All experimental procedures described below were approved by Local Ethical Committee at the Nenscki Institute of Experiemental Biology, Polish Academy of Sciences. This video shows craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two-photon microscopy in thi one-G of feet transgenic line. This approach is combined with injection of mCherry expressing adeno-associated virus, AAV, in the dorsal hippocampus.
Combined together, these techniques allow long term monitoring of experienced dependence, structural plasticity in retrosplenial cortex. In this paper, we propose implantation of the cranial window above the retrosplenial cortex as not a possible region of interest for the two-photon in vivo microscopy. In order to visualize the morphological changes in neuronal structures, we use thi one-G of B mice with the expression of fluorescent GFP protein in approximately 10 percent of neurons in the brain.
Another innovation that we propose is the injection of an AAV, expression of a fluorescent protein mCherry under a neuron-specific promoter into the deeper structures in the brain, such as hippocampus, in order to visualize projections of the structure into the retrosplenial cortex. Sterilize all tools, glass containers for liquids and cotton swabs in the autoclave. You dispensable gloves.
Clean the surgical table, the stereotactic frame and all the surrounding area with 70%ethanol. Use a sterile surgical pad to create a sterile space for all the sterilized equipment. Cut gel foam into small pieces and soak them in sterile saline.
Put the animal in the induction chamber and set isoflurane level to 5%and oxygen flow to two liters per minute. This procedure should take about three minutes. Take the animal out of the induction chamber.
Use tail or toe pinches in order to insure that the animal is fully sedated. Using precise trimmers, shave the hair from the back of the head, between the ears, up to the eyes. Place the the animal in the stereotactic frame and stabilize the head with ear bars.
Set anesthesia levels to 1.5 to 2%isoflurane and zero point three liter per minute oxygen. Apply the eye ointment. Inject the animal subcutaneously with tolfedine, four milligrams per kilogram, butomidor, two milligrams per kilogram, and Baytril, five milligrams per kilogram to prevent inflammation, pain and infection respectively.
Inject the animal intramuscularly with dexamethasone, zero point two milligrams per kilogram, to prevent brain swelling. Clean the skin using sterile cotton swabs with betadine followed by 70%ethanol. Change the gloves and spray them with 70%ethanol.
Lift the skin with forceps and using microscissors incise skin horizontal along the base of the head and then obliquely to the front point between the eyes. Remove the skin flap. Apply lidocaine ointment with sterile swab on the periosteum to prevent excessive bleeding and pain.
Use sterile cotton swabs or scalpel to remove the periosteum. Dry the skull with sterile swabs. Using sterile needle, apply dense cyanoacrylate glue on the skin edges to immobilize them and prevent from further contact with dental cement.
Wait for the glue to dry. Lay a sterile three millimeter cover glass over the skull anteriorly to the lambdoid suture. Center the cover slip at retrosplenial coordinates, anterior, posterior bregma minus-two point eight, medial lateral bregma zero.
Mark the cover slip edges by scratching skull surface with a sterile needle. Put the cover glass back into the sterile container with 70%ethanol. Use a high-speed dental drill with small-diameter bur to outline a three millimeter diameter circle.
Clean the drilling site from the bone dust with sterile saline-tipped swabs. Use the gel foam and swabs to stop the occasional bleeding and clean the bone. In between drilling, check the bone thickness with fine forceps by gently touching the bone circle and checking its mobility.
Keeping in mind that the bone is thicker on the sutures area. Stop the drilling when the bone circle is mobile and only an even thin layer of bone is left on the circumference. Clean the operational field of all the remaining bone dust with saline-tipped swabs.
Drop the sterile saline on the drilling area, covering the drilled circle. Carefully pry the bone circle with a fine forceps and then gently but firmly remove the bone by lifting it upwards. Be careful not to skew the bone circle while lifting it to prevent possible damage to the dura.
Gently apply the gel foam soaked in sterile saline on the dura to help top the bleeding. Wait until all bleeding is fully stopped. Carefully remove the gel foam not to disturb the clotting process.
Note, the suture area is high vascularized so the bleeding at this point might prove to be profound. It is essential to wait for the sufficient time for the bleeding to stop. Attach the infusion pump to the stereotactic tower and connect the controller.
