The overall goal of this procedure is to track the remodeling of the vasculature in the mouse brain during the progression of a chronic disease over extended periods of time. This method can help answer key questions in the field of neuroscience, such as the role of cerebral vasculature remodeling in diseases of the central nervous system. The main advantage of this technique is that it is minimally invasive and without neural inflammation, making it ideal to study pathologies where inflammation is actually implicated.
This technique allows the visualization of pial and and penetrating cortical vessels, complementing histological techniques direct at the visualization of microcapillaries in the parenchymal of the blood-brain barrier. Demonstrating the procedure will be Margarita Arango-Lievano. She's a post-doc from my laboratory.
To begin this procedure, clean the surgical bench and microscope plate with 70%ethanol and cover them with a clean, absorbent cloth. Next, check the appropriate level of anesthesia with paw or tail pinches. Then apply sterile ophthalmic ointment to the eyes and shave the fur from the nose to the ears with a razor blade.
Afterward, sterilize the skin with povidone iodine antiseptic solution. Place the mouse under the binocular microscope and make a skin incision from the ears to the nose. Pull the skin sideways to expose the skull.
Then tear the periosteum and repeatedly rinse the skull with sterile ACSF to stop possible bleeding. In this procedure, remove the eye ointment with a cotton tip. Then apply a drop of topical ophthalmic anesthetic.
Gently pressure the skin around the eye socket to achieve protrusion. After that, inject FITC conjugated with dextran in the retro-orbital sinus. Subsequently, apply sterile ophthalmic ointment to the eyes.
Ensure that the skin is retracted and the skull is clean and dry around the selected imaging area. Then apply a small amount of cyanoacrylic glue around the window of the head mount device and glue the head mount device around the target region. Subsequently, apply gentle pressure to the head mount for several seconds.
Pull the ends of the skin to the edge of the window of the head mount device, making sure the area is dry. Secure the skin and the edge of the window with a small amount of cyanoacrylic glue and wait until dry. Afterward, secure the animal at the stage using the head mount device.
Rinse the bone in the target area with sterile ACSF and ensure that the well between the skin and the head mount device is perfectly sealed. Following that, wrap the mouse in a survival blanket to maintain normothermia. Now perform thinning of the skull in ACSF solution to attenuate vibrations created by drilling.
And keep in mind to change the ACSF regularly to clear the bone debris and to avoid tissue overheating. Using a dental drill, start thinning the skull surface in an area with the diameter of one millimeter using regular vertical motion parallel to the skull, and remove most of the spongy bone. Note that the bone near the sutures is highly vascularized.
If bleeding occurs, apply ACSF pre-soaked gel foam to stop it. So we keep the bone to limit inflammation, but excessive pressure or heating during the thinning procedure can also result in inflammation. Use a sharp, disposable, ophthalmic microsurgical blade to continue with the thinning of the skull to result in a final region of about 0.5 millimeters in diameter with a thickness of 20 to 35 micrometers, being careful not apply excessive pressure.
Due to bone regrowth and scarring, it is necessary to further thin the window at each imaging session. In this step, transfer the mouse with the head mount stage under the two-photon laser microscope. Using a 20X water immersion objective with a numerical aperture of 1.0, locate the thin cranial window at the center of the optical field using the epifluorescent lamp.
Ensure that the objective is always immersed in ACSF. Next, start laser scanning with a mode-locked pulse laser. Set the excitation wavelength at 750 nanometers to detect the emitted fluorescence of the methoxy-X04 in the blue channel and FITC-dextran in the green channel.
Acquire a low magnification stack at 0.7X numerical zoom in order to create a 3D map for the precise re-localization of the ROI at later time points. Then acquire a mosaic of four high-digital magnification images with a two X numerical zoom. Using the imaging software, move the window in 200-micrometer steps to capture all regions.
The depth of the stacks is typically 250 micrometers starting from the pial surface in each image of the mosaic. Before removing the mouse from the microscope, take a picture or make a hand-drawn 2D map of the pial vasculature for future re-localization of the imaging field. Shown here is the comparison of two reconstructed images of the microvasculature and amyloid depositions on a five X FAD mouse at four and five months of age.
Amyloid plaques are indicated with white asterisks, and new plaques appearing at five months are indicated by yellow asterisks. A rare instance of plaque reduction is indicated with an orange asterisk. Newly formed microvessels are indicated with yellow arrows, while vascular occlusion is indicated with a red arrow.
This figure shows that the amyloid vascular deposits on large-caliber vessels. At the same time, amyloid plaques can be also observed on small-caliber vessels. Red arrows indicate occluded vessels, whereas white asterisks indicate growing amyloid plaques.
So once mastered, this technique can be performed in 1 1/2 hours, including the surgery and imaging steps. While attempting this procedure, it is important to plan the number of imaging session in order to limit excessive thinning in the first session. Overall, these procedures allow tracking changes in cerebral vasculature during disease progression.
