J.S. received Ontario Graduate Scholarship. W.I. received a New Investigator Fellowship from Mental Health Research Canada. This work is supported by operating grants to W.I from the Natural Sciences and Engineering Research Council of Canada (06106-2015 RGPIN) and the Canadian Institute for Health Research (PJT 148707).

All animal experiments are approved by the Animal Care Committee of The University of Western Ontario in accordance with the Canadian Council on Animal Care Guidelines (AUP#2014-031).
1. Solutions
<ol>
	<li>Slicing solution 
	<ol>
		<li>Refer to Table 1 for the composition of the slicing solution.</li>
		<li>Prepare a 20x stock solution in advance and store it at 4 &#176;C for up to 1 month.</li>
		<li>For 1x slicing solution, dissolve NaHCO3, glucose, and sucrose in ddH2O, and add the 20x stock. Ensure the osmolarity is between 315-320 mOsm and store the solution for no more than 1 week at 4 &#176;C.</li>
		<li>Fill two beakers with 100 mL of slicing solution and cover them with parafilm. Chill the solution in a freezer until the solution becomes partially frozen (approximately 20 min in -80 &#176;C freezer). Using a gas dispersion tube, bubble both beakers of slicing solution with 95% O2/5% CO2 for 20 min on ice.</li>
	</ol>
	</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td colspan="5">Solution Concentrations (mM)</td>
		</tr>
		<tr>
			<td></td>
			<td>Slicing</td>
			<td>Normal aCSF</td>
			<td>Low Ca2+ aCSF</td>
			<td>Sr+ aCSF</td>
			<td>Pipette/Internal</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>87</td>
			<td>126</td>
			<td>126</td>
			<td>126</td>
			<td>-</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>2.5</td>
			<td>2.5</td>
			<td>2.5</td>
			<td>2.5</td>
			<td>8</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>0.5</td>
			<td>2.5</td>
			<td>0.5</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>SrCl2</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>2.5</td>
			<td>-</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>7</td>
			<td>1.5</td>
			<td>2.5</td>
			<td>1.5</td>
			<td>2</td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>1.25</td>
			<td>1.25</td>
			<td>1.25</td>
			<td>1.25</td>
			<td>-</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>25</td>
			<td>26</td>
			<td>26</td>
			<td>26</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>25</td>
			<td>10</td>
			<td>10</td>
			<td>10</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>75</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>K-gluconate</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>116</td>
		</tr>
		<tr>
			<td>Na-gluconate</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>12</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>10</td>
		</tr>
		<tr>
			<td>K2-EGTA</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>1</td>
		</tr>
		<tr>
			<td>K2ATP</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>4</td>
		</tr>
		<tr>
			<td>Na3GTP</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
			<td>0.3</td>
		</tr>
	</tbody>
</table>
Table 1: The composition of various solutions.
<ol start="2">
	<li>aCSF (for slice recovery and maintenance)
	<ol>
		<li>Refer to Table 1 for the composition of the aCSF.</li>
		<li>Prepare a 20x stock solution in advance and store it at 4 &#176;C for up to 1 month.</li>
		<li>For 1x aCSF, dissolve NaHCO3 and glucose in ddH2O and add the 20x stock. Ensure the osmolarity is between 298-300 mOsm. Use the solution within 1 day.</li>
	</ol>
	</li>
	<li>aCSF (low Ca2+ for recording) 
	<ol>
		<li>Refer to Table 1 for the composition of the low Ca2+ aCSF.</li>
		<li>Prepare a 20x stock solution in advance and store it at 4 &#176;C for up to 1 month.</li>
		<li>For 1x low Ca2+ aCSF, dissolve NaHCO3 and glucose in ddH2O and add 20x (CaCl2 and MgCl2 free) stock, CaCl2 and MgCl2 to specified concentrations. Ensure the osmolarity is between 298-300. Use the solution within 1 day.</li>
	</ol>
	</li>
	<li>aCSF (Sr2+ for recording) 
	<ol>
		<li>Refer to Table 1 for the composition of the Sr2+ aCSF.</li>
		<li>Prepare a 20x stock solution in advance and store it at 4 &#176;C for up to 1 month.</li>
		<li>For 1x Sr2+ aCSF, dissolve NaHCO3 and glucose in ddH2O and add 20x (CaCl2 and MgCl2 free) stock, SrCl2 and MgCl2 to specified concentrations. Ensure the osmolarity is between 298-300. Use the solution within 1 day.</li>
	</ol>
	</li>
	<li>Internal solution
	<ol>
		<li>Refer to Table 1 for the composition of the K-gluconate based internal solution.</li>
		<li>To make 20 mL of internal solution, add 15 mL of molecular biology grade water to a 50 mL tube. Perform the subsequent steps on the ice.</li>
		<li>Prepare the following solutions ahead of time to 1 M stock concentrations in molecular biology grade water. Add (in mL): 2.32 K-gluconate, 0.24 Na-gluconate, 0.20 HEPES, 0.16 KCl, 0.05 K2-EGTA, 0.04 MgCl2 to the 50 mL tube.</li>
		<li>Add 100 &#181;L of 0.3 M Na3GTP.</li>
		<li>Weigh 44.08 mg of K2ATP in a 2 mL microcentrifuge tube and add 1 mL of molecular biology grade water, then add to 50 mL tube.</li>
		<li>Adjust the pH to 7.2-7.4 with 1 M KOH. Ensure the osmolarity is between 283-289 mOsm.</li>
	</ol>
	</li>
</ol>
2. Slice Preparation
<ol>
	<li>Prepare tools
	<ol>
		<li>Add 200 mL of aCSF to the recovery chamber (constructed from a 250 mL beaker with 4 wells and netting) and place the recovery chamber in a water bath (35 &#176;C).</li>
		<li>Cover the chamber with a paraffin film and constantly bubble the aCSF with 95% O2/5% CO2 using a glass dispersion tube for at least 20 min.</li>
		<li>Prepare for the dissection by setting up the tools (scalpel, angled fine scissors, forceps, fine paint brush, plastic spoon).</li>
		<li>Fill a 60 mL syringe with approximately 15 mL of the ice-cold slicing solution from step 1.1.4.</li>
		<li>Prepare the dissection platform by placing a filter paper on the lid of a well plate.</li>
		<li>Prepare the slicing chamber by placing it in the ice tray and filling the tray with ice.</li>
		<li>Set up the vibratome by securing a disposable blade in the blade holder.</li>
		<li>Make a transfer pipette by breaking the tip of a Pasteur pipette and placing a rubber bulb over the broken end.</li>
	</ol>
	</li>
	<li>Dissect mouse brain
	<ol>
		<li>Anesthetize the animal in a chamber saturated with 4% isoflurane until spinal reflexes are absent.</li>
		<li>Decapitate the animal using a guillotine and quickly remove the brain.
		<ol>
			<li>Make a midline incision with a No. 22 scalpel blade from rostral to caudal.</li>
			<li>Laterally peel the scalp on each side of the head.</li>
			<li>Use fine scissors to cut the skull on one side from caudal to rostral (including the side of the frontal bones), using caution not to damage the brain.</li>
			<li>Use forceps to lift the skull piece off the brain and quickly cool the brain with 15 mL of ice-cold slicing solution using the syringe from step 2.1.4.</li>
