The overall goal of this procedure is to accurately locate a small stroke within murine white matter to model this common form of subcortical white matter stroke. This method can help answer key questions in the stroke field, such as mechanisms of axonal injury, neuronal response to subcortical stroke, and how the cellular elements of white matter respond during both the acute and repair phases of stroke. The main advantage of this technique is that it can be used to precisely localize a stroke to the white matter, and can be used with a wide variety of murine mouse models.
Generally, individuals new to this method will struggle, because of the small amount of white matter leading to the mislocalization of the stroke. Minor adjustments to the injection angle or stereotactic coordinates for each strain of mouse, or stereotactic setup may be required. Begin by first affixing a pulled glass pipette to tubing attached to a vacuum line.
Insert the pulled end into the L-Nio solution, or L-Nio plus fluorescent tracer. Start the vacuum, and apply suction until at least two millimeters of the 0.5 millimeter diameter portion of the pipette is filled. Put the filled pipette to one side.
Then place the mouse into a stereotactic apparatus equipped with a stereotactic microscope. After anesthetizing and prepping the mouse for surgery according to approved procedures, check the depth of anesthesia with a toe pinch. There should be no response.
Next, adjust the injection arm to 36 degrees. Affix a pulled glass pipette holder to the distal end of a low volume pressure injection system, and attach it to the injection arm. After making a 1.5 centimeter midline scalp incision to expose the skull, dry the surface skull with a cotton swab.
Then, while looking through a stereotactic microscope, at one to three X magnification, use a micropoint tool to remove any overlying periosteal tissue. Mark bregma with a fine-point marker. Then use a surgical drill equipped with a fine-tipped surgical drill bit to drill a two millimeter elliptical craniotomy, beginning posteriorly at bregma, and extending anteriorly just left of the midline.
Remove bone fragments and overlying soft tissue so that the cerebral cortex can be visualized. Keep the cortical surface moist by intermittently applying drops of sterile saline. Next, affix the filled pipette to the injector arm.
Align the distal end of the pipette with bregma, and zero the stereotactic coordinates. Advance the pipette to the first injection point using the anterior, posterior, and medial-lateral coordinates listed in table one. Then, advance the pipette to the cortical surface, and zero the dorsal-ventral measurement.
Slowly lower the pipette to the dorsal-ventral coordinates. Ensure that the stereotactic microscope magnification is set to three X, and that the calibrated reticle can be clearly viewed. View the angled pipette from the side so that the air fluid meniscus has a sagittal view.
The meniscus should appear in the same focal plane of both the inner and outer wall of the pipette. Now, using a local pressure injection system set at 20 PSI for 20 millisecond pulses, inject 100 nanoliters of L-Nio into the brain. After injecting the total volume, wait five minutes to prevent reflux up the pipette track.
Then, withdraw the pipette, and repeat the injection procedure at the second and third set of coordinates provided in table one. After the final injection, remove the pipette and place enough bone wax to fill the craniotomy site. Finally, approximate the edges of the scalp wound, and bind with dermal adhesive.
After administering postoperative analgesia, place the animal in a clean cage. Supply postoperative antibiotics in the drinking water for five days, or until sacrifice if less than five days. After sacrificing and decapitating the mouse according to approved procedures, use sterile scissors to open the skull posteriorly, and then use a spatula to gently remove the overlying skull.
Next, insert a sterile four millimeter spatula at the front of the brain to sever the olfactory bulb and optic nerves. Then, gently lift the brain out of the calvarium and place into ice cold dissection buffer. Using a brain block and sterile razor blades, cut two to three millimeter slices containing the affected region, and place into cold dissection buffer.
Under a dissecting microscope, identify the white matter underlying the motor cortex in the injected hemisphere. At earlier post-stroke intervals, injection of L-Nio mixed with common laboratory dyes, such as fast green, can allow visual identification of the stroke. At longer post-stroke intervals, the region may be visually identified by focal necrosis and myelin pallor.
Once identified, use a scalpel to carefully dissect the region of white matter containing the stroke. Remove overlying cortex and underlying striatum, as desired. As shown here, immunofluorescent labeling for neural filaments, shown in red, demonstrates the degree of axonal loss seven days after stroke, using the medial approach.
Using the posterior angled approach, the white matter stroke lesion is targeted just above the lateral ventricle and shows intense microglial reactivity, as detected by immunolabeling for IBA-1. Two astrocyte intermediate filament markers, vimentin, shown in red, and glial fibrillary acidic protein, shown in green, reveal changes in morphology of white matter astrocytes after stroke. Coinjection of fluorescent dextran amine at the time of stroke induction allows identification of individual neurons with axons injured by stroke.
Most of the labeling occurs in axons within layer five and six neurons in primary sensorimotor cortices overlying the stroke. This image represents seven days after stroke. Once mastered, this technique can be done in 45 minutes, if it is performed properly.
After watching this video, you should have a good understanding of how to produce focal white matter stroke in a mouse, and prepare the brain for a variety of downstream processing, useful to answer important questions in stroke and neuro repair.
