Intravital Imaging of Fluorescent Protein Expression in Mice with a Closed-Skull Traumatic Brain Injury and Cranial Window Using a Two-Photon Microscope

All the animal related protocols were conducted in accordance with the Guide for the Care and Use of Laboratory Animals published by the National Research Council (US) Committee. The protocols were approved by the Institutional Animal Care and Use Committee of University of Massachusetts Chan Medical School (UMMS) (Permit Number 202100057). In brief, as shown in the schematic of study (Figure 1), the animal receives a virus injection, a TBI, a window implantation, and then intravital imaging in a time sequence.

NOTE: Commercial terms have been removed. Please refer to the Table of Materials for the specific equipment used.

1. Intracranial injection of AAV using a stereotaxic device

1. AAV(PHP.eB)-Syn1-EGFP
   1. Use a viral titer of 1 x 10¹³ viral genomes per milliliter (vg/mL). Syn1 refers to Synapsin1, which is a neuronal specific promotor allowing for restricted viral expression in neurons. The virus can be prepared in-house or outsourced.
2. Preparation for stereotaxic injection surgery for administration of AAV
   1. Autoclave regular scissors, a surgical caliper, surgical spring scissors, surgical forceps, mosquito forceps, gauze, a cotton tipped applicator, and a glass microliter syringe.
   2. Sanitize the surgery area using 75% ethanol. Expand the disposable sterile surgical drape to cover the surgery region on the stereotaxic platform.
      NOTE: To maintain animal body temperature during the injection surgery, use a feedback regulated heating apparatus.
   3. Weigh and record the mouse body weight.
   4. Open the oxygen tank valve and adjust the oxygen flow to 1.5 L/min. Open the anesthesia machine and set the isoflurane level value to three.
      NOTE: The carrier gas can be either room air or 100% oxygen in this step.
   5. Put the mouse into the induction chamber and let it stay for 5 min to achieve full anesthesia.
   6. Once the mouse is fully anesthetized (i.e., slow but steady breathing rates at approximately one cycle per 2 s, coupled with the absence of a tail-pinch reflex), remove the mouse from the induction chamber and place the mouse head in the stereotaxic frame.
   7. Place the anesthesia tubing nose cone over the mouse's snout.
   8. Maintain anesthesia using 1.5% isoflurane, with carrier gas at a 1 L/min flow rate until the end of surgery. Periodically check the breath frequency, and deliver a tail or toe pinch at least every 15 min throughout surgery to assess appropriate anesthesia. Adjust the isoflurane concentration as appropriate to maintain the desired depth of anesthesia.
   9. Fill a microliter syringe (10 µL) with the virus solution. Then, fix the syringe onto the matched microinjector pump of the stereotaxic device.
3. Injection surgery
   1. Administer buprenorphine (1 mg/kg, subcutaneously) using a disposable insulin syringe and apply lubricant ophthalmic ointment to the mouse's eyes.
   2. Remove the hair from the scalp on top of the head by trimming with regular scissors 1.
   3. Clean the scalp skin with gauze and cotton tipped applicators. Disinfect the skin using 75% ethanol first and then betadine. Repeat this three times (i.e., ethanol followed by betadine), leaving the final application of betadine on the skin.
   4. Re-check the anesthetic depth to ensure that the mouse is fully anesthetized (i.e., slow but steady breathing rates at approximately one cycle per 2 s, coupled with the absence of a tail-pinch reflex). Make a ~1 cm incision using regular scissors 2 along the midline to expose the right parietal skull, and remove the periosteum using a cotton tipped applicator.
   5. Mark two points on the skull using a marker pen at these coordinates: point A: 2.5 mm posterior to the Bregma and 1 mm lateral to the midline over the right hemisphere; point B: 2.5 mm posterior to the Bregma and 2 mm lateral to the midline over the right hemisphere.
   6. Carefully drill two holes through the skull at the marked coordinates using an electric dental drill with a fine EF4 carbide bit.
      NOTE: Be careful not to damage the brain tissue.
   7. Adjust the stereotaxic frame to align the microliter syringe needle tip to the skull hole that is 1 mm lateral to the midline.
   8. Lower the syringe needle to touch the brain surface, and then set that location as the zero point (for the z-axis). Lower the needle tip into the brain cortex to a depth of 0.5 mm, and slowly infuse 1 µL of the virus solution at a speed of 200 nL/min.
   9. Wait 5 min after the virus solution is completely injected into the brain tissue before withdrawing the needle to prevent backflow of the solution.
   10. Slowly withdraw the syringe needle. Repeat the injection at the other site. Suture the incision with a sterile, non-absorbable surgical suture (6-0 gauge).
   11. Administer cefazolin [500 mg/kg (333 mg/mL, usually ~45 µL), intramuscularly], and meloxicam (5 mg/kg, subcutaneously) after surgery while the animal is still anesthetized.
   12. Release the mouse from the stereotaxic frame and discontinue anesthesia. Place the mouse in a clean cage above a heating blanket, and monitor the animal until it is ambulatory (approximately 15 min). Then, transfer to the home cage.
   13. Administer buprenorphine (1 mg/kg, subcutaneously) and meloxicam (5 mg/kg, subcutaneously) again at 8 h, 16 h, and 24 h, respectively, after the first buprenorphine injection.
       ​NOTE: At 2-4 weeks after virus injection, the mouse receives TBI and cranial window implantation at the same site as the AAV injection.