			<li>Lift the brain out of the skull.</li>
			<li>Place the brain in one of the beakers filled with ice-cold slicing solution (from step 1.1.4) bubbled with 95% O2/5% CO2.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Prepare slices of mouse hypothalamus
	<ol>
		<li>Block the brain for the desired brain area and cut angle (e.g., for coronal hypothalamic slices, trim off the tissue rostral to the optic chiasm and caudal to the pons using a blade and ensure the caudal block has a flat surface perpendicular to the base of the brain).</li>
		<li>Using a cut piece of filter paper, pick up the brain from the anterior side and glue the posterior side to the holding plate using instant glue.</li>
		<li>Quickly place the holding plate into the slicing chamber and fill the chamber with slicing solution from the second beaker in step 1.1.4.</li>
		<li>Secure the slicing chamber and ice tray on the vibratome.</li>
		<li>Define the slicing area (anterior and posterior to the brain) and begin slicing 250 &#181;m thickness coronal slices. Recommended parameters: speed 0.10 mm/s, amplitude 2 mm.</li>
		<li>Trim the slices to the appropriate size for the desired brain area.</li>
		<li>Recover the slices at 35 &#176;C for 30-45 min. Then, remove the recovery chamber from the warming bath and allow the slices to recover at room temperature for an additional 30 min. Keep slices at room temperature for the rest of the day and continue to bubble the bath constantly with 95% O2/5% CO2.</li>
	</ol>
	</li>
</ol>
3. Whole-cell Patch ClampRecording
<ol>
	<li>Pull the patch pipettes
	<ol>
		<li>Using the suggested parameters for the whole-cell recording from the pipette puller&#8217;s manual, pull patch pipettes from thick walled glass to a pipette resistance of 3-5 M&#8486;.</li>
		<li>Using a microsyringe (commercial or homemade), fill a pipette tip with filtered internal solution. To make a microsyringe, burn the tip of a 1 mL syringe and allow the tip to fall creating a long fine tip.</li>
	</ol>
	</li>
	<li>Obtain the whole-cell configuration
	<ol>
		<li>Place the recording pipette just above the slice and offset pipette current in the voltage clamp mode. Apply slight positive pressure to the pipette and lock the stopcock.</li>
		<li>Select a healthy cell with an intact membrane and approach the cell with the pipette. The positive pressure should cause a slight disturbance in the tissue (i.e. a slow wave in the tissue when entering).</li>
		<li>Slowly continue to bring the pipette closer to the cell using a diagonal motion until the pipette forms a small dimple on the cell surface.</li>
		<li>Release the positive pressure lock. The cell will begin to form a seal and the resistance will increase above 1 G&#8486;. In voltage clamp, hold the cell at -68 mV.</li>
		<li>Slightly pull away from the cell diagonally to remove excess pressure from the cell.</li>
		<li>Compensate for the fast and slow pipette capacitance.</li>
		<li>Apply a brief suction through the tube connected to the pipette holder to break through the cell and obtain whole-cell configuration.</li>
		<li>Switch to Cell mode on the membrane test window in an electrophysiology Data acquisition and analysis software (e.g., Clampex).</li>
		<li>Before each voltage clamp recording, perform a membrane test using the same software and record the relevant parameters in a lab book (membrane resistance, access resistance, and capacitance).</li>
		<li>Maintain the temperature of the recording bath at 27&#8211;30 &#176;C and the flow rate at 1.5&#8211;2.0 mL/min for subsequent experiments.</li>
	</ol>
	</li>
</ol>
4. Multiplicity Experiments
<ol>
	<li>Experiment 1: estimating multiplicity using 4-AP
	<ol>
		<li>In voltage clamp, hold the cell at -68 mV. Using the same software, record the sEPSCs while perfusing the bath with low Ca2+ aCSF. Record for at least 5 min after the start of whole-cell configuration to ensure a stable baseline recording as the synaptic activity may be high shortly after the breakthrough of the membrane.</li>
		<li>Using a micropipette, add 4-AP to the aCSF and bath apply 30 &#181;M 4-AP. Record sEPSCs for at least 10 min to obtain the full drug effect.</li>
		<li>Add 0.5 &#181;M TTX and 10 &#181;M Cd2+ to the aCSF with 4-AP and record the mEPSCs for at least 10 min.</li>
		<li>For offline analysis, use the last 1 min of baseline immediately before the application of 4-AP (in low Ca2+ aCSF), the 10th min of 4-AP application and the 10th min of TTX application.</li>
	</ol>
	</li>
	<li>Experiment 2: desynchronize vesicle release using Sr2+
	<ol>
		<li>While perfusing the bath with normal Ca2+ aCSF (the same as the bath aCSF) record sEPSCs for at least 5 min.</li>
		<li>Switch from the normal Ca2+ aCSF and begin perfusing Sr2+ aCSF (from step 1.4) and record sEPSCs.</li>
		<li>For offline analysis, to determine whether the large amplitude sEPSCs are due to the synchronous release of vesicles, compare the last 1 min of baseline (in normal aCSF) to the 10th minute of Sr2+ aCSF application.</li>
	</ol>
	</li>
	<li>Experiment 3: test for multivesicular release using &#947;-DGG
	<ol>
		<li>In low Ca2+ aCSF record sEPSCs for at least 5 min.</li>
		<li>Add 30 &#181;M 4-AP to the aCSF through the perfusion system. Record sEPSCs for at least 10 min.</li>
		<li>Add 200 &#181;M &#947;-DGG to the aCSF with 4-AP and record the sEPSCs for at least 10 min.</li>
		<li>As a control experiment in a separate cell, perform steps 1-3 but apply a low concentration (125 nM) of DNQX instead of &#947;-DGG.</li>
		<li>For offline analysis, analyze the last min of each drug application.</li>
	</ol>
	</li>
	<li>Experiment 4: Stimulate afferent inputs to increase action potential firing. 
	<ol>
		<li>Record sEPSCs in normal Ca2+ aCSF.</li>
		<li>Stimulate the afferents using a monopolar glass electrode filled with aCSF at a rate of 20 Hz for 2 s and repeat 10 times with an inter-burst interval of 20 sec.</li>
		<li>For analysis, use the 5,000 ms before the first stimulus as the baseline and compare to the 10-300 ms after the final stimulus and then take the average amplitude and frequency change over 10 trials.</li>
	</ol>
	</li>
</ol>
5. Analysis
<ol>
	<li>Analyze sEPSCs and mEPSCs using a program that detects and analyzes synaptic currents (e.g., Mini Analysis software).
	<ol>
		<li>Using this software, use the suggested detection parameters for detecting AMPA Receptor EPSCs (or GABA Receptor EPSCs if recording inhibitory currents).</li>
		<li>Use the Nonstop Analysis function to detect EPSCs in the recording.</li>
		<li>Manually scan each recording to ensure the program is accurately detecting each event (e.g., ensure events are not being missed or counted twice).</li>
		<li>Export the event data by copying it to the clipboard and paste it into a data management software (e.g., Excel)</li>
		<li>Calculate the average frequency and/or amplitude for each drug treatment and perform the relevant statistical analyses.</li>
	</ol>
	</li>
</ol>

