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.

<table><tbody><tr><td>1 mL syringe</td><td>BD</td><td>309659</td><td></td></tr><tr><td>10 blade</td><td>Fisher Scientific/others</td><td>35698</td><td></td></tr><tr><td>22 blade</td><td>VWR/others</td><td>21909-626</td><td></td></tr><tr><td>22 &amp;mu;M syringe filters</td><td>Milipore</td><td>09-719-000</td><td></td></tr><tr><td>Adson foreceps</td><td>Harvard Instruments</td><td>72-8547</td><td></td></tr><tr><td>Angled sharp scissors</td><td>Harvard Instruments</td><td>72-8437</td><td></td></tr><tr><td>Clampex</td><td>Molecular Devices</td><td>pClamp 10</td><td></td></tr><tr><td>Double edge blade</td><td>VWR</td><td>74-0002</td><td></td></tr><tr><td>Filter paper</td><td>Sigma/others</td><td>1001090</td><td></td></tr><tr><td>Fine paintbrush</td><td>Fisher/various</td><td>15-183-35/various</td><td></td></tr><tr><td>Gas Dispersion Tube</td><td>VWR</td><td>LG-8680-120</td><td></td></tr><tr><td>Isoflurane</td><td>Fresenius Kabi/others</td><td>M60303</td><td></td></tr><tr><td>Krazy glue</td><td>various</td><td>various</td><td></td></tr><tr><td>Mini analysis</td><td>Synaptosoft</td><td>MiniAnalysis 6</td><td></td></tr><tr><td>Osmomoter</td><td>Wescor Inc</td><td>Model 5600</td><td></td></tr><tr><td>Parafilm</td><td>Sigma</td><td>PM-996</td><td></td></tr><tr><td>Pasteur pipette</td><td>VWR</td><td>14672-200</td><td></td></tr><tr><td>ph meter</td><td>Mettler Toledo</td><td>FE20-ATC</td><td></td></tr><tr><td>Rubber bulb</td><td>VWR</td><td>82024-550</td><td></td></tr><tr><td>Scalpel handle No. 3</td><td>Harvard Instruments</td><td>72-8350</td><td></td></tr><tr><td>Scalpel handle No. 4</td><td>Harvard Instruments</td><td>72-8356</td><td></td></tr><tr><td>Single edge blade</td><td>VWR</td><td>55411-050</td><td></td></tr><tr><td>Vibratome slicer</td><td>Leica</td><td>VT1200S</td><td></td></tr><tr><td>Water Purification System</td><td>Millipore</td><td>Milli-Q Academic A10</td><td></td></tr><tr><td>Well plate lid</td><td>Fisher/various</td><td>07-201-590/various</td><td></td></tr><tr><td>&lt;strong&gt;Chemicals/reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>4-AP</td><td>Sigma</td><td>275875</td><td></td></tr><tr><td>BAPTA</td><td>molecular probes</td><td>B1204</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;*2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma</td><td>C7902</td><td></td></tr><tr><td>CdCl&lt;sub&gt;2&lt;/sub&gt;</td><td>sigma</td><td>202908</td><td></td></tr><tr><td>DNQX</td><td>Tocris</td><td>189</td><td></td></tr><tr><td>EGTA</td><td>Sigma</td><td>E3889</td><td></td></tr><tr><td>glucose</td><td>Sigma</td><td>G5767</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>H3375</td><td></td></tr><tr><td>K2-ATP</td><td>Sigma</td><td>A8937</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P9333</td><td></td></tr><tr><td>K-gluconate</td><td>Sigma</td><td>G4500</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;*6H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma</td><td>M2670</td><td></td></tr><tr><td>Molecular biology grade water</td><td>Sigma</td><td>W4502-1L</td><td></td></tr><tr><td>Na&lt;sub&gt;3&lt;/sub&gt;GTP</td><td>Sigma</td><td>G8877</td><td></td></tr><tr><td>NaCl</td><td>Bioshop</td><td>SOD001.1</td><td></td></tr><tr><td>Na-gluconate</td><td>Sigma</td><td>S2054</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>71504</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma</td><td>S6014</td><td></td></tr><tr><td>Picrotoxin</td><td>sigma</td><td>P1675</td><td></td></tr><tr><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></tr><tr><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></tr></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.
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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. In de hersenen vormen een paar neuronen vaak meerdere synaptische contacten, die synaptische multipliciteit worden genoemd. Echter, nauwkeurig onderzoek van synaptische multipliciteit vereist technisch uitdagende experimenten. Dit protocol beschrijft een eenvoudige methode voor grove schatting van synaptische multipliciteit met behulp van hele-cel patch-klem elektrofysiologie.Deze methode kan worden toegepast op elke soort en hersenen gebied om synaptische multipliciteit te onderzoeken...

