Hey, guys, my name is Wang Yangdong, and I'm from Dr.Kiryl Piatkevich molecular bio-engineering lab in Westlake University. Today, I'm going to show you the procedure for imaging neuronal activities of hippocampus in mice, in vivo. Turn on the welding machine and heat it up to the required temperature.
Adjust the position of the imaging cannula and the injection cannula using helping hands as shown in the figure. Using the syringe with the needle, apply a small amount of appropriate type of flux onto the connection spots between imaging and injecting cannula for five seconds, and then remove the droplet. Mount soldering tin and apply it to the connection spot treated with the flux.
Avoid excess soldering tin as it will require unnecessary larger cranial to make during the surgery. Wait for soldering tin to cool down. Usually, it takes several seconds.
Confirm that the injection cannula is not blocked by inserting the dummy cannula from both directions. Apply UV curing optical adhesive on the bottom side of the imaging cannula using a toothpick or 26-gauge syringe needle. Use a fine tweezer to carefully place a cover glass of the corresponding size to the imaging cannula.
Cure the adhesive for at least an hour by UV illumination from a standard handheld UV lamp. Anesthetize the mouse with isoflurane according to-IACUC approved procedure. Place the mouse in a stereotaxic frame over a surgery heating pad.
Secure the head with ear bars. Slightly push the head in all directions to make sure the head is secured firmly. Apply eye ointment to prevent the animal's eyes from drying out during the surgery.
Use a trimmer or depilatory cream to remove the fur from the back of the neck up to the eyes. Sterilize the surgical site with betadine followed by 70%ethanol three times before making an incision. Remove the skin over the top of the skull, starting with the horizontal cut all around the base of the head, followed by two cuts in the rostral direction, almost reaching the eyelids.
Then two oblique cuts that converge at the midline. With two sterile cotton swabs, retract the connecting tissue, as well as the musculature of the back of the neck to the edges of the skull. Apply a drop of lidocaine solution to the surface of the periosteum for two minutes to avoid excessive pain.
Gently scrape the entire exposed area of the skull with a scalpel to create a dry and rough surface. Place tip of the needle onto the Lambda to see if the AP coordinate is zero to confirm that the head position is vertical, as well as if the ML coordinate is zero, to confirm the head is positioned horizontally. If not, adjust the correspondent knobs on the stereotaxic station until the AP and ML coordinates are both within 0.1 millimeters.
Move the tip of the needle to find the corresponding points for craniotomy and mark their positions on the skull, using a fine marker. Draw a circle based on four marked points, as well as the outline of the injection cannula area on the cannula side of the circle. Use a pneumatic drill at the speed of 10, 000 rounds per minute to gently drill along the outline marked on the skull.
Drill the skull until a very thin layer of bone is left, which usually starts to wiggle under gentle touch in the center. Apply a drop of sterile PBS to the center of the craniotomy. Lift the bone flap from the skull with very thin tipped forceps or two 26-gauge needles approaching it from the opposite sides.
Apply PBS followed by gentle aspirations, through a 26 gauge blunt needle several times to clean the surface of the dura. Apply gentle suction to ablate the cortex as well as the Corpus Callosum above the hippocampus. Cortex is often yellower than the Corpus Callosum.
And the Corpus Callosum is usually wider than the hippocampus. Besides, the Corpus Callosum is really easy to distinguish by neuronal fibers going in the vertical and horizontal directions when observed from the top. Bleeding at this point will affect the visibility of the brain tissue in the craniotomy.
Apply PBS, followed by gentle suction, while you're aspirating the cortex to get rid of the blood, Corpus Callosum can be recognized by the direction of the white fibers as schematically shown in the diagram. Remove Corpus Callosum fibers layer by layer by applying gentle suction in a circular motion. PBS is constantly applied to get rid of blood accumulating above exposed brain tissue.
Gently insert the implant into the craniotomy. Firmly press on top of the implant with the L-shaped needle to position the optical window of the implant as close as possible to the exposed surface of the hippocampus. Repeatedly apply PBS on the scar around the implant, followed by suction to remove blood as much as possible during implant insertion.
Apply a thin layer of quick seal between the implant and skull to prevent dental cement from penetrating under the skull. Once the quick seal is cured, apply super bond evenly on the surface of the skull, the surface of the quick seal and the surface of the implant. Once the super bond is cured, apply denture base resin above super bond, as well as the skin around the incision made at the beginning of the surgery.
After the denture base resin is cured, place the head plate on the resin around the implant and make it concentrate with the imaging cannula. Apply more denture base resin around and above the head plate to fix its position. Let it cure for several minutes.
Avoid building up a thick layer of cement around the cannula to ensure better access to the imaging window is the objective lens. Dilute the denture base resin to decrease its viscosity, thus allowing it to fill the cavities which are hard to reach with an applicator. Gently place a insulated rubber tape above the window to protect the window from possible contamination from animal banding.
Add Fast Green dye stock solution to virus solution dilute it to the desired titer in the ratio of one to nine, in a PCR tube. Withdraw 600 nano liters of the virus solution, remove the dummy cannula and insert the internal cannula connect it to the injection syringe into the guide cannula. Infuse the virus at the speed of 15 nano liters per minute, for 10 minutes in total.
Keep the internal cannula connected for 10 minutes to allow the virus to spread under the window. Gently remove the internal cannula from the guide cannula and recap it with the dummy cannula. Induce the mouse with 4%isoflurane for a few minutes.
Fix its head plate to the head fork, and then fix the head fork to the treadmill. Use a low magnification objective lens to find the best to filter view for functional imaging. Then, switch to a higher objective lens to record the neuronal activities at single cell resolution.
For calcium imaging, green fluorescent was excited by a commercially-available 470 nanometers LED using a standard GFP filter sense. To obtain fluorescent traces, the regions of interests, corresponding to neuronal cell mass were segmented manually and analyzed the by ImageJ software. For voltage imaging using a 40 X lens optical voltage recordings were performed in the near infrared channel at 830 Hertz acquisition rates.
Spontaneous neural activity recording lasted continuously for up to 25, 000 frames. Here, we describe a method for long-term imaging of the hippocampus as a A1 region in behaving mice. The method is based on the chronic implantation.
Now the custom-made imaging window, which also enables the targeted administration of viruses or drugs directly to the neurons of interest. We believe that the described protocol will facilitate the studies that aim to investigate neuronal activity with high spatial temporal resolution in the hippocampus of behaving mice using simple and affordable 1-photon imaging set-ups.
