Neuroscience
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Rat Model of Widespread Cerebral Cortical Demyelination Induced by an Intracerebral Injection of Pro-Inflammatory Cytokines
Chapters
Summary September 21st, 2021
The protocol presented here allows the reproduction of a widespread grey matter demyelination of both cortical hemispheres in adult male Dark Agouti rats. The method comprises of intracerebral implantation of a catheter, subclinical immunization against myelin oligodendrocyte glycoprotein, and intracerebral injection of a pro-inflammatory cytokine mixture through the implanted catheter.
Transcript
The overall goal of this procedure is to generate a widespread demyelination of the cortex of both cerebral hemispheres in a rat model using sub-clinical immunization against myelin oligodendrocyte glycoprotein, followed by interest cerebral injection of cytokines through an implanted catheter. This method can help answer key questions in inflammatory demyelinating diseases of the brain, such as multiple sclerosis. The main advantage of this technique is that it can generate inflammation induced gray matter demyelination in the cortex of both cerebral hemispheres without leading to white matter plugs.
The implications of this technique extend toward a better understanding of mechanisms of cortical demyelination that is observed mostly in progressive types of multiple sclerosis. To begin this procedure, assemble the catheter and the catheter cap with the inlet. Next, cut the catheter to a length of two millimeters with a scalpel.
Then shave the head of an anesthetized rat between the ears using the electric shaver. Place the rat in the stereotactic frame and secure its head using the ear bars and bite plate. Ensure that the head is horizontal and check for its stability by applying pressure on it with finger or forceps.
Afterward, apply lubricating eyedrops to prevent cornea dryness during the surgery, cover the eyes with an opaque material to prevent any surgical light exposure. Next, clean the shaven area by alternating the application of 70%ethanol and 10%povidone iodine complex. To implant the catheter, make a longitudinal incision of about two centimeters in length along the midline.
Then use bulldog clamps to hold the skin to the sides. Remove the blood using a cotton tip applicator. Afterward, remove the periosteum, clean the tissue with the cotton tip applicator, expose the skull and allow it to dry for about one minute.
Next, identify the anatomical landmarks, lambda, bregma and medial suture. With the drill installed on the stereotactic frame, position the tip of the drill bit at the bregma as the starting point. Move two millimeters posterior from the bregma, and 2.4 millimeters laterally to the medial suture.
Then, drill a 0.5 millimeter diameter hole for the catheter at this position. Drill three more holes of 1.3 millimeter diameter for the anchor screws a few millimeters away from the first hole. Remove the bone dust by irrigation with around one to two milliliters of sterile PBS and clean the skull.
Subsequently, tighten the anchor screws by two to three full turns. Insert a two millimeter length catheter through the first hole perpendicular to the skull surface. While still holding the catheter, apply a little dental cement and let it polymerize with a brief exposure to the dental curing light in order to stabilize the catheter.
Then apply more dental cement around the catheter, anchor the screws, and solidify the dental cement with the dental curing light for about 15 to 30 seconds, confirm that the cement is hardened. After implantation, close the skin with resorbable sutures anterior, and posterior to the catheter. Then inject the antidote mixture subcutaneously.
Next, administer 2.5%enrofloxacin by subcutaneous injection for prophylactic antibiotic treatment. Return the animal to the modified cage and keep it under observation for one to three hours with an application of infrared light to avoid hypothermia. Repeat the enrofloxacin treatment and administer carprofen at one milligram per milliliter subcutaneously for pain relief by injection the day after the surgery.
To prepare the immunization mixture, connect two 10 milliliter lower lock tip glass syringes to the short arms of a three-way stopcock, and close the third outlet with the long arm. Next, pipette one milliliter of IFA and 50 micrograms of rMOG together. And adjust the mixture to a final volume of two milliliters with sterile PBS.
Place the diluted IFA and rMOG mixture in the open syringe, then insert the piston gently whilst maintaining a loose pressure on the opposite piston. Emulsify the inoculum by driving it from one syringe to the other via pushing the pistons back and forth until it is white and viscous. Next, fix a one milliliter lower lock syringe to the open short arm of the three-way stopcock and fill it with inoculum.
Distribute all inoculum to one milliliter syringes and keep them on ice until the injection. Subsequently, inject 200 microliters of the IFA and rMOG mixture subcutaneously at the tail base under a temporary isoflurane anesthesia using a 21 gauge needle. For intracerebral cytokine injection, adjust the length of the connector cannula to two millimeters.
Fill a one milliliter syringe with the cytokine mixture. Then, connect the syringe to the connector cannula. Next, fill the cannula with the cytokine mixture, avoid creating any bubbles.
Then, mount the syringe onto the programmable syringe pump and program it to inject at a rate of 0.2 microliters per minute. Start the pump and keep it working in order to avoid an air bubble formation at the tip of the cannula. Afterward, remove the catheter cap with the inlet.
Insert the connector cannula into the catheter, screw and tighten it. Then allow the injection to proceed for 10 minutes before stopping the pump. Leave the cannula inside the catheter for 20 minutes to allow the injected volume to fully diffuse.
Subsequently, unscrew the connector cannula and remove it slowly to avoid a vacuum effect. Reattach the catheter cap with the inlet and screw it. Allow the animal to recover from the anesthesia in a cage.
Cortical demyelination could be assessed at different time points after a cytokine injection by immuno histochemistry for PLP. Figure 6A shows intact PLP immunoreactivity at day 15 in a moggy immunized control animal that received only sterile PBS through the implanted catheter. On day one after the cytokine injection, demyelination could already be detected in MOG prime animals.
Albeit, only in the close vicinity of the catheterized area. The PLP immunoreactivity stays intact in the contralateral cortex one day post cytokine injection. On day three, a gradual increase in the loss of the PLP immuno reactivity, which spreads in the ipsilateral cortex could be observed.
Contralateral cortical demyelination could also be detected at day three, but it is rather restricted to the area beneath the anchor screws. Between days nine to 15, demyelination affects large parts of the cortex of both hemispheres. The cortical demyelination is sustained for up to 30 days post cytokine injection in both hemispheres with only a partial remyelination.
Figure 6J shows a quantification of PLP loss in the cortical gray matter after the intracerebral cytokine injection. After watching this video, you should have a good understanding of how to generate demyelination of gray matter using subclinical immunization against myelin proteins, followed by intracerebral injection of cytokines through the implanted catheter. This animal model could help researchers in study of progressive types of multiple sclerosis, layer gray matter demyelination is a whole mark.
The implanted catheter enables multiple rounds of demyelination or intercerebral delivery of potential therapeutic drugs undergoing preclinical investigation without causing injection induced trauma.
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