Acute Imaging Preparation for Rats: Dura-Removed Cranial Window
This preparation is primarily for imaging studies lasting 1 to 4 days and provides ample opportunity for the measurement and control of physiological parameters as well as for electrophysiological recording and drug delivery. It provides excellent optical clarity and exposes a large region of cortex, i.e., typically 3 × 6
. Further, a maximal imaging depth of 500μ
m is routinely achieved with 800
nm excitation when imaging blood vessels.
Mounting a head-frame
1. Make a 4 to 5
cm incision down the midline of the scalp using a scalpel blade. Start the incision from between the eyes and cut to just caudal of the ears. Retract the scalp to the side of the head with four hemostats, two placed on either side caudally and two placed rostrally. Control bleeding with gauze.
2. Use a periosteal elevator to remove the thin periosteum from the surface of the skull (). Control bleeding from the areas of skin and soft tissue by applying constant pressure to the bleeding tissue with a pair of hemostats and removing excess blood with gauze and cotton applicators. Both the bregma (β) and lambda (λ) points should be visible on the exposed skull surface. If necessary, make additional cuts in the scalp to widen the field.
3. Stop any bleeding from vessels emanating from the skull by using light, focused abrasion with 1/2
mm dental drill burr.
4. Separate the temporal muscle from the temporal ridge, which runs along the lateral aspects of the skull (, white arrows). Apply direct pressure to the muscle where it attaches to the skull using the periosteal elevator. Avoid separating the muscle near the eye to prevent a rip to the infraorbital vein. Rather, separate the muscle as caudally as possible past the squamosal bone, which is roughly lateral to the lambda point. If desired, retract the muscle away from the skull with hemostats on both sides of the skull. Note that one can make bilateral windows to maximize use of each animal, and to potentially study bi-hemispheric changes in blood flow ().
5. Demarcate the location of the desired cranial window (, blue circles). As an example, primary somatosensory cortex, which is the part of parietal cortex, nominally lies between −1 and −5
mm relative to the bregma point and between 1 and 7
mm from the midline on the medial-lateral axis for rats (Paxinos and Watson, 1986
). However, imaging of blood flow and/or cellular dynamics can be performed in any brain region that lies near the skull. The zygomatic arch may need to be removed when imaging more lateral structures such as auditory cortex.
6. Attach a metal connector to the skull with dental acrylic for immobilization of the head during imaging (). The connector could be a custom-fabricated metal frame (Driscoll et al, 2011a
; Kleinfeld and Denk, 1999
) or simply a nut and bolt system with a horizontal cross bar, as shown (). Clean the contact regions of soft tissue to achieve a reliable connection between acrylic and bone. Apply a thin layer of cyanoacrylate glue to the bone. Introduce small self-tapping screws to the skull to reinforce the linkage of the connector to the skull, just posterior to lambda. Care should be taken when drilling pilot holes, as the transverse sinus lies directly beneath the lambda suture line.
7. Apply dental cement to the entire assembly. Cover the entire exposed skull surface with dental cement except where the window will be placed (). Fill the space between the skull and the retracted temporal muscle. Build a slight wall around the window region, which will eventually hold the agarose and keep the overlying cover glass from touching the brain surface. The wooden handle of the cotton-tipped applicator can be snapped to make a fine tip for trailing small amounts of dental cement around the window.
Dura-removed cranial window for rat
8. Using a high-speed dental drill with a 1/2
mm burr, thin the skull evenly throughout the entire window (). Flush the surface frequently to reduce heating during drilling, and continue to thin until the underlying pial vasculature becomes visible after application of mACSF. The skull should be less than 1/4th of its original thickness at this point. This requires thinning through the vasculature of the skull, which may bleed, but can be controlled by flushing with mACSF.
9. Carefully thin the edges of the window with a 1/4
mm burr until a network of fine cracks in the bone begins to show ().
10. Soak the window with mACSF for 1
minute to allow the bone to soften, and then wick the liquid away with KimWipes.
11. Use a pair of forceps to gently separate connected segments of the bone flap from the skull, but do not yet remove the bone flap completely. Again, control bleeding from the dura by flushing with mACSF.
12. Soak the window with mACSF for another minute to soften the bone, and wick the liquid away.
13. Use forceps to grasp the far corners of the loosened bone flap, and slowly peel it away from the underlying dura mater (). Peel carefully at skull suture lines, as the underlying dura may be attached to the bone.
14. Flush the dural surface with mACSF and apply small pieces of SurgiFoam presoaked in mACSF to control bleeding. The serpentine-like dural vessels will be clearly visible at this point. All bleeding must be stopped before proceeding to the next step.
