Preliminary Validation of Stereotaxic Injection Coordinates via Cryosectioning

All animal experiments were conducted in compliance with the Animal Research Reporting In Vivo Experiments (ARRIVE) guidelines and the National Institutes of Health Guide for the Care and Use of Laboratory Animals. The present study was approved by the Animal Care and Use Committee of Renji Hospital, Shanghai Jiaotong University School of Medicine. Eight-week-old C57BL/6J male mice were used for the present study. The animals were commercially obtained (see Table of Materials) and housed in standard cages (22 °C ± 2 °C, 12 h/12 h light/dark cycle, food, and water ad libitum).

1. Selection of target brain regions

NOTE: This is a sterile procedure. Ensure all surgical instruments are sterilized. Spray the gloves with alcohol prior to contact with surgical instruments. Avoid touching the tip of the instruments or the surgical area with gloves. Ensure the mouse is adequately anesthetized before starting the procedure by checking for the loss of righting reflexes and no response to cutaneous stimulation. Before the drilling and injection, closely monitor the respiratory rate (60-220 breaths per min, average 100-150 breaths per min) and the breathing amplitude (should be consistent and not excessively shallow or deep). Any significant changes in the respiratory rate or amplitude should be noted.

1. Anesthetize the mouse with an intraperitoneal injection of 1.25% tribromoethanol (250 mg/kg) using a 29 G needle. Shave the hair on the skull once the righting reflex disappears. Secure the mouse in the stereotaxic frame (see Table of Materials) by fixing the upper incisors with a nose clamp and stabilizing the head using two ear bars.
2. Apply ophthalmic ointment to prevent eye dryness. Disinfect the scalp by swabbing it three times with Anerdian (disinfectant solution; see Table of Materials). Inject 0.2 mL of 1% lidocaine subcutaneously using a 29 G insulin needle to provide local analgesia.
3. Make a 0.5 cm incision using eye scissors to expose the cranium. Utilize a cotton swab dipped with 3% H₂O₂ to scrub the exposed skull to remove the periosteum. Allow the cranium to dry.
4. Affix the drill to the stereotaxic instrument. Adjust the stereotaxic arms' knobs to position the drill tip precisely at the bregma with the help of a magnifier and record the z-axis value using the LCD digital display.
   1. Adjust the knobs and position the drill tip at the lambda and record the value of the z-axis. Adjust the nose clamp and ear bars to make two z-values closer, and repeat the steps above until the difference between the two z-values is less than 0.1 mm.
      NOTE: The x, y, and z axes on the LCD digital display correspond to mediolateral (ML), anteroposterior (AP), and dorsoventral (DV), respectively. The bregma and lambda can be considered to be on the same horizontal plane once the difference between the two z-values is less than 0.1 mm.
5. Ensure the left-right levelness of the brain. Adjust the ear bars attached to the stereotaxic frame with the same scale. Take the laterodorsal tegmental nucleus, ventral part (LDTgV), for example; its stereotaxic coordinates are -5.2 for x, +0.8 for y, -4.0 for z, according to Paxinos and Franklin's mouse brain atlas¹⁶.
   1. Adjust the ear bar to make the drill at (-5.2, +0.8) to measure the z value in the dorsal-ventral direction and move the drill to (-5.2, -0.8) to measure the z value.
      NOTE: The left-right is considered to be at the equal level when the difference between the two z-values is less than 0.2 mm. The ear bars need to be inserted into the right place because the external auditory meatus is symmetric. If the difference between the two z-values is more than 0.5 mm, the ear bar may be inserted into the wrong place.
6. Position the drill to the bregma again and set three coordinates on the LCD digital display to zero. Position the drill above the region of interest using reference coordinates. Start the drill.
   1. Gradually lower the drill using the vertical arm until it penetrates the cranium. Clear away debris and blood using cotton swabs.
      NOTE: Be careful not to damage brain tissue.