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L., Chen, Q., Feng, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+Brain+Slice+Methods+for+Adult+and+Aging+Animals:+Application+of+Targeted+Patch+Clamp+Analysis+and+Optogenetics.">Acute Brain Slice Methods for Adult and Aging Animals: Application of Targeted Patch Clamp Analysis and Optogenetics.</a> Methods in molecular biology (Clifton, N.J.). , 1183-242 (2014).</li><li>Richerson, G. B., Messer, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+composition+of+experimental+solutions+on+neuronal+survival+during+rat+brain+slicing.">Effect of composition of experimental solutions on neuronal survival during rat brain slicing.</a> Experimental neurology. 131 (1), 133-143 (1995).</li><li>Tanaka, Y., Tanaka, Y., Furuta, T., Yanagawa, Y., Kaneko, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+cutting+solutions+on+the+viability+of+GABAergic+interneurons+in+cerebral+cortical+slices+of+adult+mice.">The effects of cutting solutions on the viability of GABAergic interneurons in cerebral cortical slices of adult mice.</a> Journal of neuroscience methods. 171 (1), 118-125 (2008).</li><li>Ye, J. H., Zhang, J., Xiao, C., Kong, J. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patch-clamp+studies+in+the+CNS+illustrate+a+simple+new+method+for+obtaining+viable+neurons+in+rat+brain+slices:+glycerol+replacement+of+NaCl+protects+CNS+neurons.">Patch-clamp studies in the CNS illustrate a simple new method for obtaining viable neurons in rat brain slices: glycerol replacement of NaCl protects CNS neurons.</a> Journal of neuroscience methods. 158 (2), 251-259 (2006).</li><li>Gunn, B. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dysfunctional+astrocytic+and+synaptic+regulation+of+hypothalamic+glutamatergic+transmission+in+a+mouse+model+of+early-life+adversity:+relevance+to+neurosteroids+and+programming+of+the+stress+response.">Dysfunctional astrocytic and synaptic regulation of hypothalamic glutamatergic transmission in a mouse model of early-life adversity: relevance to neurosteroids and programming of the stress response.</a> Journal of Neuroscience. 33 (50), 19534-19554 (2013).</li><li>Su, H., Alroy, G., Kirson, E. D., Yaari, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+calcium+modulates+persistent+sodium+current-dependent+burst-firing+in+hippocampal+pyramidal+neurons.">Extracellular calcium modulates persistent sodium current-dependent burst-firing in hippocampal pyramidal neurons.</a> The Journal of neuroscience the official journal of the Society for Neuroscience. 21 (12), 4173-4182 (2001).</li><li>Frankenhaeuser, B., Hodgkin, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+action+of+calcium+on+the+electrical+properties+of+squid+axons.">The action of calcium on the electrical properties of squid axons.</a> The Journal of physiology. 137 (2), 218-244 (1957).</li><li>Xiong, G., Metheny, H., Johnson, B. N., Cohen, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Comparison+of+Different+Slicing+Planes+in+Preservation+of+Major+Hippocampal+Pathway+Fibers+in+the+Mouse.">A Comparison of Different Slicing Planes in Preservation of Major Hippocampal Pathway Fibers in the Mouse.</a> Frontiers in neuroanatomy. 11, 107 (2017).</li></ol>