15. Retract the dura to the edges of the window (). This must be done with extreme care, because the dura is thin and close to surface of the cortex. First generate a small incision in the dura using the cutting edge of a 26-G syringe needle. Bend the needle to an obtuse angle with hemostats to ensure the cutting edge punctures the dural membrane but does not damage the underlying cortex.
16. Using two sharp no. 55 forceps, gently lift the dura away from the cortical surface, starting at the incision site, and tear in small increments (). Whenever possible, tear around large dural vessels to avoid bleeding. Limit any bleeding from dural vessels with small pieces of SurgiFoam soaked in mACSF or moistened KimWipes, twisted to a fine point with the fingers. If bleeding from the dura is excessive, flush with a slow flow of mACSF across the window until bleeding ceases.
17. Tear along three edges of the window, and roll the dural flap to the side of the window (). Flush the cortical surface with mACSF. It is crucial to avoid any damage to pial vessels. Hemorrhaging will alter cerebral blood flow, accelerate brain swelling, and severely degrade imaging quality. Removal of the dura and subsequent sealing of the window should be done swiftly, i.e., within 10
minutes, to avoid brain swelling.
18. Overlay the window with 1.5% (w/v) low-melting point agarose dissolved in mACSF (
Kleinfeld and Delaney, 1996
). Dissolve the agarose in mACSF by heating in a microwave, using short periods of heating to prevent boiling. The agarose should be prepared ahead of time and maintained in a 60°C water bath. Draw the agarose mixture into a 1 mL syringe, and allow to cool slightly before applying 3 to 4 drops to the cortex; the agarose should not feel hot on the back of the hand. Then immediately seal the chamber using a precut cover glass that is larger than the size of the craniotomy. Ensure that the cover glass is not pressing against the cortical surface, as this will impede blood flow and lead to poor imaging quality over time ().
19. Apply dental cement to the edges of the cover glass (, upper panel). Build up the cement slightly to hold water for the dipping lens. Sealing the craniotomy is crucial to protect the cortex and suppress motion from cranial pressure fluctuations because of heartbeat and breathing. However, an edge of the window can remain uncovered to allow insertion of electrodes or micropipettes (, lower panel). In this case, apply ample agarose to the exposed edge as evaporation will eventually cause the agarose to dry. The perfusion of drugs over the cortical surface can also be achieved by adding tubing and an agarose-covered ‘vent' that allows fluids to directly reach the cortical surface (Nishimura et al, 2010
). Be aware that using glass pipettes with diameters >10μ
m, such as those used to inject Ca2+
indicators, can cause cortical spreading depression and impair vascular reactivity for hours.
Preparing for imaging
20. Suture the scalp so that it fits snugly around the head mount. Excess skin around the head mount should be cut early in the procedure, as bleeding from the skin takes 10 to 15
minutes to completely stop. Compress the cut edges with hemostats for 10 to 15
seconds to help minimize bleeding. When all bleeding has stopped, protect the exposed edges of the scalp by covering with dental cement. Apply additional dental cement around the metal connector for further reinforcement (Kleinfeld and Denk, 1999
21. Stabilize the animal on an optical breadboard for imaging, using the frame as a head support (). The animal should be slightly suspended by the head mount, to prevent motion artifacts from breathing. Here, we use a custom-machined cross bar with a hole for the bolt. The bolt is inserted into the hole and a nut is then used to secure the animal's head in place. This preparation is stable for anesthetized preparations, but another bolt can be introduced to the anterior aspect of the skull for increased stability. Our animal restraint apparatus, constructed from optomechanical components, can be transported between surgical and imaging suites with the animal and all physiological monitoring devices assembled as one unit.
22. Before imaging, inject 0.3
mL of 5% (w/v) fluorescent-dextran dye dissolved in saline to label the blood serum (), either through the femoral artery/vein catheter or a tail vein. Extra fluorescein-dextran dye can be frozen in aliquots.
23. If the animal is meant to survive for more than 1 day, and is showing clear signs of pain when awakened, such as sustained immobility, decreased food and water consumption, and abnormal posture, e.g., hunched back, provide analgesia by injecting buprenorphine hydrochloride solution (10 to 50μ
g per g weight of the animal) subcutaneously. Monitor animal periodically until it fully recovers from anesthesia.
