All animals were treated in accordance with the guidelines and regulations from the Animal Care and Use of Laboratory of National Institute of Health. All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of City University of Hong Kong and Incheon National University.

1. In vivo field recording

1. Animal preparation
   1. Inject urethane (0.06 g per 25 g weight) intraperitoneally to anesthetize mouse. Supplement with intramuscular injection of atropine (0.05 mg/kg). Keep the mouse in a dark quiet spot until full anesthesia takes effect.
      CAUTION: Urethane has carcinogenic potential. Handle it with care and wear protective clothing to avoid contact with exposed skin.
      NOTE: Alternatively, isoflurane can be used as anesthesia.
   2. Check for depth of anesthesia intermittently until a full surgical plane of anesthesia takes effect. Check for depth of anesthesia by performing toe pinch, ear pinch, tail pinch and corneal touch tests to observe the response of the mouse to physical stimuli.
      NOTE: A reflex or voluntary movement should not be observed when the mouse is at a full surgical plane of anesthesia.
   3. For the toe pinch test, extend either the hind or foreleg of the mouse and pinch firmly with a pair of blunt forceps or fingers. The mouse is not confirmed for full anesthesia if it withdraws the leg or shakes body, has an observable increased respiratory rate or makes vocal sounds.
   4. For the ear pinch test, pinch the ends of the pinna with a pair of blunt forceps or fingers firmly. Full anesthesia is not confirmed if the mouse moves the whiskers forward, shakes its head, makes vocal sounds or has an observable increased respiratory rate.
   5. For the tail pinch test, hold the tail of the mouse gently and firmly pinch with a blunt forcep or finger. No tail movement, vocal sound or observed increase in respiratory rate should be observed when fully anesthetized for surgical procedures.
   6. For the corneal touch test, touch the cornea of the mouse gently, with a cotton wick. No eyelid movement, whisker movement or observable increase in respiratory rate should be observed when fully anesthetized for surgery.
   7. Shave the hair on the neck and skull of the mouse.
   8. Place the fully anesthetized mouse on a heating pad set to 37 °C and insert the rectal temperature probe in the rectum. This enables the heat produced by the heating pad to adjust in response to changes of the mouse’s body temperature.
   9. Apply eye gel to moisten the eyes of the mouse.
   10. Pull the tongue out to the side of the lips gently using forceps and fix the two front teeth into the second or third teeth hole of the stereotactic instrument. Fix the skull of the mouse firmly using the eye-clamp.
       NOTE: Alternatively, a stereotactic ear clamp can be used.
   11. Observing under a microscope, separate the subcutaneous tissue and muscles at the end of the interparietal and occipital bone of the mouse with a scalpel to expose the cisterna magna. Blot the dura mater dry with a cotton swab.
   12. Gently puncture the cisterna magna by making a shallow cut with a sharp pointed scalpel blade to drain the cerebrospinal fluid (CSF). Attach a cotton swab to keep draining the CSF.
2. Craniotomy
   1. Holding the skin on the scalp with forceps, cut and remove the skin with a pair of surgical scissors. Cut enough skin to expose the bregma and lambda marks on the scalp. Keep the exposed region dry.
   2. Adjust the clamped skull of the mouse to enable the bregma and lambda points to be aligned in a horizontal level of similar height. Avoid tilting of the head in either the anterior-posterior position or the medial lateral position.
   3. Mark the points corresponding to the hippocampal region using the aid of a Vernier caliper. Use the mouse brain in a stereotactic coordinate book as a reference to help determine the exact coordinates.
      NOTE: The location to mark for incision for the hippocampus are 1 mm x 3 mm on the midline referred to as anterior-posterior (AP) using the bregma as reference point, and 3 mm x 3 mm perpendicular to the midline (ML) to connect the points on the midline. The stereotactic coordinates are given as (AP: 1,3 and ML: 3,3).
   4. Make an incision in the skull above the dorsal region of the hippocampus along the marked points using a scalpel or high-speed drill while observing under a microscope.
      NOTE: A rectangular shaped opening with a size of 2 mm x 3 mm should be obtained after incision. Keep the exposed area clean to avoid injury to the brain.
   5. Carefully take out the loose skull with forceps to expose the dura mater and use a syringe or dropper to gently apply physiological saline solution to keep the surface moist.
   6. Remove the dura mater carefully with needle or sharp pointed tip forceps.
      NOTE: Be careful not to cause any damage to the brain tissue during craniotomy. This will lead to the swelling of the brain and will affect the results.
