Fluorescence Imaging of Neuro-Immune Dynamics Using a Skull Implant

Protokol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Mouse preparation for cranial window implantation

NOTE: Various transgenic mouse lines with fluorescent tags are suitable for imaging.

1. Use CX3CR1GFP/+ mice to visualize microglia in vivo. Typically, juvenile to young adult 4 to 10-week-old mice that weigh 17-25 g are used.
   NOTE: Although this approach is even apt for pre-weaned mice, the need to return the mice to their cages with their mothers for feeding may complicate recovery if the mother does not take adequate care of the pups post-surgery. Therefore, the use of mice post-weaning is recommended.
2. Anesthetize the mouse using isoflurane (5% flow in oxygen for induction for 1 min) in an anesthetic chamber. Check that the mouse doesn’t show any movement or twitching responses to toe and/or tail pinches. Take the mouse out of the chamber and, in the open air, thoroughly shave the hair on the head between the ears from about eye level to the top of the neck region using a hair trimmer.
   NOTE: The concentration of isoflurane used would depend on the size of the induction chamber. Therefore, for smaller chambers, 3-4% isoflurane can be used to effectively induce anesthesia, while larger chambers will require up to 5%.
3. Move the mouse to the stereotactic surgery station nose cone for anesthesia (1.5-2% for maintenance during the surgery), stabilize its head using ear bars, and keep the mouse on a heating pad to keep its body temperature warm.
4. Lubricate both eyes with eye ointment. Inject 100 µL of 0.25% bupivacaine (to provide local analgesia to the mouse that will last 8-12 h) and 100 µL of 4 mg/mL dexamethasone (to reduce the inflammation that may result from the surgery procedure) subcutaneously at the incision site. Allow the mouse to sit for at least 5 min before moving to the next step.
5. Clean the shaved head with three alternating swabs of betadine and 70% alcohol. Using a surgical blade or scissors, make a midline scalp incision extending from the back of the skull region between the ears to the frontal area between the eyes. The remaining skin is cut to expose the skull.
6. Clean the connective tissue located between the scalp and the underlying skull with 3% hydrogen peroxide (H₂O₂) and localize the brain area to be imaged with stereotactic coordinates.
   NOTE: There is often some bleeding (step 1.5) from the incision on the skull surface. This bleeding usually resolves by itself within 3-5 min. Cleaning with the peroxide helps. Prior bupivacaine treatment (step 1.4) is also noted to limit the amount of bleeding during this time.

2. Mouse cranial window implantation surgery

1. Drill a circular opening ~4 mm into the skull using a dental drill bit (0.7 mm tip diameter) and carefully remove this portion of the skull using pointed forceps. For imaging the somatosensory cortex of 6-8-week-old mice, locate the center of the craniotomy at -2.5 posterior and ± 2.0 lateral to bregma. During drilling, regularly moisten the skull with sterile saline and cotton swabs to cool the brain, clean off bone debris, and soften the skull bone for eventual removal.
   NOTE: The coordinates for the craniotomy would vary depending on the region of interest and the mice's age.
2. After removing the skull, carefully place a small coverglass (size #0 at 0.1 ± 0.02 mm thickness) moistened with saline in the craniotomy. Dry off excess saline using a sterile wipe.
3. Using a pointed applicator (such as a pipette tip or the pointed end of a broken wooden cotton swab stick), apply the cyanoacrylate glue around the window and allow it to attach to the brain and skull. Apply the primer glue to the rest of the skull and cure it with a curing light for 20-40 s. Prepare a well around the window with the final glue and cure with a 20-40 s curing light.
4. Glue a small head plate onto the skull on the contralateral hemisphere of the craniotomy first with the primer glue as a primer and then with the final glue. Cure both with the curing light for 20-40 s each.
   NOTE: Sutures are unnecessary if the skull is covered with glue during this procedure.