The authors have nothing to disclose

One important requirement for a successful patch clamp electrophysiology experiment is obtaining healthy slices/cells. Our described protocol is optimized for hypothalamic slices that contain PVN neurons. Other brain areas may require modified solutions and slicing methods21,22,23,24. For the recording, it is critical to only accept stable recordings by constantly monitoring cell properties suc...

Synaptic transmission is a fundamental mechanism for communication between neurons, and hence, brain function. Synaptic transmission is also labile and can change its efficacy in an activity-dependent manner as well as in response to modulatory signals1. Thus, examining&#160;synaptic function has been a key focus of neuroscience research. Whole-cell patch clamp electrophysiology is a versatile technique that enables us to understand, by devising experimental designs and data analyses, in-depth biophysical and molecular mechanisms of synaptic transmission. A commonly used approach, perhaps owing to the simplicity of the technique and concept, is the measurement of miniature excitatory/inhibitory postsynaptic currents (mE/IPSCs) under the voltage clamp configuration2,3,4,5,6. Individual mPSCs represent the flow of ions through postsynaptic ionotropic receptors (e.g. AMPA and GABAA receptors) in response to the binding of their respective neurotransmitters released from the presynaptic terminal 7. Because the recording is obtained in the presence of the voltage-gated Na+ channel blocker tetrodotoxin (TTX), the release is action potential-independent and normally involves a single synaptic vesicle that contains neurotransmitter. Based on this assumption, the average amplitude of mPSCs is widely used as a crude estimate for the quantal size, which represents the number and functionality of postsynaptic receptors opposing a single release site. On the other hand, the frequency of mPSCs is considered to represent a combination of the total number of synapses terminating onto the postsynaptic cell and their average release probability. However, these parameters do not measure another variable-multiplicativity of synapses, or synaptic multiplicity &#8212; which is important for the efficacy of synaptic transmission.
Based on the quantal theory of synaptic transmission7,8,9, the strength of a given connection between a pair of neurons is dependent on three factors: the number of functional synapses (N), the postsynaptic response to the release of a single synaptic vesicle (quantal size; Q) and the probability of neurotransmitter release (Pr). Synaptic multiplicity is equivalent to N. The development of synaptic multiplicity or the pruning of multiplicative synapses is plastic throughout development and in different disease states3,4,6,10. For this reason, characterizing synaptic multiplicity has important implications for understanding the efficacy of synaptic transmission in health and disease. Techniques, such as electron microscopy can identify structural evidence of synaptic multiplicity by detecting multiple synaptic contacts originating from the same axon onto the same postsynaptic neuron11,12,13,14. However, these structurally identified multisynapses can be functionally silent15,16. Precise functional examination of N requires technically challenging electrophysiological approaches, such as paired whole-cell recordings that can identify whether a given connection has multiple functional release sites and minimal stimulation approaches that aim to recruit a single putative axon.
In this protocol, we describe a simple method for estimating synaptic multiplicity by adopting a method originally developed by Hsia et al2. This technique involves the measurement of spontaneous PSCs (sPSCs) and mPSCs using whole-cell patch clamp electrophysiology, which allows us to estimate the degree of synaptic multiplicity across all inputs to a given neuron.&#160; As previously defined, synaptic multiplicity reflects the number of synapses between a given pre- and postsynaptic neuron. If multiple synapses are recruited in synchrony by an action potential, there will be a high probability of temporal summation of individual (i.e. quantal) PSCs, generating a greater amplitude PSC.&#160; In mPSC recordings (in which action potentials are blocked by TTX), the probability of temporal summation of individual (non-synchronous) mPSCs is low. Using this rationale, synaptic multiplicity can be estimated by comparing the sPSC amplitude (with action potential-dependent release) to the mPSC amplitude.
To examine the existence of multiplicity we describe four experiments and their analyses using glutamatergic EPSCs as an example. However, the same approach can be used for the fast GABAergic/glycinergic transmission (IPSCs). A brief rationale for each experiment is described below. First, as explained above, synaptic multiplicity can be estimated by comparing the amplitude of sEPSCs to mEPSCs. There are two requirements for this approach; 1) presynaptic axons must fire a sufficient number of action potentials during recording, and 2) Pr must be high so that multiple synapses release neurotransmitter upon the arrival of an action potential. In order to meet these requirements, sEPSCs are first recorded in low Ca2+ artificial cerebrospinal fluid (aCSF), and then recorded in the presence of a low concentration of the K+ channel antagonist, 4-Aminopyridine (4-AP) to increase action potential firing and Pr. Then action potential firing is blocked by TTX and Pr decreased by a voltage-gated Ca2+ channel blocker Cd2+. The amplitude of sEPSCs (with 4AP) is compared to that of mEPSC (with 4AP, TTX, and Cd2+). In the second experiment, Ca2+ is replaced by equimolar Sr2+ in the aCSF to desynchronize vesicle release. As Ca2+ is required for the synchronous release of vesicles, replacement with Sr2+ should eliminate the large amplitude sEPSCs that are indicative of multiplicity. Third, mechanistically, multiplicity can result from either multiple synaptic contacts to the same postsynaptic neuron or multivesicular release (i.e. multiple vesicles released within a single synaptic contact)17,18. To differentiate between the two types of multiplicity, the third experiment uses a low affinity, fast dissociating competitive antagonist of AMPA receptors, &#947;-D-glutamylglycine (&#947;-DGG)17,18 to determine whether large sEPSC are the result of the temporal summation of independent synapses or multivesicular release acting on an overlapping population of postsynaptic receptors. If the large amplitude events arise from multivesicular release, &#947;-DGG will be less effective at inhibiting larger compared to smaller sEPSCs, whereas large sEPSCs that arise from the temporal summation of multiple synaptic contacts will be similarly affected by &#947;-DGG. In the fourth experiment, a more physiological method is used to enhance action potential firing, namely afferent synaptic stimulation. Bursts of synaptic activity can transiently increase/facilitate the spontaneous action potential firing and release probability of the stimulated afferents. Therefore, this approach allows multiplicity to manifest in a more physiological manner.
The following protocol describes the method for conducting these experiments in mouse hypothalamic tissue. Specifically, corticotropin releasing hormone (CRH) neurons of the paraventricular nucleus of the hypothalamus (PVN) are used. We describe the procedures for conducting whole-cell patch clamp electrophysiology and explain the specific experiments to test for synaptic multiplicity.

