Neuroscience
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Establishing Mouse Models for Zika Virus-induced Neurological Disorders Using Intracerebral Injection Strategies: Embryonic, Neonatal, and Adult
Chapters
Summary April 26th, 2018
Here we describe a method for establishing a model of Zika virus-induced microcephaly in mouse. This protocol includes methods for embryonic, neonatal, and adult-stage intracerebral inoculation of the Zika virus.
Transcript
The overall goal of the surgical methods presented here is to establish in vivo models of intracranial Zika virus infection. This method can help to address the key question in the field, such as pathology and the mechanism of Zika virus infection in the brain at both early and the later developmental stage as well as adult stage. The main advantage of this technique is that the direction infection of the brain can easily be achieved.
The implications of this technique extend toward therapies or diagnoses of Zika virus-induced microcephaly or other neuro-developmental disorders. Although this method can provide insight into Zika virus pathogenesis, it can also be applied to study other flavivirus infection in the brain, such as dengue virus. Just prior to surgery, load the Zika virus injection mix into a prepared glass injection needle by first assembling the gas-tight 50-microliter injection syringe Luer-lock attachment and tubing.
Saturate the assembled syringe and tubing with mineral oil. Once saturated, attach the needle and draw approximately six to seven microliters of virus into the needle. After anesthetizing a dam pregnant with E14.5 embryos, test the depth of anesthesia by pinching the toe or tail.
When a surgical plane of anesthesia is achieved, insure that the dam is laying supine on a heating pad. Shave the surface of the abdomen and sterilize with iodine and alcohol three times. Drape the skin surrounding the surgical site with sterile cloths to avoid contamination of the incision and the instruments.
Pinch and lift the skin of the abdomen with forceps, then use sterile scissors to make a 1 to 1 1/2-centimeter cut in the lower abdomen at the medial-sagittal line and expose the embryos. Next, cut a slit in the middle of a small sterile gauze and place over the surgical opening. Hydrate the opening and gauze with sterile saline.
Then, pull the embryos through the slit to rest on the gauze, taking care to not remove more than four or five embryos at once. Hydrate the embryos with sterile saline prior to and throughout the inoculation to ensure they do not dry out. Illuminate the embryo to be injected with a lamp to visualize the head and skull sutures, then gently place a spatula under the head of the embryo to be injected and manipulate the embryo until the head is pushed up directly against the uterine wall.
Hold the embryo in place with the non-dominant hand. Use the syringe to inject Zika virus or control medium into the lateral ventricles of the embryo brain. For the E11.5 stage intrauterine injection, use a one-milliliter syringe and a 27 gauge needle to inject Zika virus or a control medium into the intrauterine space.
To improve retention of the pregnancy, avoid injecting the two embryos next to the ovaries and the two embryos next to the upper vagina. If performing an intrauterine injection, inject into the intrauterine space or into the placenta. Place the embryos back into the pregnant dam and fill the abdominal cavity with approximately 0.5 milliliters of sterile saline.
Suture both the abdominal peritoneal muscle and external skin layer with 4-0 sterile sutures. Keep the dam on a heating pad and monitor while recovering from anesthesia. Thaw the virus on ice, then load the syringe by filling it with saline to reduce the dead volume in the syringe.
Draw up to 0.75 microliters of air to separate the saline solution from the injection material. To set up a warmed, humidified recovery chamber, place a closable container with a sterile gauze on a heat box set at 37 degrees Celsius and soak with saline. After setting up the microinjector pump and controller, load the injection syringe with virus and sterilize the pump head mount of the surgical setup with 70%ethanol.
Then, after cryo-anesthetizing the neonatal pups for five minutes, ensure sufficient anesthesia for injection by the lack of response to a tail or toe pinch. Place the pup on the head mount. Sterilize the surface of the head with iodine then ethanol, and then wipe dry with an alcohol wipe.
Mark lambda with a black marker, then measure the distance from lambda to the edge of the eye and, using stereotaxic coordinates, calculate the 2/5 distance from lambda to the eye. Next, use a 26 gauge needle to carefully and superficially puncture the scalp and skull at the injection location and create an opening for the injection needle. Now lower the injection needle into the puncture site and, once the needle has just breached the skin, calculate a one-millimeter depth for injection into the lateral ventricle using the stereotaxic instrument.
Once the needle is lowered to the injection depth, inject one microliter of virus at a rate of 10 nanoliters per second. Wait 30 seconds after injection, and then retract the needle in 0.5-millimeter increments by rotating the screw dorsally, waiting 30 seconds after each increment to reduce leakage of injected virus. After finishing the injection, transfer the pup to the pre-warmed, humidified chamber and monitor until recovered.
Immobilize the head of an anesthetized adult mouse in the stereotaxic instrument and place a heating pad under the mouse. After a toe or tail pinch to insure a surgical plane of anesthesia, shave the scalp starting behind the eyes to the beginning of the ears. Sterilize the exposed skin with iodine and 70%ethanol.
After making a 0.5-centimeter incision along the medial-sagittal line of the head in the sterilized location to expose bregma, clear the surface of the skull of meningeal tissues while simultaneously pushing the skin away from the injection sites. Align the surgical drill bit with bregma, and then identify the injection site by the stereotaxic coordinates. Using the control foot pedal, slowly drill into the skull until the hole is clear for the needle.
Replace the drill bit with the microinjector pump and gas-tight 10-microliter syringe. Leave a small air bubble between the saline solution and the virus and draw up four to five microliters of virus. Lower the needle to 1.5 millimeters below the brain surface from the brain surface and inject one microliter of virus at a rate of 10 nanoliters per second, injecting the one microliter over 1 1/2 minutes.
After removing the needle from the brain in increments as before, use forceps to loosen the skin and re-cover the exposed skull. Once the scalp has been sutured with 4-0 sutures, remove the mouse from the stereotax and monitor on a heating pad until recovered. Neuro-progenitor cells are immuno-labeled with Pax6, seen in red, and Zika virus via flavivirus antigens, seen in green, four days after injection at E18.5.
The embryonic brain is infected with the MEX1-44 strain of Zika virus across all layers of the developing neocortex. The TCID50 assay using tissue from post-natal day three brain tissue inoculated at E14.5 shows that the Zika virus can efficiently replicate and grow in developing brains. This image shows MEX1-44 Zika virus infection at P13 cerebral cortex after P0 injection.
Zika is detected using antibody against flavivirus group antigen seen in red. These images show the fluorescent beads that were used to practice injection location and lateral ventricle injection success in adults. While attempting this procedure, it's important to remember to keep track of the embryos injected, have appropriate controls, and keep the surgical wounds small.
Following this procedure, other methods like intraperitoneal or arterial injection can be performed to answer the additional question like how Zika virus cross the placenta. After watching this video, you should have a good understanding of how to generate the in vivo mouse model of Zika virus infection in the brain.
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