2. Administration of a repetitive TBI induction

NOTE: The TBI parameters are adjusted from previous reports^7,8, in which the TBI impact was delivered once. The protocol here applies the same parameter, except increasing the total impact number to 10.

1. TBI equipment
   1. Use a custom-built portable device for the administration of a closed-skull TBI (Figure 2A) to the mouse head on the right-side, as indicated in Figure 2B.
2. Preparation before surgery
   1. Autoclave surgery equipment, including regular scissors, a surgical caliper, surgical spring scissors, surgical forceps, mosquito forceps, headpost pieces, gauze, and cotton-tipped applicators.
   2. Weigh and record the mouse's body weight 2 h before surgery. To minimize brain edema during cranial window implantation, inject dexamethasone sodium phosphate at a dose of 4.8 mg/kg [2 mg/mL, usually ~70 µl (need to inject at two sites)] into the quadriceps muscle using an insulin syringe. After the injection of dexamethasone, but prior to starting TBI surgery, prepare the glass windows as indicated in step 3.1 for later use.
   3. Disinfect the surgical stage and TBI equipment using 75% ethanol, and expand the disposable sterile surgical drape to cover the surgery stage on the stereotaxic platform. Turn on the heating apparatus and the temperature monitor, setting the target temperature to 37 °C.
   4. Open the oxygen tank valve and adjust the oxygen flow to 1.5 L/min. Open the anesthesia machine and set the isoflurane level value to three.
      ​NOTE: 100% pure oxygen can help the animal to survive the TBI impacts.
   5. Put the mouse into the induction chamber and let it stay for 5 min to achieve full anesthesia.
   6. Once the mouse is fully anesthetized (i.e., slow but steady breathing rates at approximately one cycle per 2 s, coupled with the absence of a tail-pinch reflex), remove the mouse from the induction chamber and place the mouse head in the stereotaxic frame.
   7. Place the anesthesia tubing nose cone over the mouse's snout.
   8. Maintain anesthesia using 1.5% isoflurane mixed with 100% pure oxygen at a 1 L/min flow rate until the end of surgery. Periodically check the breath frequency, and deliver a tail or toe pinch at least every 15 min throughout surgery to assess appropriate anesthesia. Adjust isoflurane concentration as appropriate to maintain the desired depth of anesthesia.
3. TBI surgery
   1. Administer buprenorphine (1 mg/kg, subcutaneously) using disposable insulin syringes and apply ophthalmic ointment to the mouse's eyes.
   2. Remove the hair from the top of the head by trimming with scissors 1, and then applying depilatory agent for 1 min.
      ​CAUTION: Do not apply the depilatory agent product for more than 3 min on the scalp, as it is a skin irritant. Avoid getting any depilatory agent onto the mouse's eyes.
   3. Clean the scalp skin by applying gauze and cotton-tipped applicators. Then, disinfect the skin, using 75% ethanol first and then betadine. Repeat this three times (i.e., ethanol followed by betadine), leaving the final application of betadine on the skin.
   4. Re-check the anesthetic depth to ensure that the mouse is fully anesthetized (i.e., slow but steady breathing rates at approximately one cycle per 2 s, coupled with the absence of a tail-pinch reflex). Tent the skin with surgical forceps 3, and, make a 12-15 mm long midline incision starting approximately 3 mm posterior from the eyes.
   5. Excise the skin over the left and right hemisphere of the skull using spring scissors 2.
   6. Once the skull is exposed, remove the periosteum by gently rubbing with a sterile cotton-tipped applicator and flushing with sterile saline.
   7. Visually inspect the condition of the skull to ensure that it is intact, except for the two small holes, which were made for the previous virus injection surgery.
   8. Dry the skull area and mark the TBI impact site at the following coordinates: 2.5 mm posterior to the Bregma and 2 mm lateral from the sagittal suture to the right (Figure 2B).
   9. Quickly remove the mouse from the stereotaxic frame and place the head on the buffer cushion under the TBI device.
   10. Align the impactor tip with the marked impact site.
   11. Lift the metal column by pulling the tethered nylon string to 15 cm above the mouse head and then release it, allowing the weight to fall freely onto the transducer rod, which is in contact with the skull-top at the TBI site. Do not touch the mouse head when delivering the TBI impact.
   12. Move the mouse onto the heating blanket, and place it on its back while monitoring the breathing status.
   13. Once the mouse rights itself from a supine to a prone position, place the mouse into the isoflurane induction chamber for ~5 min with 3% isoflurane mixed with 100% pure oxygen at a 1.5 L/min flow rate.
   14. Once the mouse is fully sedated (i.e., slow but steady breathing rates at approximately one cycle per 2 s, coupled with the absence of a tail-pinch reflex), repeat steps 2.3.9-2.3.14 to achieve a total of 10 impacts.
       NOTE: Ten TBI impacts have been found to induce robust phenotype with low mouse mortality in our study (data not published yet). The parameters may be adjusted to achieve a different severity of brain injury by increasing or decreasing the number of impacts and/or the height from which the weight is released. All parameters are subject to approval by local IACUC.
   15. Check the skull under a surgical microscope, and remove the mouse from the study if a skull fracture has occurred.
   16. For a sham surgery, follow the same procedures as described above, including placement of the animal under the impactor, but without delivering the TBI impacts.
   17. After the mouse rights itself from a supine to a prone position following the 10^th TBI impact, place the mouse into the isoflurane induction chamber for ~5 min. Follow steps 2.2.6-2.2.8 to place the mouse head onto the stereotaxic frame for the cranial window implantation surgery described below.

3. Cranial window implantation surgery

NOTE: The cranial window implantation steps below were adopted from Goldey et al.⁹, and their specifications of the headpost and the imaging well were applied here.

