An Invasive Method for the Activation of the Mouse Dentate Gyrus by High-frequency Stimulation

Animal experimental procedures followed the institutional guidelines of the Beijing Institute of Basic Medical Sciences (Beijing, China) and the Chinese governmental regulations for the Care and Use of Laboratory Animals. The mice (adult male, 26 ~ 30 g) were housed and kept at a constant temperature of 23 °C, with water and food ad libitum, under a 12-h light/12-h dark cycle (lights on at 7:00 a.m.). All experimental procedures were performed during the light cycle.

1. Surgical Preparation

NOTE: A custom electrode was homemade using the method modified from Halpern's report²³.

1. Use 2 copper micro-wires (with an outer diameter of 200 µm) as the parallel bipolar electrode. Wrap the electrodes around each other to form cylindrical electrodes for the stimulation.
   NOTE: Here, the electrode length within either the PP or DG was varied according to the anatomical depth of the nucleus.
2. Sterilize the electrodes using an Ethylene oxide disinfection cabinet. Handle and store the homemade 2-channel micro-wire electrodes without any contamination. The electrodes should be strictly sterilized before any surgical implantation.
3. Autoclave the surgical instruments.
4. Before the surgery, inject the mouse with pentobarbital (20 – 50 mg/kg) intraperitoneally, to achieve a surgical plane of anesthesia.
5. Inject the mouse with a sterile penicillin G procaine suspension (75,000 U, I.M.) and an analgesic agent (carprofen, 0.05 – 1 mg/kg, SC) to decrease the risk of infections and pain, respectively²⁴.
6. After the animal anesthesia, apply an eye lubricant to both eyes to prevent eye dryness. Ensure the induction of the sedation and the depth of the anesthesia by the animal's reflexes in response to toe pinching and proceed with the operation only when no response occurs.
   NOTE: Vet ointment is another option to prevent eye dryness.
7. Shave most of the fur on the top of the mouse head with electrical hair clippers or a razor blade, sterilize the shaved area with three alternating scrubs of 70% alcohol and betadine solution and proceed with the operation after the betadine has dried on the skin.

2. High-Frequency Stimulation Surgery^25,26

NOTE: In this step, an electrode is unilaterally implanted into the dorsal DG or medial PP area, a part of the hippocampus with critical roles in episodic learning and memory.

