Slice Patch Clamp Technique for Analyzing Learning-Induced Plasticity

All animal housing and surgical procedures were in accordance with the Guidelines for Animal Experimentation of Yamaguchi University School of Medicine and were approved by the Institutional Animal Care and Use Committee of Yamaguchi University.

1. Animals

1. Use 4- to 5-week-old male Sprague-Dawley rats (postnatal 28 to 31 days of age).
2. House the rats in individual plastic cages (40 cm × 25 cm × 25 cm) maintained at a constant temperature (23 °C ± 1 °C) under a 12-h light/dark cycle. Give rats ad libitum access to water and food.

2. Rotor rod test

1. To investigate motor skill learning, subject each rat to the rotor rod test (rod diameter 7 cm; lane width 8.9 cm; fall height 26.7 cm) for 1 or 2 consecutive days (Figure 1A in Kida et al., 2016⁷). Perform the task in a quiet, temperature-controlled room (23 ± 1 °C). Do not disturb or handle rats prior to the test.
2. Set the rotor rod to acceleration mode, which increases linearly from 4 rotations/min to 40 rotations/min (8 π/min to 80 π/min) in 5 min.
3. Put the rat on the resting rotating rod. Confirm that all limbs are on the rod.
4. Measure the latency to fall from the rotating rod to evaluate motor performance.
5. Allow each rat 10 attempts (trials) with 30-s intervals.
6. If the rat falls from the rotating rod, set it on the rod again after a 10-20 s interval.
7. Sacrifice the rat with an overdose of pentobarbital (400 mg/kg) 30 min after the final trial. Inject untrained control rats with the same dose of anesthesia in their home cages.

3. Inhibitory avoidance test

1. To investigate contextual learning, subject rats to an inhibitory avoidance (IA) test (Figure 1D in Mitsushima et al., 2011, 2013^10,11) Avoid any episodic experiences on the day of the experiment such as contact with others, cage changes or cleaning. Perform the task in a quiet, temperature-controlled room (23 ± 1 °C).
   NOTE: The IA training apparatus is a two-chambered acrylic box (length 33 cm; width 58 cm; height 33 cm). It has a lighted safe side and a dark shock side that are separated by a trap door (Figure 1D).
2. Place the rat in the safe (lit) side of the illuminated box. Handle the rat gently without stress.
3. Wait a short time (10 to 20 s) to acclimate the rat to the environment.
4. Open the sliding door to allow the rat to enter the dark box at will.
5. Measure the latency (s) before the rat enters the novel dark side of the box. The latency of the first trial represents the rat's performance before training.
6. After entry into the dark side, close the door and apply a scrambled electrical foot shock (2 s, 1.6 mA) via electrical steel rods set into the floor of the box. Allow walk-through rats to explore the training apparatus for 1 min without being shocked. House unpaired rats in an illuminated shock cage for several days and suddenly give the shock without episodic experiences. Handle gently without stress in any groups.
7. Keep each rat in the dark box for 10 s before returning it to the home cage.
8. At 30 min after the foot shock, again place the rat into the lighted side of the box. Measure the latency to enter the dark side.
9. Return the rat to the home cage.
10. At 60 min after the shock, sacrifice the rat with an overdose of pentobarbital (400 mg/kg). Handle the rat gently and inject the anesthesia intraperitoneally. In untrained control rats, inject the anesthesia in their home cages without the experience described above.

4. Dissection buffer

1. Dissolve crystals of 0.195 g NaH₂PO₄-2H₂O, 0.188 g KCl, 0.074 g CaCl₂, 1.423 g MgCl₂-6H₂O, and 12.579 g choline chloride in ultrapure water (900 mL to 950 mL). See Table 1.
2. Dissolve crystals of 2.340 g ascorbic acid, 0.342 g pyruvic acid sodium salt, 2.100 g NaHCO₃, and 4.500 g glucose.
3. Add water up to 1000 mL. The range of osmolality will be between 290 mOsm/L and 300 mOsm/L. Adjust osmolality by adding ultrapure water, if it is over the range.
4. Bubble the solution with 5% CO₂/95% O₂ gas mixture at ice-cold temperatures for 5 min before use.

5. Artificial cerebrospinal fluid (aCSF)

1. Dissolve crystals of 0.186 g KCl, 6.700 g NaCl, and 0.156 g NaH₂PO₄-2H₂O in ultrapure water (900 mL to 950 mL). See Table 2.
2. Bubble with the gas mixture for 5 min.
3. Dissolve crystals of 1.800 g glucose and 2.184 g NaHCO₃ and then add 4 mL MgCl₂ and 4 mL CaCl₂ from 1 M stock solutions.
4. Add water up to 1000 mL. The range of osmolality will be between 290 mOsm/L and 295 mOsm/L. Adjust osmolality by adding ultrapure water, if it is over the range.
5. Bubble with the gas mixture prior to use.