   7. Keep the exposed brain tissue moist by applying physiological saline or inert oil using a dropper or syringe.
3. In vivo recording
   1. Fix and position the stimulation and recording electrode firmly in the stereotactic holder. Adjust the stereotactic instrument according to the position of the corresponding AP and ML coordinates for the stimulation and recording electrodes above the CA1 dorsal hippocampus.
      NOTE: Use the mouse brain in stereotactic coordinates as a guide in locating the coordinates for the dorsal longitudinal CA1 hippocampal region. For example, a stereotactic coordinate for the CA1 hippocampal region will be (AP 1.5, ML 1.0) for the recording electrode and (AP 1.7, ML 1.5) for the stimulation electrode.
   2. Locate the stimulating electrode lateral to the recording electrode in a longitudinal direction.
      NOTE: Ensure that both stimulation and recording electrodes are clean before usage. This prevents the introduction of noise when recording.
   3. For recordings, use multichannel electrodes. First, locate the stereotactic coordinates for the CA1 hippocampal region using the first channel. Position the remaining electrodes such that the angle of stimulation and recording electrodes are in the range of 30° to 60° in relation to the midline from bregma point.
      NOTE: This angle corresponds to the longitudinal orientation of the CA1 hippocampal region.
   4. As a control, locate the recording electrode above the CA1 region and the stimulation electrode above the CA3 region of the dorsal hippocampus. For example, using the mouse brain in stereotactic coordinates as a guide, a stereotactic coordinate for CA1-CA3 hippocampal region will be (AP 1.8, ML 1.0) for the recording electrode and (AP 1.5, ML 1.5) for the stimulation electrode.
      NOTE: This step is an alternative control and should be done in a separate experiment.
   5. Place the reference electrode at a distal part of the exposed brain region or under the skin of the mouse.
   6. Turn on the recording system.
   7. Open the software for recording and data acquisition.
      NOTE: Different laboratories have their preferred software for recording and data acquisition.
   8. Observing under a microscope, lower the recording and stimulation electrodes slowly using the micromanipulator until it just touches the surface of the brain. Mark the point as the zero point to start calculating the accurate depth to the hippocampal region.
      NOTE: The micromanipulator can be used to monitor the depth at which the electrode is inserted at any given time. Observe an increase in the impedance just when the electrode touches the brain surface.
   9. Slowly lower the electrodes to the approximate depth corresponding to the chosen stereotactic coordinates for CA1 hippocampus.
      NOTE: Use the mouse brain in stereotactic coordinates as a guide to obtain the approximate depth corresponding to the chosen stereotactic coordinates.
   10. Give stimulation (100 μs duration, repeated at 30 s intervals) and adjust the electrode depth in steps of 50 μm or less until a stable evoked field excitatory postsynaptic potential (fEPSP) is observed.
       NOTE: A stimulus of 20 μA is usually enough to evoke an observable response.
   11. Ensure that a stable fEPSP has been evoked by varying the stimulus intensity. A notable increase in the slope or amplitude of the fEPSP should be observed with each increased stimulus intensity. This is termed as the input-output curve. Create an input-output (I-O) curve to detect maximum stimulus intensity at which there is no more increase in the slope of the evoked fEPSP (Figure 6).
       NOTE: Discard data and change electrode position if the fiber volley disappears during experiment.
   12. Use the input-output curve to set the baseline intensity to 40 – 50% of the maximum. Use the corresponding stimulus intensity for baseline recording.
   13. Record the local field potential as a baseline for 20 – 30 min.
   14. Use the same input-output curve to set the stimulus intensity for evoking high frequency stimulation (HFS) or tetanus to 75% of the maximum intensity. Alternatively, when inducing long-term depression, maintain the same stimulus intensity used for recording the baseline when evoking low frequency stimulation (LFS).
   15. Apply a tetanic stimulation of 100 Hz pulses 4 times with a 10 s interval to induce LTP.
       NOTE: Use this protocol only when working on a long-term potentiation experiment.
   16. Apply low frequency stimulation of 5 Hz (900 stimuli during 3 min), 1 Hz LFS (900 stimuli during 15 min), or 1 Hz paired-pulse (50 ms paired-pulse interval, 900 pairs of stimuli during 15 min) to induce LTD according to already established protocols.
       NOTE: Use these protocols only when working on a long-term depression experiment.
   17. Record the local field potential for 1 h after HFS or LFS, alternatively.
   18. Export data and analyze using the software.
   19. Verify the position of the recording and stimulation electrode by giving a stimulation of 10 μA current for 30 s to lesion the recorded areas. Transcardially perfuse the mouse with 4% paraformaldehyde and harvest the brain for slicing and staining with Cresyl Violet according.
