All animal experiments were performed in accordance with guidelines approved by the IACUC at the University of Nebraska Medical Center.

1. Before surgery

1. Prepare pipettes for viral injections. Pull borosilicate glass capillaries using a pipette puller and bevel the pipette at a 20° angle. Sterilize the pipettes overnight.
2. Prepare fresh pieces of sterile gel foam by cutting them into small squares. Submerge the gel foam in a sterile microfuge tube containing 0.5 mL of saline. Soak the gel foam in saline for at least 30 min before use.
3. Prepare fresh pieces of sponge strips by cutting them into thin strips.
4. Twenty minutes before surgery, inject 4-6-week-old GLAST-CreER/GCaMP6f mice intraperitoneally with 0.2 mg/kg dexamethasone to prevent brain swelling and 5 mg/kg carprofen to reduce inflammation.
   ​NOTE: To induce the recombination and expression of GCaMP6f in approximately 40% of the astrocytes, GLAST-CreER/GCaMP6f mice were injected with 100 mg/kg tamoxifen at 3 weeks for 5 consecutive days.

2. Start of surgery

1. After 20 min, anesthetize the mice with 3% isoflurane with oxygen at a flow rate of 1 L/min for approximately 1.5 min.
2. Once anesthetized, place the mouse on a stereotaxic frame that sits on a water re-circulating blanket to maintain a body temperature of 37 °C throughout the surgery. Secure the mouse to the frame using the snout clamp and ear bars. Ensure that the snout clamp and ear bars are stable enough that the head will not move during the procedure, but not so tight that the skull is damaged.
3. Maintain anesthesia with 1-1.5% isoflurane with oxygen at a flow rate of 1-2 L/min. Note that some animals may need more or less isoflurane to maintain sedation. Monitor the breathing of the animal as well as its reflexes to toe and tail pinches, and adjust the isoflurane appropriately.
4. Apply eye ointment to each eye using a cotton tip applicator to prevent the eyes from drying out during the procedure.
5. Using rodent trimmers, shave the hair from the neck to just past the eyes. Use caution to ensure that the whiskers of the animal are not accidentally trimmed.
6. Clean the shaved area with one iodine prep pad, followed by one alcohol prep pad.
7. Using sterile tissue forceps and surgical scissors, cut and remove the skin from the frontal sutures anterior to the bregma all the way posteriorly to lambda.
8. Once the skin is removed, add approximately 0.1 mL of 1% xylocaine (lidocaine with epinephrine 1:200,000) to the skull to minimize bleeding of the skull.
9. Using a sterile size 11 carbon steel surgical blade attached to a handle, gently scrape the connective tissue from the skull. Take extra care when removing the connective tissue near the edge of the skull, as any remaining tissue could prevent strong adhesion of the glass or head plate later on.
10. Use sterile tissue forceps to remove loose connective tissue from the skull.
11. Using sterile cotton tip applicators, clean off any remaining xylocaine from the skull.
12. Once completed, thoroughly degrease the skull using a sterile cotton tip applicator dipped in acetone. Use compressed air to immediately blow-dry the skull.

3. Craniotomy

1. Using a fine-point marker and a ruler, measure the appropriate size for the opening at the desired location. Confirm the size of the opening by comparing it to the size of the cover glass (3 or 5 mm in diameter); ensure that the opening is slightly smaller than the size of the cover glass.
2. Using a dental drill, gradually trace the outline of the opening, thinning the bone. Drill slowly to prevent drilling through the bone, and drill in concentric circles to facilitate even thinning of the bone.
3. After each concentric pass, add a drop or two of sterile saline to the drilled area. Allow the saline to sit for at least 10 s to prevent the bone from overheating and damaging the underlying dura.
4. Use a sterile cotton tip applicator to absorb any remaining saline. Blow compressed air over the area to ensure that all remaining saline has evaporated before resuming drilling. Do this to blow away bone debris that remains.
5. Repeat steps 3.2-3.4 until the bone is adequately thinned for removal.
6. Ensure that the bone is adequately thinned for removal by gently pushing on the central area of the bone with fine forceps to check that the thinned skull moves. The underlying vasculature should be visible where the drilling occurred. Observe that the bone may appear as if it will crack where thinning occurred.
   NOTE: Training at this stage is crucial to identify when the bone has been thinned appropriately.
7. Before attempting to remove the bone, check the mouse to make sure it is completely sedated. If not, gradually increase the isoflurane maintenance to prevent brain swelling when removing the bone.
8. Add a small piece of gel foam saturated in saline to the thinned skull. Add one or two additional drops of saline to the thinned skull.
   NOTE: Having plenty of saline over the thinned skull helps to reduce bleeding and protect the dura when removing the bone flap.
9. Using a miniature 15° pointed blade, carefully insert the blade into the thinned bone and cut along the thinned bone. Keep the gel foam soaked in saline, ready to stop any bleeds that may occur.
10. Using forceps, carefully lift and remove the bone. Be as gentle as possible to avoid damage to the underlying tissue and vasculature. Take extreme care not to damage the dura mater.
11. Once the bone flap is removed, add a fresh piece of gel foam saturated in saline to the cortex. Add a few drops of saline to prevent the gel foam from drying out, as this will cause damage to the underlying tissue.

