solution to add 300 µL to each well. Here, dilute the fluorescent secondary antibodies 1:500 (Figure 3).
10. After the final TBS wash, add 300 µL of the secondary antibody solution to each well and incubate in the dark for 1 h at room temperature in a stationary position. From this point on, all steps must take place in a dark room.
11. Wash the cells three times with TBS for 10 min each in a stationary position.
12. During the last 10 min wash, dilute the nuclear marker DAPI nuclear stain to the recommended concentration in TBS (typically 1:1,000 – 1:5,000). Make enough solution to add 300 µL to each well.
13. After the final wash, add 300 µL of DAPI solution to each well and incubate for 5 min at room temperature in a stationary position. Then wash once with 0.5 mL of TBS.
14. Use an anti-fade mounting medium for fluorescence and 1 drop (10-15 µL) per coverslip that will be placed on the slide. Carefully use tweezers to remove coverslips from the 24-well plate wells and place the cell surface down on the mounting medium drop on the slide.
15. Once the coverslips are placed on the slide, place the slide in a flat slide-holder. Make sure to label slides will all relevant information needed to identify the cells on that slide, then cover the holder with aluminum foil to keep slides protected from light.
16. Store the slides at 4 ˚C and allow the mounting medium to solidify overnight. The following day, observe the slide using an epifluorescent microscope with UV-2E/C filter (340-380 nm), GFP HYQ BP filter (450-490 nm), or Y-2E/C Texas Red filter (540-580 nm), 20X magnification.