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			<td>1 ml syringe</td>
			<td>BD</td>
			<td>309659</td>
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			<td>10 blade</td>
			<td>Fisher Scientific/others</td>
			<td>35698</td>
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			<td>22 blade</td>
			<td>VWR/others</td>
			<td>21909-626</td>
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			<td>22 uM syringe filters</td>
			<td>Milipore</td>
			<td>09-719-000</td>
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			<td>Adson foreceps</td>
			<td>Harvard Instruments</td>
			<td>72-8547</td>
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			<td>Angled sharp scissors</td>
			<td>Harvard Instruments</td>
			<td>72-8437</td>
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			<td>Clampex</td>
			<td>Molecular Devices</td>
			<td>pClamp 10</td>
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			<td>Double edge blade</td>
			<td>VWR</td>
			<td>74-0002</td>
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			<td>Filter paper</td>
			<td>Sigma/others</td>
			<td>1001090</td>
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			<td>15-183-35/various</td>
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			<td>LG-8680-120</td>
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			<td>M60303</td>
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			<td>Krazy glue</td>
			<td>various</td>
			<td>various</td>
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			<td>Mini analysis</td>
			<td>Synaptosoft</td>
			<td>MiniAnalysis 6</td>
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			<td>Model 5600</td>
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			<td>Pasteur pipette</td>
			<td>VWR</td>
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			<td>ph meter</td>
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			<td>82024-550</td>
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			<td>Scalpel handle No. 3</td>
			<td>Harvard Instruments</td>
			<td>72-8350</td>
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			<td>Scalpel handle No. 4</td>
			<td>Harvard Instruments</td>
			<td>72-8356</td>
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			<td>Single edge blade</td>
			<td>VWR</td>
			<td>55411-050</td>
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			<td>Vibratome slicer</td>
			<td>Leica</td>
			<td>VT1200S</td>
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			<td>Millipore</td>
			<td>Milli-Q Academic A10</td>
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			<td>4-AP</td>
			<td>Sigma</td>
			<td>275875</td>
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			<td>BAPTA</td>
			<td>molecular probes</td>
			<td>B1204</td>
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			<td>CaCl2*2H2O</td>
			<td>Sigma</td>
			<td>C7902</td>
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			<td>CdCl2</td>
			<td>sigma</td>
			<td>202908</td>
			<td></td>
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			<td>DNQX</td>
			<td>Tocris</td>
			<td>189</td>
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			<td>EGTA</td>
			<td>Sigma</td>
			<td>E3889</td>
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			<td>glucose</td>
			<td>Sigma</td>
			<td>G5767</td>
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			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375</td>
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			<td>K2-ATP</td>
			<td>Sigma</td>
			<td>A8937</td>
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			<td>KCl</td>
			<td>Sigma</td>
			<td>P9333</td>
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			<td>K-gluconate</td>
			<td>Sigma</td>
			<td>G4500</td>
			<td></td>
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			<td>MgCl2*6H2O</td>
			<td>Sigma</td>
			<td>M2670</td>
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			<td>Molecular biology grade water</td>
			<td>Sigma</td>
			<td>W4502-1L</td>
			<td></td>
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			<td>Na3GTP</td>
			<td>Sigma</td>
			<td>G8877</td>
			<td></td>
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			<td>NaCl</td>
			<td>Bioshop</td>
			<td>SOD001.1</td>
			<td></td>
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			<td>Na-gluconate</td>
			<td>Sigma</td>
			<td>S2054</td>
			<td></td>
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		<tr>
			<td>NaH2PO4</td>
			<td>Sigma</td>
			<td>71504</td>
			<td></td>
		</tr>
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			<td>NaHCO3</td>
			<td>Sigma</td>
			<td>S6014</td>
			<td></td>
		</tr>
		<tr>
			<td>Picrotoxin</td>
			<td>sigma</td>
			<td>P1675</td>
			<td></td>
		</tr>
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			<td>SrCl</td>
			<td>Sigma</td>
			<td>255521</td>
			<td></td>
		</tr>
		<tr>
			<td>sucrose</td>
			<td>Bioshop</td>
			<td>SUC507.1</td>
			<td></td>
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			<td>TTX</td>
			<td>Alamone Labs</td>
			<td>T-550</td>
			<td></td>
		</tr>
		<tr>
			<td>yDGG</td>
			<td>Tocris</td>
			<td>6729-55-1</td>
			<td></td>
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	</tbody>
</table>