6. Intracellular solutions

1. For current-clamp recordings (Table 3), dissolve 0.0746 g KCl, 6.089 g K-gluconate, 0.476 g HEPES, 0.0456 g EGTA, and 500 µL MgCl₂ from 1 M stock solution in 180 mL ultrapure water (adjust pH to 7.2 with KOH).
   1. Add 0.4408 g Na₂-ATP, 0.0418 g Na₃-GTP, and 0.510 g Na-phosphocreatine. Add water to 200 mL and adjust the pH to 7.35 with KOH.
   2. Adjust the osmolality to around 290 mOsm/L by adding ultrapure water.
   3. Store as 1-mL aliquots in the freezer (-30 °C).
2. For voltage-clamp recordings (Table 4), dissolve 5.244 g CsMeSO₃, 0.672 g CsCl, 0.476 g HEPES, 0.0456 g EGTA, and 500 µL MgCl₂ from 1 M stock solutions in 180 mL ultrapure water. Adjust the pH to 7.2 with CsOH. For mEPSP and mIPSP recordings, use modified concentration of 5.814 g CsMeSO₃ and 0.252 g CsCl to adjust the reversal potential of the GABA_A receptor response¹¹.
   1. Add 0.4408 g Na₂-ATP, 0.0418 g Na₃-GTP, and 0.510 g Na-phosphocreatine. Add water to 200 mL and adjust the pH to 7.35 with CsOH.
   2. Adjust the osmolality to around 290 mOsm/L by adding ultrapure water .
   3. Store as 1-mL aliquots in the freezer (-30 °C).

7. Slice preparation

1. Prior to sacrifice, cool-down all dissection tools with crushed ice (Figure 2A). Add about 500 mL of cold water into the crushed ice container to increase the contact surface area. This procedure was described previously^10,11,12.
   NOTE: The tools here are: Large scissors, iris scissors, a spatula, a micro spatula, forceps, tweezers, a stainless steel 200-mL beaker, a blade for brain trimming, a 120-mL cardiac perfusion syringe filled with dissection buffer treated with the gas mixture, a silicone tube (20 cm) connected to a flattened 18-gauge needle, a stainless brain dissection stage (thickness = 3 mm, ϕ = 12 cm), and a mounting stage for the vibratome (ϕ = 5 cm).
2. Sacrifice the rat 30 min after completing the behavioral paradigm by anesthetizing it with an overdose of pentobarbital (400 mg/kg bodyweight). Perform the slice preparation rapidly to ensure that the slices are as healthy as possible^10,11,12. The brain extraction protocol meets all veterinary standards for our university.
3. Fill a 120-mL syringe with ice-cold dissection buffer (Table 1) bubbled with a 5% CO₂/95% O₂ gas mixture. Remove any air bubbles prior to perfusion.
4. After exposing the heart, insert the needle into the posterior part of the left ventricle.
5. Perform transcardial perfusion of the brain manually using the syringe. Larger rats require more dissection buffer for perfusion. Submerge the brain with ice-cold dissection buffer for 5 min. Bubble the buffer continuously during the submersion.
6. Trim the posterior side of the brain at an angle parallel to the dendritic orientation of the target cortical region using a blade. Since the brain is stand on dissection stage with the cut end bottom, the initial angle determines the angle of all subsequent brain slices. This step is critically important (Figure 2B). An incorrect angle may cut through the target pyramidal neurons.
   NOTE: The tools here are: a blade for brain trimming, a filter paper (ϕ = 10 cm), a stainless brain dissection stage (thickness = 3 mm, ϕ = 12 cm), a spatula, a superglue, a dropper, and a mounting stage for the vibratome (ϕ = 5 cm).
7. Cut 350-µm thick coronal brain slices using a vibratome. Fill the dissection chamber with ice-cold buffer bubbled with a 5% CO₂/95% O₂ gas mixture (Figure 2C). Bubble the buffer continuously during the brain slice.
8. Trim the periphery of the target area using iris scissors.
9. Wash the trimmed slices gently in room temperature aCSF bubbled with 5% CO₂/95% O₂ (Table 2).
10. Maintain the trimmed slices in an interface chamber until the recording is performed (Figure 2D and E). Incubation for 1 h in the chamber improves the condition of the cells, but the phenotypes change if the slices are incubated for more than 10 hours. Close the lid of the chamber to enclose the gases and the small liquid drops of aCSF.

8. Whole-cell patch clamp

NOTE: Whole-cell recordings require an amplifier and a low-pass filter that is set to a cutoff frequency of 5 kHz. The signals are digitized and stored in a PC. The stored data are analyzed offline (Figure 3A).

1. Create glass electrodes using a horizontal puller. Fill the electrodes with a suitable solution (Tables 3 and 4) using a regular polyethylene 1-mL syringe attached to a fine glass tube and a 0.22-µm filter.
2. Prior to contact with the cell, maintain positive pressure and adjust the pipette current to zero.
3. After forming a gigaohm seal, apply negative pressure to rupture the cell membrane (whole-cell configuration in Figure 3F).

