The overall goal of this procedure is to model zika virus infection in the developing human brain using stem cell derived cerebral organoids. This method can help answer key questions in the biology field such as, which cells are capable of being infected and which proteins are involved with the infection pathway. The main advantages of this technique are that it mimics early human infection, can be utilized in high throughput assays and can be combined with focus genetic techniques such as CRISPR.
This method can provide insights into the mechanism of zika virus infection. It can also be applied to other systems such as, drug screens and virus pseudotyping. Using regular stem cell maintenance methods, bring cultures to between 50%and 70%confluence.
Inspect the cultures using bright field microscopy at 10 to 20x magnification and ensure that the colony has a healthy morphology with no detectable differentiation. Bring the xeno free stem cell maintenance media to 37 degree celsius in a hot water bath and thaw two milliliters of enzymatic detachment reagent and a 50 microliter vial of the stock rock inhibitor at room temperature. Next, prepare an ultra low attachment U bottom 96 well plate, a multi channel p200 pipette and a 25 milliliter reagent reservoir.
Next, aliquot 45 milliliters of the media into a 50 milliliter conical tube. Add 45 microliters of the rock inhibitor and mix in the inhibitor thoroughly by triterating the mixture, followed by two to four in versions. Vacuum aspirate two wells of a six well plate containing stem cells and quickly add one milliliter of enzyme free detachment reagent to each well, then, incubate the plate at 37 degree celsius for four minutes.
Next, vacuum aspirate the treated wells and add one milliliter of enzymatic detachment reagent to each well. After a five minute incubation at 37 degree celsius, remove the plate from the incubator. Gently tap on the plate to break up any aggregated cells.
Next, inspect the cells under the microscope. If large cell clusters are still visible, incubate the plate for two additional minutes at 37 degree celsius. Repeat the step until clusters are no longer visible by the naked eye.
Once the cells are properly dissociated, add one milliliter of xeno free stem cell maintenance media to each well, to deactivate the dissociation enzymes. Pull the cell suspension into a 50 milliliter conical tube and take a small aliquot for cell counting. While centrifuging, count the cells and determine the re-suspension volume needed to achieve 60, 000 cells per milliliter suspension.
Carefully remove the supernatant and re-suspend the cells at 60, 000 cells per milliliter in media containing the rock inhibitor. Using a five milliliter pipette, mix the cells gently five to 10 times in order to ensure a uniform cell suspension. Immediately add the cell suspension into reagent reservoir and use a multi channel p200 pipette to transfer 150 microliters into each well of an ultra low attachment you bottom 96 well plate.
Next, centrifuge the plate at 150 x G for one minute and then place the plate into a 37 degree celsius incubator. Do not disturb the plate for 48 hours. Starting on day two, prepare 17 milliliters of media containing 17 microliters of the stock rock inhibitor in a 50 milliliter conical tube.
Warm the mixture to 37 degree celsius in a hot water bath. Take the organoid plate from the incubator and using a multi channel p200 pipette, slowly draw up 75 microliters of medium from the edges of the wells in the first row. Then, quickly expel it back into the well to lift out loose cells and organoids.
Repeat this dispersion process for each row. Wait for 15 seconds to ensure that the organoids have sunk to the bottom of each well and then slowly and carefully aspirate 75 microliters of medium from each well using the multi channel p200 pipette and dispense it into a waste reservoir. Next, transfer media containing the rock inhibitor into a reagent reservoir, dispense 150 microliters of media into each organoid well and then incubate the cells at 37 degree celsius.
Warm 17 milliliters of fresh culture medium to 37 degree celsius, this time without any rock inhibitor. Using a multi channel p200 pipette set to 100 microliters, repeat the dispersion process previously shown and wait 15 seconds to ensure that the organoids have sunk to the bottom of their wells. Then, carefully aspirate 125 microliters of medium from each well and replace with 150 microliters of warmed media.
Place the plate into the 37 degree celsius incubator. Prepare the medium according to the recipe shown here and then sterile filter the mixed solution in a 500 milliliter filter. From day four until the cells are infected on about day 24, conduct regular maintenance every other day using neural induction medium.
Once filtered, warm 17 milliliters of the neural induction medium to 37 degree celsius and store the rest at four degree celsius. Using a multi channel p200 pipette, set to 100 microliters, disperse all wells of the organoid plate has previously shown. Wait 15 seconds to ensure that organoids have sunk to the bottom of their wells.
Carefully aspirate 125 microliters of medium from each well and replace it with 125 microliters of fresh neural induction medium. Repeat this neral induction feat every other day until the organoids are ready for zika virus infection. Bring earle's 1x balanced salt solution, 1x PBS and neural induction medium to 37 degrees celsius in a hot water bath.
Next, dilute zika virus of a known concentration in 1x earle's solution to the target multiplicity of infection, which is typically between 0.1 and 10. Using a p200 pipette, carefully remove all medium from each organoid well. Then, quickly wash the organoids with 200 microliters of warmed 1x PBS.
When removing the media from each well, it is critical that one does not touch the organoid or suck it into the pipette. This can cause irreparable damage to the organoid. Should this happen, it is best to discard the organoid.
Next, carefully remove all the remaining liquid from each organoid well using a single channel p200 pipette. Then, quickly add 50 microliters of either 1x earle's solution for a mock infection or zika virus solution to each well. Ensure that each organoid is completely submerged when finished, place the plate back into a 37 degree celsius incubator for two hours.
After a viral exposure is completed, wash each well of the organoids with 200 microliters of PBS. Allow organoids to settle for 15 seconds and then remove the PBS using a single channel p200 pipette. Finally, add 200 microliters of fresh neural induction medium to each well and place the plate back into the incubator.
Staining with DAPI, phospho-vimentin, TBR2 and MAP2 can be combined to show the cortical structures forming within the organoids. These structures include the ventricular zone, the sub-ventricular zone, the intermediate zone and the cortical plate within each rosette. Culturing the organoids to day 108, allows one to observe the later stages of development.
While cortical layering is not as prominent in this type of organoid, the natural switch to gliogenesis does occur much like it does in-vivo. Three days after zika virus infection, a difference in organoid size becomes visible. The scale of this difference depends on the viral multiplicity of infection that is applied to the organoids.
This difference in organoid size will increase over the following week until the infected organoids begin to break apart. After three days, there will also be an increase in cellular debris in the infected wells compared to the mock wells. While attempting this procedure, it's important to follow safe laboratory practices since viable infectious materials are being used.
Following this procedure, other methods like immunohistochemistry or RNAC can be performed in order to answer additional questions about the biology involved in neural developmental zika virus infection.