1. Window preparation
   NOTE: Complete the window preparation in step 2.2.2 before starting the TBI surgery. The window is made from two round glass coverslips (one 3 mm and one 5 mm in diameter, as indicated by Figure 2C) that are joined together by transparent optical adhesive, as described below.
   1. Sanitize the glass coverslips by submerging them in 75% ethanol for 15 min. Take the glass coverslips out of the ethanol and leave them to dry on a sterile surface (i.e., the lid of a sterile 24-well plate) for ~10 min.
   2. Fill an insulin syringe with transparent optical glue while avoiding bubble formation. Under microscope 1 (Table of Materials) at 0.67x magnification, put a small drop (~1 µL) of optical glue at the center of the 5 mm coverslip. Then, immediately place the 3 mm coverslip on top, and center it with the 5 mm coverslip.
   3. Gently apply pressure using fine forceps to spread the glue evenly. Discard the glass window if a bubble or wall forms around the 3 mm cover slip.
   4. To cure the glue, place the coverslips into a UV box for 150 s at a power of 20 x 100 µJ/cm². Check that the two coverslips are securely bonded, by using forceps 4 to gently nudge the side of the 3 mm slip. If the coverslip moves, place it back in the UV box for an additional 60 s. Store the glass window in a sterile 24-well plate for later use.
2. Rough the skull surface.
   NOTE: From here on, carry out all steps in section 3 under a surgical microscope. Start with 10x, and adjust to the desired magnification.
   1. Gently and slowly drill the surface of the skull using a FG4 carbide bit at a low rotor speed (i.e., output number set to ~1-2) to remove the remaining periosteum and create a rough skull surface, such that the dental cement binds securely with the skull. A scalpel can also be used here instead of a drill as an alternative.
   2. Use saline to flush and clear bone dust from the skull surface.
3. Separate the muscles.
   NOTE: Muscle separation serves to increase the surface area of the exposed skull bone that will serve as a contact point for the dental cement, thereby ensuring structural integrity of the implant. This muscle separation step usually exposes ~3 mm of the temporal skull plate.
   1. At approximately 5 mm posterior to the eye, where the suture connecting the parietal and temporal skull is located, gently insert the closed fine tips (#5/45 forceps) to separate the lateral muscles from the skull, and gently move the closed tips in the posterior direction until the Lambdoid suture.
   2. Separate the lateral muscles on the side where implantation of the cranial window will occur.
      NOTE: Be careful not to separate the muscles too close to the eye, to avoid injuring the ophthalmic artery; otherwise, severe and persistent bleeding may occur. Do not separate muscles from the occipital bone, as the mouse requires these muscles to raise its head.
   3. Wash away the debris from the surgical site using saline and dry the area with gauze. Gel foam can be applied to the surgical site to stop the bleeding.
4. Implant the headpost.
   1. Use a custom-made titanium headpost (Figure 2D), described in a previous publication⁹, to fix the mouse head securely while performing the craniotomy surgery, and for subsequent two-photon imaging.
   2. Use a marker pen and a surgical caliper to trace the circumference of the craniotomy on the clean and dry skull on the right side. The center point of the traced circle is 2.5 mm posterior to the Bregma and 1.5 mm off the sagittal suture on the right hemisphere. The diameter of the traced circle is about 3.2-3.5 mm, slightly larger than the 3 mm glass coverslip. Ensure the craniotomy circle can cover the virus injection sites (the two holes made while injecting the virus can be used as a reference) and the TBI site.