4. Viral injections

1. Perform viral injections using a stereotaxic apparatus.
   1. Prepare AAV1.CaMKII.0.4.Cre (titer of 3 × 10¹³ genome copies (GC)/mL) at 1:5,000 by serial dilutions in saline.
   2. Prepare the mixture of AAV1.CaMKII.0.4.Cre (1:5,000) and AAV1.CAG.FLEX.tdTomato (titer of 5 × 10¹² GC/mL) on a small piece of parafilm.
   3. Fill a sterile beveled glass pipette with a tip size of 20 µm with the virus mixture by gently placing the pipette on the solution.
   4. Lower the pipette so that it just touches the surface of the brain and continue to lower for an additional 200-300 µm for L2/3 injections. Using an intracellular microinjection dispense system (see the Table of Materials), pressure-inject 12-15 times over 2 min (20 psi, 9 ms pulse duration). Observe the meniscus in the pipette drop with each injection to ensure that the pipette is not blocked.
   5. Once injected, leave the pipette in the brain for 4-5 min to prevent backflow. Slowly retract the pipette.
   6. Repeat the injections at 2-3 sites (approximately 500 µm apart).
   7. Discard the used glass pipettes, parafilm, pipette tips, and microfuge tubes used for serial dilutions of the Cre virus into 10% bleach.

5. Implantation of cranial window

1. Remove any gel foam, and use a small strip of sponge strips to soak up any remaining saline.
2. Add a few drops of the antibiotic enrofloxacin (22.7 µg/mL) to the opening and allow it to remain there for 1 min. After 1 min, use a fresh piece of sponge strip to absorb any remaining enrofloxacin. Repeat this process two more times.
3. After the third wash, add a few drops of saline to the opening and allow it to remain there for 1 min. After 1 min, use a fresh piece of sponge strip to absorb any remaining saline. Repeat this wash process two more times and after the third wash, add a small drop of saline to the opening.
4. Place the cover glass over the opening. Use forceps to ensure that the glass is flush over the opening to obtain images of good quality.
5. While gently holding the glass in place, apply cyanoacrylate adhesive gel (Table of Materials) around the perimeter of the glass to seal the edges of the window to the skull. Make sure not to let the glue seep under the cover glass, and keep as much glue off the surface of the cover glass as possible.
6. Apply a layer of super glue over the adhesive gel. Add a layer of dental cement liquid over the glue to allow it to harden.
7. Take an appropriately sized helicopter-type head plate and apply a thin layer of glue around the central opening. Place the head plate over the cover glass, and allow the glue to dry briefly.
8. In a 1.5 mL microfuge tube, add dental cement powder to the 0.1 mL mark on the tube. Add 7-8 drops of fast-curing, instant adhesive, and mix. Draw into a 1 mL syringe with a 19 G needle that has been cut to create a larger opening.
9. Inject the mixture through the lateral holes of the helicopter bar until it seeps from either side. Apply the dental cement/adhesive mixture to the rest of the exposed skull to fasten the headplate to the skull, which will reduce movement artifacts during imaging.
10. Allow the mixture to air-dry for at least 15 min.
11. Inject the mouse with 0.5 mL of saline subcutaneously to aid in recovery.

6. Post operation

1. Remove the mouse from the stereotaxic frame and return it to its cage.
2. Place a portion of the cage on a water re-circulating blanket, space gel heating pads or isothermal pads to assist with recovery.
3. Provide the animal with a small helping of food pellets and wet food, in addition to their regular diet, to further aid in recovery. Add new food pellet and wet food each day after surgery for the duration of recovery.
4. The day after surgery, inject mice once with 5 mg/kg carprofen and 5 mg/kg enrofloxacin.
5. On days 2-6, inject the mice once with 5 mg/kg carprofen and twice with 5 mg/kg enrofloxacin, pending approval by the local IACUC. Separate the enrofloxacin injections by at least 8 h.
6. On days 7-20, inject the mice daily with 5 mg/kg carprofen.