2. Preparing dye solution and brain microinjection

1. Take 25 µL of SDS-PAGE loading buffer containing bromophenol blue (see Table of Materials) into a 200 µL microcentrifuge tube. Add 50 µL of ddH₂O to the tube to prepare the dye solution.
2. Cover the cranial window with a saline-moistened cotton pad. Ensure microsyringe patency by aspirating and expelling saline repeatedly. Load 0.3 µL of dye into the micro syringe. Attach the syringe and the motorized stereotaxic microinjector to the stereotaxic arm.
3. Position the syringe tip at the bregma and set the coordinates to zero on the LCD digital display. Adjust the needle into the region of interest according to reference coordinates. Set the injection speed to 0.1 µL/min and start the motorized microinjector, delivering 0.3 µL of blue dye into the target region. Leave the syringe in position for at least 10 min before slowly withdrawing the injector.
   NOTE: Different color dyes can be used in the same mice to distinguish closely located regions. Test other dyes for suitable concentrations based on their physical properties. It's recommended to use the bromophenol blue formula and thoroughly rinse the syringe before and after use to prevent blockages. Adjust injection volumes based on dye diffusion.

3. Brain tissue harvest and cryosectioning

1. Release the mouse from the stereotaxic frame. Anesthetize the mouse with an intraperitoneal injection of 50 mg/kg pentobarbital sodium using a 29 G needle. Secure the mouse on a foam board with tape.
2. Open both sides of the thorax to induce rapid suffocation. Make a 1-2 mm incision in the right atrial appendage. Insert an infusion needle into the left ventricle. Infuse 20 mL of pre-cooled saline and 20 mL of iced 4% paraformaldehyde solution.
3. Decapitate the mouse with tissue dissecting scissors (following institutionally approved protocols). Remove the skin by making an incision along the midline from the neck to the nose and then expose the skull. Clear off residual muscle on the skull with forceps or scissors.
4. Position the tip of one blade of the fine scissors between the foramen magnum and the brain, with the sharp edge facing the bone, at the rostral end of the mid-sagittal suture. Proceed to slide and cut the cranium along the mid-sagittal suture. Employ forceps to remove the skull, revealing the exposed brain.
   NOTE: Exercise caution to avoid causing any damage to the brain during the operation. Before removing the bone chips, carefully peel off any meninges that may be attached to the skull. This precaution is crucial to prevent inadvertent slicing of the brain tissue during the process.
5. Turn on the power of the rotary microtome cryostat (see Table of Materials) and set the chamber temperature to -20 °C. Pour a little OCT compound to cover the surface of the specimen disc evenly. Place it into the cryo-chamber and allow the OCT to solidify.
6. Attach the disc to the specimen head. Bring the blade closer to the block and trim the OCT block to make a flat plane that is parallel to the disc.
7. Cut the brain in a coronal direction away from the injection site in a brain slice mold by a shaving blade. Place the brain tissue containing the injection site with the cut side down on the OCT flat plane.
   1. Pour OCT onto the brain and put the disc into the cryo-chamber, allowing the specimen block to solidify. Pour OCT repeatedly onto the brain until the brain is fully embedded with OCT.
      NOTE: Add the OCT compound layer by layer. Once the previous layer of OCT is frozen, pour the subsequent layer of OCT compound onto the brain.
8. Trim the specimen block until the target region is approaching. Make several sections from the injection site level. Collect the coronal brain slices into a 6-well plate containing room-temperature PBS using a small writing brush kept at -20 °C.
   NOTE: Set the cutting thickness to a larger one to trim the block, and then set the cutting thickness to 40 µm as the blade approaches the injection site. If neural circuit tracers are used, the mice are not sacrificed immediately. Place the mouse in a sterile, postoperative recovery cage over a heating blanket for recovery from anesthesia. Return the mice to the original cage or a new cage once awake.

4. Imaging

1. Wash the OCT with room temperature PBS. Use a brush to pick up brain sections and place them on a slide. Allow sections to air dry at room temperature.
2. Observe the injected region visually or use a microscope when the target nuclei are minuscule. Compare the location of the injected dye with the corresponding region in the brain atlas.
   NOTE: Adjust the values of the x, y, and z axes according to the direction and distance of the injection point deviation. It may be necessary to repeat all steps in this protocol multiple times to determine the coordinates accurately.