evaluation of synaptic multiplicity using whole-cell patch-clamp electrophysiology

In the central nervous system, a pair of neurons often form&#160;multiple synaptic contacts and/or functional neurotransmitter release sites (synaptic multiplicity). Synaptic multiplicity is plastic and changes throughout development and in different physiological conditions, being an important determinant for the efficacy of synaptic transmission. Here, we outline experiments for estimating the degree of multiplicity of synapses terminating onto a given postsynaptic neuron using whole-cell patch clamp electrophysiology in acute brain slices. Specifically, voltage-clamp recording is used to compare the difference between the amplitude of spontaneous excitatory postsynaptic currents (sEPSCs) and miniature excitatory postsynaptic currents (mEPSCs). The theory behind this method is that afferent inputs that exhibit multiplicity will show large, action potential-dependent sEPSCs due to the synchronous release that occurs at each synaptic contact. In contrast, action potential-independent release (which is asynchronous) will generate smaller amplitude mEPSCs. This article outlines a set of experiments and analyses to characterize the existence of synaptic multiplicity and discusses the requirements and limitations of the technique. This technique can be applied to investigate how different behavioral, pharmacological or environmental interventions in vivo affect the organization of synaptic contacts in different brain areas.

The above protocol describes a method for using whole-cell patch clamp electrophysiology to examine the degree of synaptic multiplicity, using mouse hypothalamic neurons as an example. This slice preparation technique should yield healthy viable cells that do not have a swollen membrane or nucleus (Figure 1). Each step in the protocol is important for the health of the tissue and quality of the recordings.
<img ...

Watch this Scientific Journal Video about Evaluation of Synaptic Multiplicity Using Whole-cell Patch-clamp Electrophysiology at JoVE.com

Evaluation of Synaptic Multiplicity Using Whole-cell Patch-clamp Electrophysiology

Here, we present a protocol for evaluating the functional synaptic multiplicity using whole-cell patch clamp electrophysiology in acute brain slices.

In the central nervous system, a pair of neurons often form&#160;multiple synaptic contacts and/or functional neurotransmitter release sites (synaptic ...

evaluation-synaptic-multiplicity-using-whole-cell-patch-clamp

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Neuroscience

12.8K Views.  University of Western Ontario. Here, we present a protocol for evaluating the functional synaptic multiplicity using whole-cell patch clamp electrophysiology in acute brain slices.

Video: Evaluation of Synaptic Multiplicity Using Whole-cell Patch-clamp Electrophysiology

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Culturing and Electrophysiology of Cells on NRCC Patch-clamp Chips

Due to its exquisite sensitivity and the ability to monitor and control individual cells at the level of ion channels, patch-clamping is the gold standard of electrophysiology applied to disease models and pharmaceutical screens alike 1. The method traditionally involves gently contacting a cell with a glass pipette filled by a physiological solution in order to isolate a patch of the membrane under its apex 2. An electrode inserted in the pipette captures ion-channel activity within the membrane patch or, when ruptured, for the whole cell. In the last decade, patch-clamp chips have been proposed as an alternative 3, 4: a suspended film separates the physiological medium from the culture medium, and an aperture microfabricated in the film replaces the apex of the pipette. Patch-clamp chips have been integrated in automated systems and commercialized for high-throughput screening 5. To increase throughput, they include the fluidic delivery of cells from suspension, their positioning on the aperture by suction, and automated routines to detect cell-to-probe seals and enter into whole cell mode. We have reported on the fabrication of a silicon patch-clamp chip with optimized impedance and orifice shape that permits the high-quality recording of action potentials in cultured snail neurons 6; recently, we have also reported progress towards interrogating mammalian neurons 7. Our patch-clamp chips are fabricated at the Canadian Photonics Fabrication Centre 8, a commercial foundry, and are available in large series. We are eager to engage in collaborations with electrophysiologists to validate the use of the NRCC technology in different models. The chips are used according to the general scheme represented in Figure 1: the silicon chip is at the bottom of a Plexiglas culture vial and the back of the aperture is connected to a subterranean channel fitted with tubes at either end of the package. Cells are cultured in the vial and the cell on top of the probe is monitored by a measuring electrode inserted in the channel.The two outside fluidic ports facilitate solution exchange with minimal disturbance to the cell; this is an advantage compared to glass pipettes for intracellular perfusion.
<img alt="Figure 1" src="/files/ftp_upload/3288/3288fig1.jpg" />
Figure 1. Principle of measurement using the NRCC patch-clamp chip
We detail here the protocols to sterilize and prime the chips, load them with medium, plate them with cells, and finally use them for electrophysiological recordings.