1. Place the anesthetized mouse onto the stereotactic frame. To keep the mouse's head immobile, align the ear bars properly and install them quickly. Gently tighten the ear bars so that they support the centered head and secure the mouse slightly.
2. Use a thermostatic pad to maintain the animal's body temperature at 37 ± 0.5 °C.
3. Use forceps to grasp the skin anterior to the mouse ears and cut it with sterilized scissors to remove about 1-cm² area of skin on the top of the mouse's skull.
   NOTE: A small amount of bleeding could be observed along the edges of the cut skin.
4. Remove the muscle and other attachment overlying the skull with a cotton swab to expose the skull surface. Use a cotton swab and dissecting chisel to clean all periosteum, tendons and connective tissue from the skull’s surface completely to identify the sagittal and lambda sutures lines.
5. Identify the bregma and lambda point. The intersection of the coronal suture and the sagittal suture on the superior middle portion of the calvaria is where the bregma is located. The intersection of the lambdoid suture by the sagittal suture is where the lambda is visible. Use the bregma as the original point to set the other nucleus position.
6. After the identification of the bregma and lambda position on the skull, adjust these two points in the horizontal level. Make sure that the height of these two points is almost the same in the dorsal-ventral direction (any errors should be < 0.05 mm). Also, make sure that the skull is level in the medial-lateral direction in the same way²⁶.
7. For a unilateral surgery, mount a homemade 2-channel copper micro-wire electrode into the rotating electrode holder on the stereotactic surgical frame.
8. Place the electrode vertically above the bregma point at first, and set it as original site. Use the digital displayed stereotactic surgical frame to indicate the anterior-posterior (AP), medial-lateral (ML), and dorso-ventral (DV) positions.
   NOTE: To prevent the electrode from getting bent, do not touch the electrode tip onto the skull during the operation.
9. Move the electrode gently to the correct position (Figure 1A, AP-2.1/ML ± 1.0 for DG and AP-3.8/ML ± 3.0 for PP, respectively)²⁷ above the skull without any touch. When the desired site was identified for electrode insertion, use the dorsoventral stereotactic adjustments to higher the electrodes holder, and mark the position with a black marker.
   NOTE: The exact location in reference to the stereotaxic coordinates may vary by mouse strain, weight, and sex. Adult mice of the same sex should be used to minimize any variation in the location. If possible, pre-operational anatomical scans should be used to identify the location on an individual subject basis²⁷.
10. Use a hand-held micromanipulator-assisted drill (burr size = 0.5 mm) to make burr holes at the marked position precisely. Sterilize the tip of the burr with 70% ethanol before drilling. Move up the mini drill bit through the skull along the Z-axis 0.8 – 1.0 mm without changing its X-Y position. Use a sterile cotton to stop any bleeding during the operation.
    NOTE: To avoid over-drilling and excessive heat generation during the drilling, ensure the drilling speed is 15,000 – 18,000 revolutions per minute (RPM), and frequently withdraw the drill bit from burr hole every 8 – 10 s.
11. Keep drilling until the dura is exposed. Use a bent needle to remove any bone scraps that are generated during the drilling and make a pinhole on the dura without damaging the underlying soft brain tissue.
    NOTE: To avoid damaging the brain tissue, the bent blunt needle can also be replaced with a fine blunt forceps.
12. To confirm that the burr holes have been made properly at the desired position, reduce the electrode height by an adjustment of the stereotactic surgery frame and ensure that the electrode can be inserted smoothly through the burr hole without touching any obstacles. If so, slowly insert the electrodes to a depth of 1.8 mm for the DG (or 3.0 mm for the PP) from the surface of the skull located at the medial PP to the DG (PP-DG).
    NOTE: Wipe off any cerebrospinal fluid or blood outflows from the burr holes with sterile cotton during the process of the electrode implantation.
13. After inserting the micro-wire electrode into the skull hole, hold it in place with adequate dental cement using a small spatula. As the cement thickens, mold it around the inserted electrode and screws to form a nice smooth cap. Debride and disinfect the wound edges.
14. Disengage the electrode from the electrode holder. Remove the mouse from the stereotactic frame and return it to a warm cage.
    NOTE: After surgery, wait with returning the animals to their cages until they have fully recovered from the anesthesia with sufficient consciousness.
15. Let the mouse move freely and allow it to recover for 2 days in the cage. Then, connect the electrode implanted in the mouse brain to a programmable stimulator via leads. Set up the software interface as presented in Figure 2. Set the HFS parameters as 6 series of 6 trains of 6 pulses at 400 Hz (Figure 1B, 100 ms between the trains, 20 s between the series)²⁸. Set the stimulation intensity to 200 μA.
16. Deliver an HFS for the desired period of time as set up. Deliver the stimulation 2x a day at 8:00 a.m. and at 8:00 p.m.
17. For the control group, implant electrodes in the mice and do not perform a stimulation.
18. For the c-fos staining, stimulate the experimental mice acutely (2x per day). For the BrdU labeling, divide the experimental mice into an acute stimulation group (applied 2x per day) and a chronic stimulation group (applied 10x in 5 days, 2x per day). During the course of the stimulation, make sure the mouse is awake. Remember to ensure the leads are not twisted.
    NOTE: At the last day of the procedure, the mice in the acute stimulation group were stimulated simultaneously.
19. When the stimulation is done, carefully disconnect the electrode pin from the electrode holder and the connector of the recording system.