   20. Euthanize mouse by cervical dislocation or injection of lethal anesthetic dosage after experiment.

2. In vitro field recording

1. Preparing oxygenated slicing and artificial cerebrospinal fluid (ACSF) solutions
   1. For 2 L of slicing solution, add approximately 1 L of double distilled water in a volumetric flask and stir vigorously on a stirrer plate.
   2. Add the following slicing solution components (in mM): 87.0 NaCl, 2.5 KCl, 1.3 NaH₂PO₄, 25.0 NaHCO₃, 25.0 glucose, 75.0 sucrose, 7.0 MgCl₂.6H₂O and 0.5 CaCl₂∙2H₂O (Table 1).
      NOTE: Alternatively, MgCl₂∙6H₂O and CaCl₂∙2H₂O can be excluded in this stage and added later in a ready to use volume.
   3. Top up to 2 L with double distilled water while stirring vigorously.
   4. For 2 L of ACSF, add approximately 1 L of double distilled water in a volumetric flask and stir vigorously on a stirrer plate.
   5. Add the following ACSF solution components (in mM): 125.0 NaCl, 2.5 KCl, 1.3 NaH₂PO₄, 25.0 NaHCO₃, 25 glucose, 1.0 MgCl₂∙6H₂O and 2.0 CaCl₂∙2H₂O (Table 1).
      NOTE: Alternatively, MgCl₂.6H₂O and CaCl₂∙2H₂O can be excluded in this stage and added later in a ready to use volume.
   6. Top up to 2 L with double distilled water while stirring vigorously.
2. Setup and brain slicing
   1. Pour 400 mL of already prepared slicing solution into a separate flask and oxygenate (95% O₂/5% CO₂) for approximately 20 min.
   2. Store the rest of the 1.6 L solution in a 4 °C refrigerator. Keep solution up to 1 week after which it must be discarded if not used to avoid fungal growth.
   3. Pour 200 mL of the oxygenated slicing solution in the flask, cover with parafilm and transfer to a -80 °C freezer for approximately 20 min to make a slush.
   4. Pour the remaining 200 mL of slicing solution in a brain slice holding chamber and keep in a 32 °C water bath with continuous bubbling.
   5. Prepare the bench for slicing. Place paper towels down and surgical tools on top of bench. Arrange the surgical tools in order of use to facilitate a fast and efficient process (Figure 7).
   6. Take out the chilled slicing solution from the freezer and pour approximately 50 mL in a beaker. Pour approximately 10 mL in a Petri dish containing filter paper to moisten it. Place them on ice by the dissection area.
   7. Pour the rest of the slushed slicing solution into the slicing chamber and fix it on the vibratome.
   8. Anesthetize the mouse with isoflurane using guidelines and regulations on Animal Care and use of laboratory of National Institute of Health, and the approved methods of Institutional Animal Use and Care Committee of City University of Hong Kong and Incheon National University.
   9. Decapitate the anesthetized mouse with a pair of scissors and place the head on tissue paper. Cut the skin covering the skull of the mouse and cut through the cutaneous muscles with scissors. Cut the skull plates along the midline to the occipital bone using surgical scissors.
   10. Open the skull with blunt forceps to expose the brain. Gently scoop out the brain with a spatula and place in the chilled slicing solution in the beaker. Wait about 30 s.
   11. Take out the brain using the spoon and carefully place it on the previously moistened filter paper in the Petri dish.
   12. Separate the two brain hemispheres along the midline with a scalpel blade. Isolate the hippocampus by gently detaching it from the cortex with a spatula and place it carefully on the moistened filter paper. Cut out the septal and temporal end of the isolated hippocampus using the scalpel.
   13. Pick up the isolated hippocampus using a blunt spatula and brush. Gently dab the spatula on a tissue paper to remove excess water.
   14. Apply a small amount of glue to the slicing plate of the vibratome.
   15. For a longitudinal CA1 hippocampal slice, attach the CA3 region of the hippocampus to the slicing plate with the glue. Quickly but carefully place it in the slicing chamber of the vibratome containing the chilled oxygenated slicing solution.
   16. For the transverse slice that will serve as a control, attach the ventral end of the hippocampus to the slicing plate of the vibratome. Quickly but carefully place it in the slicing chamber containing the chilled oxygenated dissection solution.
       NOTE: This step is an alternative to the step above and should be performed separately.