We show how planar patch-clamp chips fabricated at the National Research Council of Canada are sterilized, primed, loaded with medium, plated with cells, and used for electrophysiological recordings.

Due to its exquisite sensitivity and the ability to monitor and control individual cells at the level of ion channels, patch-clamping is the gold standard ...

Whole Cell Patch Clamp for Investigating the Mechanisms of Infrared Neural Stimulation

It has been demonstrated in recent years that pulsed, infrared laser light can be used to elicit electrical responses in neural tissue, independent of any further modification of the target tissue. Infrared neural stimulation has been reported in a variety of peripheral and sensory neural tissue in vivo, with particular interest shown in stimulation of neurons in the auditory nerve. However, while INS has been shown to work in these settings, the mechanism (or mechanisms) by which infrared light causes neural excitation is currently not well understood. The protocol presented here describes a whole cell patch clamp method designed to facilitate the investigation of infrared neural stimulation in cultured primary auditory neurons. By thoroughly characterizing the response of these cells to infrared laser illumination in vitro under controlled conditions, it may be possible to gain an improved understanding of the fundamental physical and biochemical processes underlying infrared neural stimulation.

Infrared nerve stimulation has been proposed as an alternative to electrical stimulation in a range of nerve types, including those associated with the auditory system. This protocol describes a patch clamp method for studying the mechanism of infrared nerve stimulation in a culture of primary auditory neurons.

It has been  demonstrated in recent years that pulsed, infrared laser light can be used to  elicit electrical responses in neural tissue, independent of ...

Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells

Mauthner cells (M-cells) are large reticulospinal neurons located in the hindbrain of teleost fish. They are key neurons involved in a characteristic behavior known as the C-start or escape response that occurs when the organism perceives a threat. The M-cell has been extensively studied in adult goldfish where it has been shown to receive a wide range of excitatory, inhibitory and neuromodulatory signals1. We have been examining M-cell activity in embryonic zebrafish in order to study aspects of synaptic development in a vertebrate preparation. In the late 1990s Ali and colleagues developed a preparation for patch clamp recording from M-cells in zebrafish embryos, in which the CNS was largely intact2,3,4. The objective at that time was to record synaptic activity from hindbrain neurons, spinal cord neurons and trunk skeletal muscle while maintaining functional synaptic connections within an intact brain-spinal cord preparation. This preparation is still used in our laboratory today. To examine the mechanisms underlying developmental synaptic plasticity, we record excitatory (AMPA and NMDA-mediated)5,6 and inhibitory (GABA and glycine) synaptic currents from developing M-cells. Importantly, this unique preparation allows us to return to the same cell (M-cell) from preparation to preparation to carefully examine synaptic plasticity and neuro-development in an embryonic organism. The benefits provided by this preparation include 1) intact, functional synaptic connections onto the M-cell, 2) relatively inexpensive preparations, 3) a large supply of readily available embryos 4) the ability to return to the same cell type (i.e. M-cell) in every preparation, so that synaptic development at the level of an individual cell can be examined from fish to fish, and 5) imaging of whole preparations due to the transparent nature of the embryos.

We have developed an intact brain-spinal cord preparation to record and monitor electrical activity via patch clamp recording from the Mauthner neurons and other reticulospinal cells in zebrafish embryos. Thus, we are able to record excitatory and inhibitory synaptic currents, voltage-gated channel activity and action potentials from key neurons in a developing embryo.

Mauthner cells (M-cells) are large  reticulospinal neurons located in the hindbrain of teleost fish. They are key  neurons involved in a characteristic ...

A Computer-assisted Multi-electrode Patch-clamp System

The patch-clamp technique is today the most well-established method for recording electrical activity from individual neurons or their subcellular compartments. Nevertheless, achieving stable recordings, even from individual cells, remains a time-consuming procedure of considerable complexity. Automation of many steps in conjunction with efficient information display can greatly assist experimentalists in performing a larger number of recordings with greater reliability and in less time. In order to achieve large-scale recordings we concluded the most efficient approach is not to fully automatize the process but to simplify the experimental steps and reduce the chances of human error while efficiently incorporating the experimenter's experience and visual feedback. With these goals in mind we developed a computer-assisted system which centralizes all the controls necessary for a multi-electrode patch-clamp experiment in a single interface, a commercially available wireless gamepad, while displaying experiment related information and guidance cues on the computer screen. Here we describe the different components of the system which allowed us to reduce the time required for achieving the recording configuration and substantially increase the chances of successfully recording large numbers of neurons simultaneously.

Multi-electrode patch-clamp recordings constitute a complex task. Here we show how, by automating of many of the experimental steps, it is possible to accelerate the process leading to qualitative improvement in performance and number of recordings.

The patch-clamp technique is today the most  well-established method for recording electrical activity from individual  neurons or their subcellular ...