3. Immunofluorescence Staining and Bromodeoxyuridine Labeling²⁹

1. For the c-fos and Notch1 staining, about 3 h after the last HF-DBS stimulation, anesthetize the mice by injecting pentobarbital (20 – 50 mg/kg, I.P.) to achieve a surgical plane of anesthesia.
2. For the BrdU labeling, about 12 h after the last HF-DBS stimulation, give 6 injections of BrdU to the control and the stimulated mice at 2-h intervals (50 mg/kg in 0.9% saline, I.P.). 36 h after the last injection, anesthetize the mice by injecting pentobarbital (20 – 50 mg/kg, I.P.) to achieve a surgical plane.
3. Make incisions using a scalpel through the rib cage to expose the heart and then pass a 22-gauge blunt perfusion needle to the left ventricle. Make a small incision in the right atrium. Transcardially perfuse it with 50 mL of cold phosphate-buffered saline (PBS) followed by 50 mL of cold 4% paraformaldehyde (PFA) in 1x PBS (pH 7.4).
   NOTE: Switch the PBS to the PFA when the mouse blood in vivo is replaced and the fluid runs clear. A good perfusion can be judged by the clearing of the liver. The perfusion should be stopped once fixation tremors have been observed³⁰.
4. Decapitate the mouse with ophthalmic scissors, make an incision on the skin and expose the skull. Using forceps, chip off the skull slightly and carefully dissect the whole mouse brain. Post-fix the brain in 4% paraformaldehyde for an additional 2 days and transfer it to a 30% sucrose solution at 4 °C overnight.
5. Using a cryostat, cut the brains into 20-µm coronal sections. Transfer and mount the sections to the glass slide with a brush and store them at -20 °C.
   NOTE: Animals with electrode misplacements or excessive mechanical damage will be excluded from the subsequent analysis.
6. Transfer the slides into normal PBS (0.1 M, pH 7.4) containing no fixative. Wash the slices with PBS for 3 – 4x for 5 min each to remove any excess fixative.For the BrdU-labeled group, incubate the sections in 2-N HCl for 30 min at 37 °C, and rinse them in a 0.1-M borate buffer (pH 8.4; 10 min). Then wash the slices 2x for 5 min in PBS.
   NOTE: Immunostaining is then performed.
7. Treat slices by incubation in blocking solution (1 % normal goat serum, 3 % bovine serum albumin <BSA> , 0.5 % Triton X-100 and 0.02 % sodium azide in 1 x PBS) for 1 h at room temperature (RT).
8. Incubate the slices with anti-c-fos polyclonal antibody (1:500; rabbit), anti-Notch1 antibody (1:50; goat), or anti-BrdU antibody (1:800; rat) in the blocking solution at 4 °C overnight.
9. Wash the slices 3x in PBS, then incubate them with Alexa Fluor 568-conjugated anti-rabbit (1:500), Alexa Fluor 488-conjugated anti-goat (1:1,000), or Alexa Fluor 488-conjugated anti-rat (1:500) secondary antibody for 1 h at RT.
10. Wash the samples 3x in PBS (10 min each time). Cover the immuno-labeled brain sections using an anti-fade reagent mounting medium containing DAPI. Let the slices air-dry and store them protected from light at 4 °C.
11. Acquire images of the hippocampal areas from the stimulated and unstimulated sides of the brain with a high-resolution multi-channel (sequential) scanning confocal microscope. Set the imaging parameters consistently between the control and experimental groups.
12. Perform a mosaic imaging using a high-magnification objective (20X air objective, NA 0.45) to acquire continuous 3D (XYZ) images of adjacent fields of view using the motorized stage and utilize appropriate software to stitch them together to make large composite images.
    NOTE: Tiling functions include true stitching and smoothing options for improved seamless images. Composite images are quickly and easily prepared using the stitching function, to form an image over a wide area.
13. For c-fos and Notch1 staining, perform a positive nuclei quantitative analysis of the immunofluorescent density²¹ of the stitched images. Randomly select 3 areas in the rostral, dorsal, and ventral hippocampal DG subregions and measure the immunofluorescent density in a double-blinded manner using Image J software (http://rsb.info.nih.gov/ij/)³¹.
14. For a BrdU labeling, analyze the slices around the drilling point through the dorsal hippocampus (-1.7 mm to -2.7 mm relative to the bregma) in a double-blinded manner.
15. Count only BrdU-positive neurons in the DG. Include the cells lying within 2-cell diameters of the granule cell in the count.
    NOTE: Here, counting was performed using a 40X air objective. Randomly, 3 – 5 sections per animal were counted, and the counts were averaged and expressed as means per DG¹⁹.