   3. Loosen the ear bar and rotate the head so that the craniotomy plane is perfectly horizontal, and then tighten the ear bar again.
   4. Use the wood stick of a cotton tipped applicator to add two small drops of superglue to the front and back edge of the headpost.
   5. Position the titanium headpost over the center of the craniotomy, and quickly adjust it to rest within the same plane as where the cranial window will be implanted. Apply light pressure until the superglue is dried; this usually takes ~30 s.
   6. Prepare dental cement in a pre-cooled ceramic mixing dish (staying at least 10 min in a -20 °C freezer): combine 300 mg of cement powder, six drops of quick base liquid, and one drop of catalyst, and then stir the mixture until thoroughly mixed (~15 times).
      NOTE: The cement needs to be pasty. If it is too thin, stir in a little more cement powder. If it is too thick, stir in quick base liquid one drop at a time until a pasty consistency is achieved.
   7. Quickly apply a generous amount of dental cement mixture to the outside perimeter of the traced circumference, and cover any exposed bone surface. However, do not cover the site of the craniotomy. Allow ~15 min for the dental cement to dry and harden before proceeding.
   8. Release the ear bar and secure the headpost to the metal frame to ensure that the head is stable for precise drilling along the marked craniotomy circumference. If there is cement over the craniotomy site, use the FG4 carbide bit to drill and remove it.
5. Craniotomy
   1. Use a surgical caliper to verify the diameter of the marked circle, as defined in step 3.4.2. Adjust as necessary, so that the cranial window will fit snugly inside the craniotomy.
   2. Using an electric dental drill, etch and thin the skull along the outside of the marked circle, using a FG4 carbide bit first (speed set to an of output ~9-10). This creates a "track" within which to thin the skull.
      CAUTION: To minimize heat injury and achieve a smooth track, keep moving the drill bit. Do not drill the same place for more than 2 s.
   3. Periodically, stop drilling and irrigate the whole area with sterile saline, to reduce heating from the drill and to wash away the bone dust.
   4. Continue to thin the skull using an FG1/4 carbide bit, as described in step 3.5.2, until the skull is paper-thin and transparent.
      NOTE: Using two hands to hold the drill can make it easier to control the drill and avoid inserting the drill bit into the brain.
   5. Complete the skull thinning using an EF4 carbide bit. When a crack occurs between the bone flap and the surrounding skull bone, sometimes there is a release of cerebral spinal fluid (CSF), indicating that the skull has been completely drilled through.
   6. Continue thinning and drilling through the rest of the skull along the track. Avoid drilling through the point where an obvious vasculature crosses under the skull to prevent bleeding.
   7. Insert a fine forceps tip (forceps 1) ~0.5 mm through the cracked place, and lift the bone flap gently upward without indenting the underlying brain.
   8. After the bone flap is removed, irrigate the craniotomy area with saline. The brain surface could be 1-2 mm higher than the craniotomy edge.
   9. Starting from this step, always cover the exposed brain with saline to protect the brain tissue.