7. Animal habituation for imaging

1. On day 14 after surgery, examine the cranial window for optical clarity. Do not use mice with unclear or otherwise damaged windows (i.e., excessive angiogenesis).
2. Check for tdTomato expression under a fluorescence microscope. If labeled cells can be identified, proceed with animal habituation.
3. First day of habituation (handling)
   1. Hold the mouse for a few minutes and return it to its cage after handling. Repeat the handling three times with a 15 min interval between the sessions.
   2. After the third trial, habituate the mouse by wrapping the animal in a small piece of cloth.
   3. Hold the mouse wrapped in cloth for approximately 1 min.
   4. After the trial, return the mouse to its cage. Repeat this process two more times, allowing a 15 min interval between each trial.
4. Second day of habituation
   1. Weigh and record the mouse weight.
   2. Wrap the mouse in cloth, and secure it via its head plate to the head fixation arm of the air-lifted home cage.
   3. Leave the home cage exposed to the light.
   4. Allow the mouse to remain secured in the mobile home cage for 15 min.
   5. After habituation, remove the mouse from the home cage and turn off the airflow.
   6. Weigh the mouse and return it to its cage. Take care to only include mice that do not lose more than 10% of their body weight. Exclude mice that lose more than 10% of their weight during any day of habituation from imaging experiments.
5. Day three of habituation and beyond
   1. Weigh and record the mouse weight before securing it to the air-lifted home cage.
   2. Secure the mouse to the air-lifted home cage.
   3. Cover the home cage with something that will provide a dark interior, such as a box, and allow the mouse to remain secured for 30 min.
   4. After 30 min, uncover the home cage, remove the mouse, and turn off the air flow.
   5. Weigh and record the mouse weight after habituation.
   6. Repeat this process every day, increasing the duration the mouse is secured to the home cage by 15 min. Continue this process until the mouse can be secured to the home cage for 1.5 h as this is the approximate duration of a two-photon imaging session.
   7. To habituate the mouse to noise, perform some habituation sessions on the microscope to acclimate the mouse to the sounds of the laser scanning mirrors. Exclude mice that fail to habituate sufficiently (i.e., vocalizations and stress-induced defecation).

8. Multiphoton imaging

1. Commence imaging 3 weeks following surgery to allow for the window to clear.
2. Perform imaging on a custom-made or commercially available multiphoton microscope equipped with a Ti:Sapphire laser. Acquire images using a high numerical aperture (NA) water immersion objective.
   NOTE: Two-channel imaging is achieved by using a 565 nm dichroic mirror and two external photomultiplier tubes. A 535/50 bandpass filter is used to detect GCaMP6f emission, and a 610/75 bandpass filter is used to detect tdTomato. A 25x water immersion objective (1.05 NA) and resonant scanners were used to acquire images shown in Figure 1 and Figure 2. Each region of interest consisted of a stack of images separated axially by 1 µm. Each optical section was collected at 512 x 512 pixels, 0.18 µm/pixel. Images were acquired from the forelimb region of the primary motor cortex as determined by stereotaxic coordinates.
3. Set the wavelength of the laser to 920 nm (990 nm if a red-shifted GECI is used).
4. Place the mouse on the mobile home cage, and clamp the headplate to the head fixation arm.
5. Add a few drops of water to the center of the window.
6. Using the wide-field mode of the microscope, select 2-3 positions of easily identifiable vasculature. Save the images of these blood vessels and record the X- and Y-coordinates that appear on the motor controller to relocate the tdTomato-labeled dendrites for subsequent imaging sessions. Adjust the X- and Y-coordinates each time to realign with the saved images of the blood vessels.
7. Once the vasculature has been imaged and positions recorded, locate the regions with neurons expressing tdTomato (Figure 1). Image the synaptic structures (dendrites and axons) of neurons and GCaMP6f activity in astrocytes.
   NOTE: The dendrites will serve as landmarks to reliably return to the same regions and image the same dendrites and astrocytes over repeated days. No detectable shift in the imaging plane was observed with stable head fixation and after habituation to head fixation. Displacement that occurs in the X and Y directions is motion-corrected.
8. After imaging, return the mouse to its cage.
   NOTE: Mice are typically kept on the scope for a maximum of 2 h. When the imaging lasts for a long time, the water under the objective may dry. Water should be added during the experiments. Alternatively, an ultrasonic gel can be used instead of water.