Anticipated results for rat
24.A window generated over the somatosensory cortex will reveal large proximal branches of the middle cerebral artery that can be several 100μ
m in diameter. These large arteries traverse the entire window and eventually form anastamoses with arterioles branching from the anterior cerebral artery near the midline of the brain (
Blinder et al, 2010
; Schaffer et al, 2006
25.A web of smaller pial arterioles, typically <100μ
m in diameter, link these major branches and form redundant vascular loops over the cortical surface. In a 3 × 6
cranial window one might see ~100 penetrating arterioles that dive to feed subsurface microvessels (Blinder et al, 2010
26.Below the surface, penetrating arterioles begin to ramify into fine precapillary arterioles as shallow as 10μ
m below the cortical surface, and continue to branch off over the depth of cortex (Tsai et al, 2009
) (). Larger penetrating arterioles that feed deeper structures tend to branch less within the imaging depth accessible by TPLSM.
27.Capillary networks drain into penetrating venules and, like penetrating arterioles, run perpendicular to the cortical surface. Penetrating venules typically outnumber penetrating arterioles by a factor of 1.8 × in rat (Nguyen et al, 2011
28.Penetrating venules drain into pial surface venules and, like arterioles form an intricate network on the surface. These venules either drain medially toward the superior sagittal sinus, or laterally toward the rhinal vein (Scremin, 1995
). Venules can be differentiated from arterioles during imaging by a number of attributes, including: (i) a mottled appearance caused by slower moving RBCs; (ii) greater abundance of branches and cortical penetrations; (iii) a generally larger lumen diameter; (iv) a slower flow for the same lumen diameter; and (v) the direction of flow, as penetrating arterioles support flow into the brain while penetrating venules support outward flow. As arteriovenous anastomoses are not present in the normal brain, the individual arterial and venous networks can be traced without ambiguity once a vein or artery is identified.
29.For troubleshooting, refer to .
Troubleshooting for rat cranial window and imaging
Chronic Imaging Preparation for Mice: Polished and Reinforced Thinned-Skull Window
This procedure allows repeated imaging for up to 3 months over a large cortical region, i.e., 2 × 2
, without disrupting the intracranial milieu. Imaging depths of up to 250μ
m are routinely achieved. It also permits the introduction of an electrode or micropipette to the brain through a separate drill hole.
Mounting a head frame
1. Remove the scalp over the entire dorsal skull surface (). Use a scalpel blade to remove the thin periosteum from the surface of the skull.
2. Clean and dry the skull surface. Apply a thin layer of cyanoacrylate glue to the surface and allow the glue to dry thoroughly.
3. Attach a metal connector to the skull, away from the area of the desired window with a small dab of cyanoacrylate glue and allow the glue to dry thoroughly. Then secure the connector with a layer of dental cement. We adhere a small nut (no. 2-56) that can be later secured to the imaging setup using a bolt (). Seal the backside of the nut with tape to ensure that glue does not enter the threads.
4. Alternatively, attach a custom-made connector, in this case with two attachment points (). This greatly reduces the degrees of freedom and simplifies relocation of the same imaging field in longitudinal studies. A wide crossbar gives ample room for electrode placement and stimulation of vibrissae.
5. Cover the rest of the 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.
6. Stabilize the mouse in a head mount (). As with the restraint apparatus for rats describe above, a mouse apparatus can be made from commercially available miniature optomechanical components from Qioptiq (Rochester, NY, USA) or ThorLabs (Newton, NJ, USA).
Generating a polished and reinforced thinned-skull (PoRTS) window
7. Thin a 2 × 2-mm2
region over the somatosensory cortex with a 1/2-mm burr. Alternate between wetting the skull with mACSF and then drying the skull surface with a gentle stream of air; wet for cooling, and dry for thinning. The skull may bleed from the vessels in the inner cancellous layer, but can be controlled by flushing with mACSF (). The skull begins to flex without breaking under the slight pressure of the drill when it is ~50μ
m thick, and the pial vessels should begin to be visible through the wet bone.
8. At this point, the bone must be thinned even further, and the speed and sharpness of the drill burr are critical. Change to a new drill burr and lightly shave the skull surface with small controlled movements while holding the drill like a pen. We find that a drill speed of ~1,000
r.p.m. is appropriate at this stage. Small white spots within the bone, normally visible when moistened bone is ~50μ
m thick, will not be visible at the final skull thickness of approximately 10 to 15μ
9. Polish the window region with tin oxide powder (). Attach a premade drill bit that has been dipped in silicone aquarium sealant and withdrawn, leaving a tapered whip (inset, ). Place a small pinch of powder on the window along with a drop of mACSF. Agitate the slurry over the window for up to 10
minutes by gently touching the tip of the moving whip to the skull surface. Surface irregularities and adherent bone chips left by drilling in the previous steps should be removed after polishing. Flush away the tin oxide powder thoroughly from the window using mACSF and dry the bone thoroughly with a gentle stream of air.
