Long-term Potentiation of Perforant Pathway-dentate Gyrus Synapse in Freely Behaving Mice

This protocol is appropriate for mice of 3 and 18 months of age and approximate body weight of 30-50 g). Mice can be obtained from The Jackson Laboratory (Bar Harbor, ME). All surgical and experimental protocols were approved by the Trinity College Animal Care and Use Committee and were in accordance with the NIH Guide for the Care and Use of Laboratory Animals.

1. Animal Preparation and Surgical Procedures

1. Prepare anesthesia cocktail containing ketamine (25 mg/ml), xylazine (2.5 mg/ml) and acepromazine (0.5 mg/ml). Inject 1 ml/kg body weight in the lower right quadrant of the abdomen supplemented by 0.2 ml/kg injections every 45 min thereafter to maintain a stable depth of anesthesia using the pedal withdrawal reflex.
2. Prepare anesthetized mouse for surgery by shaving the fur on its cranium using an electric shaver. The shaved area should include the middle of the eyes to the middle of the ears. Make sure not to shave the whiskers as they are part of the mouse barrel sensory system.
3. Use cotton swabs to apply isopropyl alcohol followed by Betadine to the shaved area on the head. Then, also using cotton swabs, apply a small amount of mineral oil on the eyes to prevent the eyeballs from drying.
4. Use infant rat ear cuffs instead of regular ear bars to mount the animal’s head on the stereotaxic frame apparatus. To do this, first mount the left ear on the left ear cuff. Then gently ease the right ear onto the right cuff by gently supporting the animal’s head with one hand and advancing the ear cuff with the other hand. Next, place a heating pad under the animal’s body and set it to 86 °F to maintain body temperature.
5. Then, check for bilateral rigidity by attempting to move the head side to side. The animal is properly mounted on the stereotaxic frame when the head is rigid and cannot be moved side to side but can easily pivot up and down.
6. Gently rest the mouse’s snout on the nose/tooth bar assembly making sure that the incisors are resting gently but securely within the tooth aperture of the nose/tooth bar.
7. Using a sterile scalpel, make a midline incision starting from the middle of the eyes to the middle of the ears. Make sure the scalpel is be held at a 45° angle and apply enough pressure to cut through the underlying fascia but not through the skull as that will cause excessive bleeding.
8. Use cotton swabs to separate the fascia and expose the skull. Skull landmarks bregma and lambda should be clearly visible (Figure 1A). Ensure that the animal is in the skull flat position by adjusting the nose/tooth bar such that bregma and lambda landmarks are at same the dorsoventral (DV) position (+ 0.1 mm).
9. Apply a small amount of sterile saline to the exposed area and use cotton swabs to gently clean the skull. Then wait 2-3 min for the skull to completely air dry before proceeding.
10. Mount a needle on the left stereotaxic arm to measure the DV position of bregma and lambda. The DV positioning of these two landmarks should be within 0.1 mm of each other. If not, the nose bar of the stereotaxic frame can be adjusted up and down to achieve this.
11. Position the needle on bregma to record its anterior-posterior (AP), lateral (LAT) and dorsoventral (DV) measurements. Be sure the needle just touches the skull and does not penetrate it.
12. Use a mouse brain atlas, such as “The Mouse Brain in Stereotaxic Coordinates”¹, to determine the coordinates for the target structures: medial perforant pathway (mPP) in the angular bundle and dentate gyrus (DG) of the hippocampus, relative to lambda and bregma, respectively (also see Table 1). Then, use a fine-point pen or pencil to mark these points on the skull.
13. Also, mark two more points on the contralateral side of the skull. These points which will serve as ground (GND) and reference (REF) should be about 3 mm from the midline and positioned longitudinally relative to each other (Figure 1A, also Table 1).
14. Remove the needle marker from the left stereotaxic arm and replace it with an electric dental drill equipped with a stereotaxic mount. Position the drill over each marked point and make small burr holes of approximately 0.5 mm in diameter. When drilling, use a repetitive up-and-down motion and check frequently whether the drill has penetrated the skull to expose the brain surface.
15. Gently but firmly drive a screw electrode (#0-80 1/8 in stainless steel machine screws, slotted fillister head) in each of the contralateral holes (GND and REF) such that they just touch the cortical surface without penetrating it (Figure 1A). These two screw electrodes serve as ground and reference.
16. Mount stimulating and recording electrodes in electrode holders on the left and right stereotaxic arms, respectively.
17. Connect the stimulating electrode terminals to an electrophysiological stimulator with current isolation and the recording electrode terminal to an electrophysiological differential amplifier (gain = 1,000, bandpass filter = 1 Hz–3 kHz) and then to a digital oscilloscope for visual inspection of evoked responses.  Also, connect GND and REF electrode terminals to the differential amplifier.
18. Gently and slowly lower both stimulating and recording electrodes in 0.5 mm increments while visually monitoring the evoked response to a stimulus shock of 600-800 uA. Continue to lower each electrode until they reach their respective target DV position and a stereotypical signal is observed on the oscilloscope (Figure 1B).
19. Afterward, use dental acrylic cement to make a cap to hold the electrode terminals in place; any dental cement that touches the skin should be wiped off immediately.  Once the dental cement is completely cured transfer the animal from the stereotaxic frame to a clean rodent cage. Use a heating lamp or pad to maintain core temperature.
20. Monitor the animal every hour until it regains consciousness. An injection of flunixin may be administered as an analgesic.

2. LTP Induction

1. Allow the animal 5-7 days to recover from surgery. Place the animal in the recording environment consisting of a faraday cage (122 cm x 43 cm x 43 cm) outfitted with soundproofing material and a 5-channel rotating commutator connecting the electrode leads to the stimulating/recording setup.
2. Allow the animal to acclimate for 1-2 hr in the recording environment before connecting the electrodes to the stimulating and recording instruments (refer to step 1.17).
3. Set the stimulator controls to output 400 uA and record the amplitude of an average of 10 evoked responses making sure that at least 10 sec elapse between stimuli. Use the method shown in Figure 1C to quantify the evoked response. Repeat this procedure for 600, 800, 1,000, 1,200, and 1,400 uA.
4. Construct an input/output curve by plotting the average amplitude of the evoked response versus stimulus intensity. From this plot determine the stimulus intensity which corresponds to 50% of the maximum amplitude measured. Use the 50% intensity for the remainder of the experiment.
5. Then, using the 50% stimulus intensity, obtain a baseline by recording the average amplitude of 5 evoked responses every minute for 15 min. Again make sure that at least 10 sec elapse between stimuli.
6. Subsequently deliver the tetanic stimulation consisting of 10 bursts of 10 pulses delivered at 400 Hz with a burst rate of 5 Hz to the medial perforant pathway. Monitor the animal closely for signs of seizure, including wet dog shakes. If the animal has a seizure the experiment should be terminated and all data from that animal should be excluded from the study results.
7. Then, continue to record the average amplitude of 5 evoked responses every minute for 30 min posttetanization. After that, compare this amplitude to the baseline amplitude obtained in step 2.5 by calculating the percent change from baseline (Figure 2).
8. Following the conclusion of experiments, the animal is euthanized by means of inhalation of isoflurane—a method which is consistent with the Guidelines on Euthanasia of the American Veterinary Medical Association.