Whole-cell Patch-clamp Recordings from Morphologically- and Neurochemically-identified Hippocampal Interneurons

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal population (10-20%), but show a high level of physiological, morphological, and neurochemical heterogeneity, reflecting their diverse functions. Therefore, investigation of interneurons provides important insights into the organization principles and function of neuronal circuits. This, however, requires an integrated physiological and neuroanatomical approach for the selection and identification of individual interneuron types. Whole-cell patch-clamp recording from acute brain slices of transgenic animals, expressing fluorescent proteins under the promoters of interneuron-specific markers, provides an efficient method to target and electrophysiologically characterize intrinsic and synaptic properties of specific interneuron types. Combined with intracellular dye labeling, this approach can be extended with post-hoc morphological and immunocytochemical analysis, enabling systematic identification of recorded neurons. These methods can be tailored to suit a broad range of scientific questions regarding functional properties of diverse types of cortical neurons.

Cortical networks are controlled by a small, but diverse set of inhibitory interneurons. Functional investigation of interneurons therefore requires targeted recording and rigorous identification. Described here is a combined approach involving whole-cell recordings from single or synaptically-coupled pairs of neurons with intracellular labeling, post-hoc morphological and immunocytochemical analysis.

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Subcellular Patch-clamp Recordings from the Somatodendritic Domain of Nigral Dopamine Neurons

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding these fundamental functions will shed light on the information transfer in these neurons. Dendritic patch-clamp recordings provide the possibility to directly examine the electrical properties of dendrites and underlying voltage-gated ion channels. However, these fine structures are not easily accessible to patch pipettes because of their small diameter. This report describes a step-by-step procedure to collect stable and reliable recordings from the dendrites of dopaminergic neurons in acute slices. Electrophysiological measurements are combined with post hoc recovery of cell morphology. Successful experiments rely on improved preparation of slices, solutions and pipettes, adequate adjustment of the optics and stability of the pipette in contact with the recorded structure. Standard principles of somatic patch-clamp recording are applied to dendrites but with a gentler approach of the pipette. These versatile techniques can be implemented to address various questions concerning the excitable properties of dendrites.

Subcellular patch-clamp recordings offer the possibility to investigate the functional properties of dendrites. However these fine structures are not easily accessible due to their small diameter. The protocol described here aims to facilitate the collection of stable and reliable recordings from the dendrites of dopamine neurons in slices in vitro.

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding ...

Whole-cell Patch-clamp Recordings for Electrophysiological Determination of Ion Selectivity in Channelrhodopsins

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate electrical processes in target cells. In this context, ion selectivity of a channelrhodopsin is of particular importance. This article describes the investigation of chloride selectivity for a recently identified anion-conducting channelrhodopsin of Proteomonas sulcata via electrophysiological patch-clamp recordings on HEK293 cells. The experimental procedure for measuring light-gated photocurrents demands a fast switchable &#8211; ideally monochromatic &#8211; light source coupled into the microscope of an otherwise conventional patch-clamp setup. Preparative procedures prior to the experiment are outlined involving preparation of buffered solutions, considerations on liquid junction potentials, seeding and transfection of cells, and pulling of patch pipettes. The actual recording of current-voltage relations to determine the reversal potentials for different chloride concentrations takes place 24 h to 48 h after transfection. Finally, electrophysiological data are analyzed with respect to theoretical considerations of chloride conduction.

This article describes how the ion selectivity of channelrhodopsin is determined with electrophysiological whole-cell patch-clamp recordings using HEK293 cells. Here, the experimental procedure for investigating chloride selectivity of an anion-selective channelrhodopsin is demonstrated. However, the procedure is transferable to other channelrhodopsins of distinct selectivity.

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate ...

Slice Patch Clamp Technique for Analyzing Learning-Induced Plasticity

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze motor-learning induced plasticity, we trained rats using an accelerated rotor rod task. Rats performed the task 10 times at 30-s intervals for 1 or 2 days. Performance was significantly improved on the training days compared to the first trial. We then prepared acute brain slices of the primary motor cortex (M1) in untrained and trained rats. Current-clamp analysis showed dynamic changes in resting membrane potential, spike threshold, afterhyperpolarization, and membrane resistance in layer II/III pyramidal neurons. Current injection induced many more spikes in 2-day trained rats than in untrained controls.
To analyze contextual-learning induced plasticity, we trained rats using an inhibitory avoidance (IA) task. After experiencing foot-shock in the dark side of a box, the rats learned to avoid it, staying in the lighted side. We prepared acute hippocampal slices from untrained, IA-trained, unpaired, and walk-through rats. Voltage-clamp analysis was used to sequentially record miniature excitatory and inhibitory postsynaptic currents (mEPSCs and mIPSCs) from the same CA1 neuron. We found different mean mEPSC and mIPSC amplitudes in each CA1 neuron, suggesting that each neuron had different postsynaptic strengths at its excitatory and inhibitory synapses. Moreover, compared with untrained controls, IA-trained rats had higher mEPSC and mIPSC amplitudes, with broad diversity. These results suggested that contextual learning creates postsynaptic diversity in both excitatory and inhibitory synapses at each CA1 neuron.
AMPA or GABAA receptors seemed to mediate the postsynaptic currents, since bath treatment with CNQX or bicuculline blocked the mEPSC or mIPSC events, respectively. This technique can be used to study different types of learning in other regions, such as the sensory cortex and amygdala.

The slice patch clamp technique is an effective method for analyzing learning-induced changes in the intrinsic properties and plasticity of excitatory or inhibitory synapses.