视网膜缺血再灌注损伤的小鼠模型通过眼压升高

The overall goal of this technique, is to provide an experimental model of Retinal Ischemia-Reperfusion Injury, and it's pathologic consequences. This method is commonly used to investigate the mechanisms underlying Retinal Ischemia-Reperfusion, as well as to evaluate potential therapeutic approaches for treating associated pathologies. The main advantage of this technique is that it creates an injury highly targeted to ocular structures, while causing minimal damage to surrounding tissues.

The usefulness of this technique extends towards the understanding and three month of multiple manifestations of Retinal Ischemia-Reperfusion Injury, including:inflammation, oxidative stress, and stare dysfunction in the retinal neurovascular unit. Video demonstration of this method is critical because the surgical techniques for cannula placement may be difficult to reproduce without a visual guide. Before beginning the procedure, warm an empty cage on the table.

Then insert the sharp end of the primary set pre-pierced Y-site tubing into a bottle of 0.1%Heparin sodium, and hang the bottom of Heparin sodium from an IV pole extension. Snap open the air filter cap on the tubing. Then elevate the Heparin sodium bottle to 163 centimeters from the tabletop to the peak of the sodium Heparin drip, to produce a Heparin sodium flow pressure of 120 mililiters of mercury.

Open the primary set pre-pierced Y-site tubing, to allow the Heparin sodium to fill the tubing, and connect the primary set pre-pierced Y-site tubing to the five valve manifold. Then, using Luer male to Luer male tube fittings, connect 30 gauge half-inch needles to the five port manifold. Insert each of the needles into its own ten inch segment of 30 gauge tubing.

Then, using hemostats, break new 30 gauge half inch needle tips and insert their blunt ends into open ends of the 30 gauge tubing. Next, use tape to arrange the needle tubes such that the tubes connected to the inner ports are positioned on top of the tubes connected to the outer ports to prevent tangling of the tubing during the Anterior Chamber Cannulation. Now, turn on the 0.1%Heparin sodium flow to the five valve manifold and to each individual port for two to three minutes to remove any air bubbles.

Then, turn off all of the ports on the five valve manifold. To perform the cannulation, after administering the anesthesia, place all of the mice into the warm cage on the surgery table, confirming the achievement of deep anesthetization by lack of response to toe pinch after five to ten minutes. Then administer one drop of Tropicamide for pupil dilation, and one drop of Proparacaine for local anesthesia into each of the animals'eyes, and arrange the mice in the order of anesthetization, so that the first mouse anesthetized will be the first mouse to be cannulated.

While waiting for the eye drops to take effect, prepare four inch straight pieces of tape, pulling tightly on the tape before tearing, and setting aside one piece of tape per animal. Note that failing to pull tightly on the tape, will result in curling of the tape. When the first mouse is ready, place the animal under a surgical microscope.

Then, turn on the first 0.1%Heparin sodium port on the five valve manifold, and use a pair of forceps to gently proptose one eye. Next, insert the 30 gauge cannula needle into the anterior chamber approximately halfway between the zonular fibers and the apex of the cornea. Taking care to avoid penetrating the conrea a second time.

Note that a gentle twisting motion is used to insert the cannula deeply into the anterior chamber. Now, press the 30 gauge tubing against the tabletop to minimize any movement of the inserted cannula while reaching for a piece of tape, and use the tape to secure the tubing to the table. Once the tube has been secured, record the start time of the surgery.

Use the microscope to verify that there is no leakage, and that the Ischemia-Reperfusion eye is larger than the contralateral eye, to establish the presence of ocular distension. The apply Hypromellose to both eyes every 30 minutes, until the end of the experiment to lubricate the cornea, and to seal any micro leaks. Should an animal stir during the procedure, lift the tail and administer a Ketamine booster intraperitoneally.

90 minutes after cannulating the first animal, gently pull the cannula from the anterior chamber, and lubricate both eyes with lubricating jelly. As subsequent cannulae are removed, place each mouse back into the cage on the surgical table, until the animals are fully recovered. After all of the cannulae have been removed, disinfect them with alcohol wipes.

Then use a 60 milliliter syringe filled with air to expel the Heparin sodium from the five valve manifold, 30 gauge needles, 30 gauge tubing, and cannulae. Next, rinse all of the instruments with a 60 milliliter syringe filled with distilled water. Finally, expel the distilled water from all of the materials with 60 milliliters of air, and store the disinfected and rinsed equipment for later use.

NeuN immunolabeling of neuronal nuclei reveal significant nuronal cell loss, following Ischemia-Reperfusion insult, by confocal microscopic imaging, seven days after Ischemia-Reperfusion Injury. Indeed, manual quantification of the NeuN positive cells in control, and Ischemia-Reperfused eyes, demonstrates a decrease in the gangelion cell layer neuron counts after injury, indicating the presence of Ischemia-Reperfusion induced cell death. Lower a-wave and b-wave amplitudes documented by electroretinography also signify the occurrence of Ischemia-Reperfusion induced impairments in retinal neuronal function compared to control, non-injured eyes.

Once mastered, this technique can be performed on ten animals in approximately five hours, including the final two hours spent waiting for the mice to recover from anesthesia. While attempting this procedure, it's important to maintaim deep anesthesia. As an effective sedation will prevent displacement of the cannula by animal movement.

Following this procedure, other methods, like electroretinography can be performed to answer additional questions about retinal function. This technique has enabled researchers in the ophthalmology neuroscience and vascular biology fields to explore retinal neurovasuclar pathogenesis, and therapeutic intervention in rodents. After watching this video, you should have a good understanding of how to create a targeted retinal injury, though the cannulation of the ocular anterior chamber.

Don't forget, that working with surgical instruments can be extremely hazardous, and that precautions, such as using an effective sharps disposal, should always be taken while performing this procedure.