9. Current-clamp analysis

1. Properties of the cell membrane
   1. Fill the patch recording pipettes with the intracellular solution for current-clamp recordings (Table 3). The resistance of the pipette is between 4 MΩ and 7 MΩ in the aCSF.
   2. After the membrane ruptures, hold the membrane voltage at -60 mV in V-CLAMP mode. Then, switch from "bath" mode to "cell" mode in the membrane test using software to measure the intrinsic cell properties such as membrane capacitance, resistance, and time constant.
2. Current injection study
   1. After recording the intrinsic cell properties, switch the mode from V-CLAMP to TRACK (I = 0) /I-CLAMP NORMAL for the current injection. Note that the liquid junction potential should not be corrected¹⁰.
   2. Inject current into the cell for 300 ms. Change the intensity of the current stepwise from −100 pA to +550 pA with 50-pA increases. Count the number of spikes (action potentials) elicited by the current injections.
   3. Measure the minimum voltage needed to induce an action potential (this is the threshold voltage).
   4. Calculate the afterhyperpolarization amplitude as the difference between the voltage at spike initiation and the lowest voltage attained during afterhyperpolarization⁷.

10. Voltage-clamp analysis

1. The AMPA/NMDA ratio
   NOTE: The AMPA/NMDA ratio is a conventional way to evaluate postsynaptic plasticity at glutamatergic excitatory synapses^7,8,9,10,11. However, note that concomitant increases in both components may not change the ratio¹³.
   1. Perfuse the recording chamber with physiological solution bubbled with the gas mixture and maintain the temperature at 22 °C to 25 °C. Add 0.1 mM picrotoxin to the solution to block the GABA_A-mediated response and add 4 µM 2-chloroadenosine to stabilize the evoked neural response¹⁴.
   2. Fill the patch recording pipettes with the intracellular solution for voltage-clamp recordings (Table 4). Check the resistance of the recording pipette in the aCSF. The resistance is between 4 MΩ and 7 MΩ.
   3. For recording in layer II/III pyramidal neurons in the M1, place a bipolar tungsten stimulating electrode 200 µm to 300 µm lateral to the cells to be recorded, below the pial surface in the region of the forelimb representation (2-mm lateral to the midline)^15,16,17.
   4. For recording in a CA1 pyramidal neuron, place the stimulating electrode 200 µm to 300 µm lateral (Schaffer collateral fiber) or medial (temporoammonic pathway) to the cells that will be recorded (Figure 3B).
   5. Increase the stimulus intensity up to the synaptic response > 10 pA.
   6. Calculate the AMPA/NMDA ratio as the ratio of the peak current measured at −60 mV to the current measured at +40 mV at 150 msec after the stimulus onset. Note that 50 to 100 traces should be averaged to calculate the ratio.
2. Miniature postsynaptic current recordings
   Note: Miniature excitatory postsynaptic currents (mEPSCs) are thought to correspond to the responses elicited by the presynaptic release of a single vesicle of glutamate¹⁸. In contrast, miniature inhibitory postsynaptic currents (mIPSCs) are thought to correspond to the responses elicited by the presynaptic release of a single vesicle of GABA¹⁸. Increases in the amplitudes of mEPSCs and mIPSCs reflects postsynaptic transmission strengthening, while increases in the event frequency reflect increases in the number of functional synapses or the presynaptic release probability¹¹.
   1. Fill the patch recording pipette with modified intracellular solution (Table 4) to adjust the reversal potential of the GABA_A receptor-mediated current to -60 mV.
   2. Add 0.5 µM tetrodotoxin to the bath to block spontaneous action potentials.
   3. Hold the voltage at -60 mV to record the mEPSC events for 5 min.
   4. Change the holding potential to 0 mV to record mIPSC events for 5 min. Because M1 neurons show slightly higher reversal potential for AMPA receptor-mediated currents, the mIPSCs of M1 neurons are recorded at +15 mV with 0.1 mM APV.
   5. Wait for a few minutes for the current to stabilize.
   6. Record the mIPSC events for 5 min.
   7. Detect the miniature events using the software, and use events above 10 pA for the analysis. Count the number of mEPSCs or mIPSCs events for 5 min to determine the frequency. Average the amplitudes of the events to obtain the mean amplitude.
   8. Confirm whether bath treatment with 10 µM CNQX or with 10 µM bicuculline methiodide blocks the mEPSCs and mIPSCs events, respectively.
3. Paired-pulse analysis
   NOTE: Presynaptic plasticity can be analyzed using paired-pulse analysis. An increase in the paired-pulse rate suggests a decrease in the presynaptic glutamate or GABA release probability^7,10,11.
   1. To analyze excitatory synapses, add 0.1 mM picrotoxin and record the response at -60 mV. Although we added 4 µM 2-chloroadenosine to the bath, we need to keep in mind that the drug affects the presynaptic release probability¹⁴ .
   2. To analyze inhibitory synapses, add 0.1 mM APV and 4 µM 2-chloroadenosine to the bath and record the response at 0 mV. In M1 neurons, record the response at +15 mV.
   3. Apply paired pulses with an inter-stimulus interval of 100 ms or 200 ms.
   4. Record 50-100 sequential traces at each holding potential and average the values.
   5. Calculate the paired pulse ratio as the ratio of the second peak to the first peak of the postsynaptic current.