The overall goal of this procedure is to generate a stable, thin skull window over the mouse cortex to allow repeated imaging without the disruption of the intracranial environment. This is accomplished by first attaching a head mount that will allow the head to be held stably under the imaging apparatus. Next, carefully thin the skull in a region over the dorsal surface of the brain and polish the surface with fine grit.
Then cover the window with a small dab of cyanoacrylate glue and a cover glass to reinforce the window. The final step is to prepare the animal for imaging under the two photon microscope. Ultimately in vivo, two photon microscopy is used to show fine structures such as arterials, vees capillaries, and neuronal dendrites, which can be imaged up to 250 micrometers deep into the cortex over weeks.
Main advantage of this cranial window technique is that the skull overlying the cortex is unreached. First, we thin the skull to obtain optical clarity, and then we reinforce the skull with cyanoacrylate glue and the cover slip. This method should be useful to study a diversity of fluorescent transgenic mice in the anesthetized or the OIC states.
To begin this procedure autoclave the required surgical tools, anesthetize a mouse at about three to six weeks old according to the methods in the accompanied manuscript. Check the reflex by toe pinching to ensure the mouse is in the surgical plane of anesthesia. Next, secure the mouse in a stereotaxic frame.
Maintain the mouse's body temperature at 37 degrees Celsius. Using a feedback regulated rectal probe and heat padd. Apply ophthalmic ointment to its eyes to retain moisture.
Then shave the scalp with the small electrical razor. After that, clean the scalp with Betadine, followed by swabbing it with 70%isopropyl alcohol. Then warm an aliquot of sterile artificial cerebral spinal fluid to 37 degrees Celsius under a microscope.
Remove the scalp over the entire dorsal skull surface with a pair of surgical scissors. Trim the skin laterally to the edges of the temporal muscles on either side of the skull and posterior to the muscles of the neck. Next, use a scalpel blade to remove the thin periosteum from the surface of the skull.
Then dry and clean the skull. After that, apply a thin layer of cyanoacrylate glue to the skull surface. Allow the glue to dry thoroughly.
This layer of glue is required for proper adherence of the dental cement. In subsequent steps. For the awake imaging preparation drill holes and introduce two number 0 0 0 self-tapping screws into the contralateral skull surface.
Then attach a rigid custom made connector with two attachment points to the skull. This greatly reduces the degrees of freedom and simplifies the relocation of the same imaging field in longitudinal studies. A wide crossbar also gives ample room for electrode placement and stimulation of vibrancy.
Next, cover the entire skull surface, excluding the location of the window with a layer of dental cement, ensure that all exposed edges of the skin are covered by cement. Before thinning the skull, ensure that drill burs are sharp and avoid reusing them. Use a dental drill at low speed to thin a two millimeter by two millimeter region over the somatosensory cortex with a one half millimeter burr.
Then thin the dry skull surface for only a few seconds at a time with the skull with a CSF periodically to prevent heating. Then thin the bone. Further switch the drill to a slower speed and shave the skull surface with only slight touches.
The drill should approach the surface at approximately a 30 degree angle from the vertical. Continue thinning the bone past the vascular layer into the second layer of bone. Above the brain surface.
Hold the drill with small controlled movements and only apply force in the lateral direction. When the bone is about 50 microns thick, the peel vessels should be clearly visible through the wet bone. In this depth, small white spots within the bone will become visible for a few seconds.
Immediately after the dry skull surface is moistened, the final bone thickness should be about 10 to 15 microns. When the bone is sufficiently thin, the small white spots in the bone will no longer be visible when the dry skull surface is moistened. Now polish the window with tin oxide powder.
Use a small pinch of powder roughly one quarter the size of the window, agitate a slurry of powder with a CSF using a pre-made drill bit with silicone whip. Polish the surface for 10 minutes, flush away the tin oxide powder from the window thoroughly and dry the bone. This process should help reduce any rough edges left by the drill bit during the thinning procedure.
Next, cut a few square pieces of number zero cover glass with the size slightly smaller than the size of the window. Use a scribe to gently score the separated horizontal and vertical lines in the cover. Slip with a straight edge without breaking the glass.
Then place the cover slip in a Petri dish. Separate the glass pieces by knocking the dish against a table edge. Afterward, apply a small dab of Sano ACRL glue over the window using the wooden tip of a broken cotton tipped applicator.
Quickly place a pre-cut piece of cover glass atop the glue. Avoid creating bubbles underneath the cover glass after the glue has been allowed to dry thoroughly over 15 minutes. Remove excess cyanoacrylate glue from the upper surface of the cover glass with a scalpel blade.
Seal the edges of the window with dental cement. Then form a slightly raised well to hold water for the dipping lens At the end. Place the animal in a warm cage until it fully recovers from anesthesia.
In this step, stabilize the animal on an optical breadboard for imaging and use the frame as a head support. Our separate plate can transport the animal and all physiological monitoring devices assembled as one unit between the surgical and two photon imaging sites. If blood flow imaging is desired, inject 0.05 milliliters of 5%fluorescent dextran dye through the infraorbital vein in order to label the blood serum.
This must be done under general anesthesia and the dye usually remains in the circulation for several hours. Inject lactated ringer solution intraperitoneal at a volume of three microliters per gram every two hours to maintain body fluids and energy requirements. Awaken the mouse by discontinuing the isoflurane.
One hour after awakening, the mouse is ready for imaging. Close the cage over the microscope to prevent light contamination.Shown. Here is an example of the cortical vasculature of a thigh, one YFP mouse labeled by intravenous injection of dextran conjugated Texas red dural vessels are often visible slightly above the cortical surface in the dura mater.
Large peel arteries and es lie on the cortical surface. Penetrating vessels branch from this surface network and dive into the cortex where they ramify into a dense capillary bed that feeds the cortical tissue. This movie shows the vasculature at the peel surface in an awake mouse.
Spontaneous oscillations in arterial diameter can be seen when the animal is in a quiet resting state. This method should be useful to study a wide variety of fluorescent transonic animals in either the anesthetize or the awake state.