10. Cut small square pieces of no. 0 cover glass, roughly 2 × 2
, by gently scoring separated horizontal and vertical lines in the cover glass with a scribe. Then place the cover glass in a Petri dish and shake vigorously to separate the glass pieces.
11. Apply a small dab of cyanoacrylate glue over the window using the wooden tip of a broken cotton-tipped applicator and quickly place an appropriately sized piece of no. 0 cover glass atop the glue (). Using forceps, push the glass so that it is in contact with the skull surface. Avoid creating bubbles underneath the cover glass. Allow the glue to dry thoroughly for 10
minutes before proceeding. Excess cyanoacrylate glue can be removed from the upper surface of the cover glass with a scalpel after it is dried.
12. Seal the edges of the cover glass with dental cement and form a slightly raised well to hold water for the dipping lens ().
13. For some experiments, it may be desirable to inject dyes or insert electrodes into the tissue volume beneath the PoRTS window. A small hole can be added adjacent to the window, through which pipettes or electrodes can be introduced using a stereotaxic arm or Sutter manipulator (Stosiek et al, 2003
). This hole can be resealed with bone wax after the experiment if the animal is to be imaged again in future sessions.
Preparing for imaging
14. Stabilize the animal on an optical breadboard for imaging, using the frame as a head support. Our separate plate can be transported between surgical and imaging suites with the animal and all physiological monitoring devices assembled as one unit ().
15. Inject 0.05
mL of 5% (w/v) fluorescent-dextran dye dissolved in saline either through a femoral artery/vein catheter, tail vein, or infraorbital vein to label the blood serum (). For tail vein or infraorbital injections, use an ultrafine 0.3-mL insulin syringe with a 29.5-G needle.
16. If the animal is meant to survive for more than 1 day, and is showing clear signs of pain (see step 23 of last section), provide buprenorphine hydrochloride solution subcutaneously for analgesia.
Additional notes for awake imaging preparations
17. More rigid head mounts are necessary for imaging awake animals. For increased stability, two self-tapping #000 screws can be added to the contralateral hemisphere of the skull before application of the dental cement, as shown in step 3 ().
18. Habituation to head fixation is important to reduced animal movement during imaging. A new animal can be gradually accustomed to head restraint over a period of 3 to 7 days, starting with 15
minutes sessions and working up to several hours (Drew et al, 2011
). Before blood flow imaging, the mouse can be briefly anesthetized with isoflurane for an infraorbital injection of fluorescent-dextran dye. The dye will remain in circulation for several hours, and supplements can be given as necessary if the animal is re-anesthetized.
Anticipated results for mouse
19. The cortical vasculature of the mouse is very similar to that of a rat, and can be thought of as a cropped, rather than scaled, version of the rat pial network (Blinder et al, 2010
). In our example, we imaged a portion of the pial network after an intravenous injection of Texas red-dextran (), concurrent with yellow fluorescent protein (YFP)-labeled dendrites in the Thy1-YFP mouse line () (Feng et al, 2000
; Zhang et al, 2005
). The average density of penetrating arterioles, and average surface area occupied by arterioles loops is very similar to the rat, but the total surface area of the mouse cortex is three times smaller (Blinder et al, 2010
). As with rat, the subsurface microvasculature is highly tortuous and forms densely packed loops.
20. Detailed studies from mouse have revealed that neurons lie, on average, 15μ
m away from the nearest microvessel, despite the fact that that cerebral vasculature only accounts for between 1% and 3% of the total cerebral volume (Tsai et al, 2009
21. Two-photon imaging the vasculature through a PoRTS window requires transmission though the thinned bone and the dura, which attenuates the laser light and adds optical aberrations at greater depths (Drew et al, 2010b
). However, despite this drawback, a well-prepared window will allow imaging of depths up to 250μ
m below the pial surface for months (), and should show no sign of damage, such as angiogenesis and microglial reactivity.
22. In our hands, the success rate of the PoRTS window can be as high as 80%. In failed cases, the skull is breached or subarachnoid hemorrhaging occurs. These cases should be discarded.
23. The age of the mouse has a strong influence on the success of bone thinning. In adult mice 4 weeks or older, the rigidity of the more developed skull appears to prevent the bone from breaking, which is necessary to achieve the 10 to 15μ
m optimal thickness. In younger mice, the soft skull is prone to break during the thinning process. Bleeding from the cancellous layer of the skull is also more prominent in older animals.
24. For troubleshooting, refer to .
Troubleshooting for mouse PoRTS window and imaging