   10. Bleeding may occur when lifting the bone flap at the original AAV injection sites due to the tissue adhesion and vasculature growth after the injection surgery. If this happens, gently press the site with a dry cotton tipped applicator for ~2 min (or longer as needed). Gel foam can be used to stop the bleeding. Do not use chemicals or heating forceps to stop the bleeding, as this can cause injury.
   11. Use the forceps 1 to gently remove the visible arachnoid matter.
6. Implant glass window
   1. Use the surgical forceps 2 to pick up the sterile glass window, with the 3 mm glass coverslip facing down. Place and adjust the glass window above the craniotomy site to make sure the window can fit snugly to the craniotomy edge. The 5 mm glass coverslip is on top.
   2. Prepare dental cement as follows: combine 100 mg of cement powder, two drops of quick base liquid, and one drop of catalyst. Stir the mixture until thoroughly mixed (~15 times). Wait ~6 min until the cement becomes pasty and thick. If the cement is too thin, it may seep into the space under the window in step 3.6.4 and obscure the window.
   3. While waiting for the cement to become pasty and thick, apply an adequate amount of pressure to the window through a stereotaxic manipulator, to check that the skull can securely and tightly contact the glass window. Ensure that the dental cement liquid in step 3.6.4 will not reach the space underneath the glass and thus obscure the window.
   4. Use an adjustable precision applicator brush to add a small amount of cement alongside the window edge to seal the glass window with the skull. Wait for ~10 min to let the cement completely dry, and then gently release and remove the manipulator above the window. As of this step, it takes ~4 h to finish the 10 TBI impacts and implant the headpost and cranial window.
   5. Trim the dental cement using the dental drill with an FG4 carbide bit if there is excess cement covering the window.
      NOTE: Excessive cement around the window may prevent the two-photon objective lenses from approaching the window surface.
7. Implanting the imaging well
   NOTE: An imaging well (Figure 2D) is a rubber ring with an outside diameter of ~1.6 cm, that matches the headpost top surface and holds water above the cranial window for two-photon imaging.
   1. After the window implantation is completed, drill away the dental cement debris on the headpost top surface, and clean the area using wet surgical gauze. Allow the area to dry for ~3 min.
   2. Use a cotton tipped applicator to dip a small amount of superglue and paste it on the headpost top surface. Quickly place the rubber ring onto the headpost. Apply medium pressure on the rubber ring for ~2 min to ensure close contact with the headpost. Use superglue sparingly, otherwise, it can adhere to the glass window and obscure the two-photon imaging.
8. Analgesic and antibiotic administration
   1. Immediately after implanting the imaging well, but prior to discontinuing isoflurane, administer cefazolin [500 mg/kg (333 mg/mL, usually ~45 µL), intramuscularly], and meloxicam (5 mg/kg, subcutaneously) using disposable insulin syringes.
   2. After drug administration, discontinue the anesthesia, release the mouse from the stereotaxic frame, and return the mouse to its home cage, which is above a heating blanket.
   3. Closely monitor the mouse for ~15 min until it is ambulatory.
      NOTE: House the mice individually, as they may bite the rubber imaging well of other mice. Provide food and water gel close to the mouse in the cage for access ad libitum.
   4. Administer buprenorphine (1 mg/kg, subcutaneously) and meloxicam (5 mg/kg, subcutaneously) again every 8 h after the previous dose until 48 h after the cranial window implantation surgery.