Insert the 35-guage needle into the fill syringe. Flush the syringe 10 times with ethanol to sterilize it and 10 times with sterile saline to remove traces of ethanol. Remove air bubbles from the syringe.
Insert the syringe into the pump. Fill a single dose of AV preparation and keep it on ice. Fill the syringe with virus solution.
Center the needle on bregma and then gently insert into the hippocampus using the following coordinates:Anterior, posterior minus-two, medial lateral plus minus-one, dorsal ventral minus-one. These coordinates will be located near the edge of the cranial domain. Wait for five minutes for the tissue to stabilize.
Inject zero point seven microliters of AV solution at the rate of 50 nanoliters per minute. Wait 10 minutes for the virus to fully absorb. Gently remove the needle.
Blot with gel foam if bleeding occurs. Repeat with the contralateral side. Lay the sterile dry cover glass on the top of the dura in the drilled circle frame.
Hold the cover glass with the foreceps to gently flatten the dura and bring cover glass edges closer to the skull surface. Using a sterile needle, apply the dense cyanoacrylate glue on the cover glass edges to attach them to the skull. Wait for the glue to dry.
Place a fixation bar, anton nut or a custom-made design in the front part of the skull. Note, the fixation bar should be placed in a position that will enable horizontal positioning of cranial window during imaging session. Apply the glue over the edges of the bar.
Wait for the glue to dry. Note, the fixation bar should be placed as distant as possible from the window. If it is placed too close to the window, the bar and the screw connecting it with the custom-made holder might pose to be an obstacle for the objective during the imaging process.
Prepare the dental acrylic and apply it on the skull surface around the glass. It is helpful to form a crater-like shape around the window. It will create a cavity for the water applied later for imaging with the water objective.
Create a cap with the dental acrylic, covering the rest of the operational area skin edges fixation bar re-enforcing the crater around the cranial window. Wait for the dental cement to harden. Remove the animal from the stereotactic frame and put it into the recovery chamber.
Wait for the animal to recover from the surgery while observing the physiological functions. Apply postoperative anesthesia cyroproferrin, 10 milligrams per kilogram, and antibiotics treatment baytril, five milligrams per kilogram, for 48 hours. Start the ti sapphire laser, power up the microscope.
The system used in this experiment is equipped with a two-photon laser, OPO system and dual-gallium arsenide falsified PMT. Put the animal in the induction chamber and induce anesthesia. Remove the animal from the induction chamber and place in the gas anesthesia mask under the microscope.
Decrease the oxygen flow to zero point three liters per minute and the isoflurane concentration to 1.52%Fix the animal to the custom microscope frame with an MT screw or another custom system. Level the cranial window. Note, it is possible to use the microscope manufacturer's head fixation system, although the specific custom frame gives better results, improved head stability, constant positioning in multiple sessions during a chronic experiment.
Using the wide-field microscope settings, a low magnification objective, center the view on one of the sides of the retrosplenial cortex and focus it on the cover slip surface. Apply a droplet of water into the crater-like acrylic well. Switch to a long-distance water immersion objective.
Move the objective towards the cranial window until the water meniscus connects the specimen and objective. Switch to two-photon settings and begin scanning the specimen top to bottom using lowest zoom. The crossing of dura matter will be visible as glare of high non-specific signal.
Adjust microscope acquisition settings in both channels, GFP and mCherry according to the signal strength from florescent cells in order to cover the entire dynamic range. After finding a suitable neuron with the dendritic tree separated from other cells, perform an initial scan using only GFP filters set with lowest zoom and Z distance of five microns. Obtain a maximum projection of the scan stack and print it for annotations using inverted colors.
Set zoom to a value that will allow to image the desired morphological details. Image the entire dendritic tree in GFP and mCherry channel using maximum projection as guide. Using this protocol, it will be possible to repeatedly monitor both pre and post-synaptic structures in the retrosplenial cortex.
This approach could be utilized with a chronic experiment where behavioral manipulations expected to correlate when changes in morphology of dendrites, synaptic spines or pre-synaptic neutrons. The imaging sessions can be performed at any frequency, but 24-hour time interval is preferred in order to allow proper recovery of the animal and to minimize the effect of anesthesia. If applied properly, this technique allows performing multiple imaging sessions over the course of several months.