3. Post-surgery care

1. Allow the mouse to wake up without anesthesia (recovery done on a heating pad shortens the recovery time) and return it to its home cage once fully awake. Inject one subcutaneous dose of buprenorphine SR (0.5 mg/kg) as post-operative analgesia that is sufficient for 72 h.
2. To facilitate a healthy recovery from the surgery, provide the mouse with extra-soft food, such as regular solid chow soaked in water to soften it or food in the form of a gel.
   NOTE: A one-time provision of soft food immediately after the surgery is sufficient.
3. Monitor the mouse daily for health and proper recovery for the first 72 h of the surgery procedure. Afterward, imaging from the window implantation surgery will be performed as early as 2 weeks.
   NOTE: If done well, mice recover well showing normal ambulatory behaviors, sufficient cage exploration, good hydration, stable weight gain, and extensive interactions with other mice in the cage and other items in the cage. Mice showing lethargy, dehydration, and greater than 10% weight loss following the surgery are euthanized and removed from the study.

4. Two-photon brain mapping for initial imaging

1. Anesthetize the mouse (Isoflurane, 5 % induction and 1.5 % maintenance). Stabilize the head using screws to mount the headplate on the two-photon microscope stage, being maintained on a heating plate at 35 ˚C. Inject intraperitoneally 100 µL of blood vessel dye such as Rhodamine B (2 mg/mL).
   NOTE: Imaging could also be done in awake mice without anesthesia. However, recent studies indicate that anesthesia affects microglial surveillance dynamics. Head fixation for two-photon imaging in awake mice increases stress even during chronic imaging for at least 25 days.
2. Clean the surface of the cranial window gently using a cotton swab dabbed in 70% ethanol. Put a few drops of water or saline on the cranial window and lower the objective lens into the solution since the objective is an immersion lens.
3. Hand-draw a coarse map to denote the major blood vessel landmarks in a lab notebook while looking through the eyepiece by epifluorescence. Use this drawing to identify the specific regions during two-photonn imaging. Alternatively, take pictures of the blood vessels either through a camera fitted to the microscope or through a hand-held camera or phone.
   NOTE: These hand-drawn images and pictures are to facilitate revisiting the same broad regions under the microscope before two photon imaging. These are not precise image mapping.
4. Under two photon imaging, collect images of florescent cells and vessels as needed. Take careful notes with appropriate coordinates to ensure that that the precise regions can be revisited for subsequent imaging. Collect several fields of view in this initial imaging session e.g. acquire z-stack images every 1-2 µm through a volume of tissue.
   NOTE: While collecting images by two photon, the blood vessel landmarks are used for coarse mapping. If fine mapping is needed, YFP-labeled dendrites from Thy1-YFP mice are used.
   1. Use these recommended parameters for imaging: a wavelength of 880-900 nm is optimal; for GFP and/or dsRed / Rhodamine excitation, a 565 nm dichroic mirror with 525/50 nm (green channel) and 620/60 nm (red channel) emission filters are used; for GFP and YFP separation, a 509 nm dichroic mirror with 500/15 and 537/26 nm emission filters are used; the power at the brain is maintained at 25 mW or below; image resolution is 1024 x 1024 pixels, the field of view taken with a 25X 0.9 NA objective at a 1.5X zoom factor is 295.24 x 295.24 µm.
5. At the end of the imaging, take the mouse off the stage, allow it to wake up from anesthesia and return to its home cage until a future imaging session.

5. Two-photon imaging and re-imaging

1. For future subsequent imaging sessions, which could be anywhere from a few hours to months after the initial imaging session, anesthetize the mouse (Isoflurane, 5% induction and 1.5% maintenance), mount on the two-photon microscope, maintain on a heating plate and re-inject 100 µl a blood vessel dye such as Rhodamine B (2 mg/mL).
2. Open the previously obtained images in ImageJ and, using these images as well as the notes from the previous session, identify the previously imaged areas and carefully re-image them.
3. Repeat this for as long as the imaging window is clear or as essential for the extent of the study.

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