4. Intravital two-photon imaging

1. Use microscope 2 (Table of Materials), equipped with a tunable coherent multiphoton laser and a 20x magnification objective (NA 1.0; water-immersion), for intravital imaging¹⁸. The filter used for the EGFP signal is "BP 500-550".
2. Prepare the cranial window mouse for intravital imaging.
   1. Starting from the surgery day (designated as day 0), carry out two-photon imaging at designed time points post-TBI. Place the cranial window mouse into the anesthesia induction chamber, and administer 3% isoflurane mixed with carrier gas at a 1.5 L/min flow rate for 5 min.
      NOTE: The carrier gas can be either room air or 100% oxygen.
   2. Once the mouse is fully anesthetized (i.e., slow but steady breathing rates at approximately one cycle per 2 s, coupled with the absence of a tail-pinch reflex), remove the mouse from the induction chamber and quickly clamp the headpost to a bracket. Let the mouse's torso lay on a round plastic plate (19 cm diameter) that is equipped with the bracket.
      NOTE: A hand-warm pad was placed under the mouse's torso to provide heat support during intravital imaging.
   3. Apply lubricant ophthalmic ointment onto the mouse's eyes and place the anesthesia tubing nose cone over the mouse's snout. Maintain anesthesia by using 1.5% isoflurane with carrier gas at a 1 L/min flow rate.
   4. Periodically check the breath frequency, and deliver a tail or toe pinch at least every 15 min throughout imaging to assess appropriate anesthesia. Adjust isoflurane concentration as appropriate to maintain the desired depth of anesthesia.
   5. Align the mouse head to ensure that the cranial window is directly underneath the two-photon objective lens. Add some water into the imaging well above the cranial window. Lower the objective so that it is immersed in the water.
3. Intravital imaging of intracranial window mice
   1. Turn on the scope mercury lamp. View the brain with epifluorescence through the ocular first.
      NOTE: The vasculature appears black. Select an area where the window is clear. Turn off the epifluorescence, set the laser wavelength to 860 nm for the EGFP signal, and adjust the laser setting for optimal (i.e., bright but not saturated) signal.
   2. Set the scan mode to Frame and the line step to 1. Set the averaging number to 16, the bit depth to 8-bit, the mode as Line, and the method as Mean.
   3. Use the vasculature pattern as a "reference map" to image the same brain region for subsequent longitudinal imaging. Image the superficial level (layer I, less than 100 µm from the meningeal surface) for three planes where the cortical vasculature dominates through a z-stack mode, with an interplane interval of 10 µm, as indicated in Figure 3A.
   4. Image six planes at deep level (layer IV and V, ~400 µm deeper than the superficial level), through a z-stack mode as indicated in Figure 3A, with an interplane interval of 10 µm and a scanning speed of 8.
   5. After finishing the imaging (~20 min per mouse), discontinue the anesthesia, release the mouse from the frames, and return it to its home cage that is above a heating blanket. Monitor the mouse consecutively until it is ambulatory, which usually takes ~7 min.
   6. Longitudinally image the mouse at day 0, 1 week, and 4 months post-TBI surgery.