   17. Position the blade angle to 90°. Set the vibratome parameters to a speed of 0.05 mm/s, an amplitude of 1.20 mm and a slice thickness of 400 µm.
   18. Slice the attached hippocampus with the vibratome.
       NOTE: A good longitudinal CA1 hippocampal brain slice will have one layer of CA1 and 2 layers of dentate gyrus. A maximum of 2 good hippocampal slices can be obtained. Alternatively, a transverse hippocampal brain slice will have the dentate gyrus, CA3 and CA1 regions intact.
   19. Transfer the longitudinal hippocampal brain slices from the vibratome with a pipette and incubate it in the brain slice holding chamber in the water bath.
   20. Incubate slices in the water bath for 20 min at a temperature of 32 °C and bring out to room temperature for 30 min of recovery.
       NOTE: Alternatively, transfer the brain slice when out of the water bath and gently place it in the recording chamber with flowing oxygenated ACSF at a temperature of 32 °C for 30 min of recovery.
   21. While incubating in the water bath, pour 400 mL of ACSF solution into a separate flask and oxygenate (95% O₂/5% CO₂) for approximately 20 – 30 min before transferring the brain slice into the recording chamber. Continuously superfuse the ACSF into the recording chamber at a speed of 2 mL/min.
   22. Turn on the temperature controller to heat the flowing ACSF to 32 °C.
   23. Store the rest of the 1.6 L solution in a 4 °C refrigerator. Keep the solution up to one week after which it must be discarded if not used to avoid fungal growth.
3. In vitro recording
   1. Set up a recording rig by turning on all needed hardware.
   2. Transfer the brain slice from the slice holding chamber into the recording chamber. Adjust the position of the brain slice with blunt forceps and hold it in place with a harp. Allow the ACSF to run for at least 20 min. This enables the brain slice to be stable before recording.
   3. Fill the recording pipette with ACSF as an internal solution.
   4. Check the recording pipette resistance with the designated software. The recording pipette resistance should be within the range of 3-5 MΩ.
   5. Turn on the software for data acquisition. This software should have similar features that enable data acquisition in a similar way as the in vivo recordings shown earlier.
   6. Fix the recording and the stimulation electrodes firmly in the stereotactic instrument holder. Place the reference electrode in the ACSF in the recording chamber.
   7. For longitudinal slices, position the stimulation and recording electrode in the stratum oriens (S.O.). Keep the distance between electrodes to about 300 to 500 µm. Place the stimulation electrode on either the septal or temporal side of the brain slice and record from the same layer (Figure 8).
   8. Alternatively, position the stimulation and recording electrode in the stratum radiatum (S.R.). Place the stimulation electrode on either the septal or temporal side of the brain slice.
   9. For transverse slices as a control, position the stimulating electrode on the CA3 (Schaffer collateral pathway) region, and the recording electrode on the CA1 region (Figure 9).
      NOTE: This is an alternative control step and should be performed in a separate experiment.
   10. Turn on the isolated stimulus generator and give stimulation (100 μs duration, repeated at 30 s intervals). Adjust the recording electrode depth and/or position till a stable evoked excitatory postsynaptic field potential is observed.
   11. Ensure that a stable fEPSP has been evoked by varying the stimulus intensity. A notable change in the slope of the fEPSP should be observed with each change of stimulus intensity. This is termed as the input-output curve. Create an input-output (I-O) curve to detect maximum stimulus intensity at which there is no more increase in the slope of the FEPSP (Figure 6).
       NOTE: Discard data and change electrode position if the fiber volley disappears during experiment.
   12. Use the input-output curve to set the stimulus intensity for baseline recording and for evoking high frequency stimulation (HFS) or tetanus to 30-40% of the maximum evoked fEPSP.
   13. Alternatively, for experiments regarding LTD, use the input-output curve to set the baseline intensity for baseline recording and for evoking low frequency stimulation (LFS) to 70% of the maximum evoked fEPSP.
   14. Record the local field potential as the baseline for 20 – 30 min.
   15. Apply HFS of 100 Hz pulses twice with a 30 s interval to induce LTP.
   16. Alternatively, for LTD experiments, apply low frequency stimulation of 5 Hz (900 stimuli during 3 min) or 1 Hz LFS (900 stimuli during 15 min) or 1 Hz paired-pulse (50 ms paired-pulse interval, 900 pairs of stimuli during 15 min) to induce LTD according to already established protocol.
   17. Record the local field potential for 1 h after HFS or LFS.
   18. Export data and analyze.