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

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 ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

Electrophysiological and Morphological Characterization of Neuronal Microcircuits in Acute Brain Slices Using Paired Patch-Clamp Recordings

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with simultaneous intracellular biocytin filling allows a correlated analysis of their structural and functional properties. With this method it is possible to identify and characterize both pre- and postsynaptic neurons by their morphology and electrophysiological response pattern. Paired recordings allow studying the connectivity patterns between these neurons as well as the properties of both chemical and electrical synaptic transmission. Here, we give a step-by-step description of the procedures required to obtain reliable paired recordings together with an optimal recovery of the neuron morphology. We will describe how pairs of neurons connected via chemical synapses or gap junctions are identified in brain slice preparations. We will outline how neurons are reconstructed to obtain their 3D morphology of the dendritic and axonal domain and how synaptic contacts are identified and localized. We will also discuss the caveats and limitations of the paired recording technique, in particular those associated with dendritic and axonal truncations during the preparation of brain slices because these strongly affect connectivity estimates. However, because of the versatility of the paired recording approach it will remain a valuable tool in characterizing different aspects of synaptic transmission at identified neuronal microcircuits in the brain.

Patch-clamp recordings and simultaneous intracellular biocytin filling of synaptically coupled neurons in acute brain slices allow a correlated analysis of their structural and functional properties. The aim of this protocol is to describe the essential technical steps of electrophysiological recording from neuronal microcircuits and their subsequent morphological analysis.

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with ...

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 ...

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.

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze ...

Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ganglion cells and project it to the visual cortex. In addition, relay neurons receive top-down excitation from the cortex. The two main excitatory inputs to the relay neurons differ in several aspects. Each relay neuron receives input from only a few retinogeniculate synapses, which are large terminals with many release sites. This is reflected by the comparably strong excitation, the relay neurons receive, from retinal ganglion cells. Corticogeniculate synapses, in contrast, are simpler with few release sites and weaker synaptic strength. The two synapses also differ in their synaptic short-term plasticity. Retinogeniculate synapses have a high release probability and consequently display a short-term depression. In contrast, corticogeniculate synapses have a low release probability. Corticogeniculate fibers traverse the reticular thalamic nuclei before entering the lateral geniculate nucleus. The different locations of the reticular thalamic nucleus (rostrally from the lateral geniculate nucleus) and optic tract (ventro-laterally from the lateral geniculate nucleus) allow stimulating corticogeniculate or retinogeniculate synapses separately with extracellular stimulation electrodes. This makes the lateral geniculate nucleus an ideal brain area where two excitatory synapses with very different properties impinging onto the same cell type, can be studied simultaneously. Here, we describe a method to investigate the recording from relay neurons and to perform detailed analysis of the retinogeniculate and corticogeniculate synapse function in acute brain slices. The article contains a step-by-step protocol for the generation of acute brain slices of the lateral geniculate nucleus and steps for recording activity from relay neurons by stimulating the optic tract and corticogeniculate fibers separately.

Here, we present protocols for the preparation of acute brain slices containing the lateral geniculate nucleus and the electrophysiological investigation of retinogeniculate and corticogeniculate synapses function. This protocol provides an efficient way to study synapses with the high-release and low-release probability in the same acute brain slices.

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ...

Whole-Cell Patch Clamp Electrophysiology: A Method to Study Electrical Properties of Neurons

Source: Roy, B., et al. <a href="https://www.jove.com/v/50551/" target="_blank">Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells</a>. J. Vis. Exp. (2013).
This protocol describes a technique called whole-cell patch clamp electrophysiology, which is a method to study electrical properties of Mauthner neurons and other reticulospinal cells in zebrafish embryos.

Source: Roy, B., et al. Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells. J. Vis. Exp. (2013).
This protocol describes a technique called ...

JoVE Encyclopedia of Experiments

Biology

Danio rerio (zebrafish)

Embryo

Studying the Excitability of Fluorescent Neurons using a Whole-Cell Patch Clamp

Source: Briz, C. G., et al., <a href="https://app.jove.com/v/55139">In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity.</a> J. Vis. Exp. (2017)
The video demonstrates a method to study the excitability of neuronal cells expressing fluorescent protein using a whole-cell patch current clamp. In the current-clamp mode, action potentials indicative of neuronal excitability are generated.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Electrophysiology
NOTE: The ...

Neurophysiology

Electrophysiology Techniques

Establishing a Whole-Cell Voltage-Clamp Configuration for Electrophysiological Recordings in Brain Slices

Source: Sunstrum, J. K., et al. <a href="https://app.jove.com/v/59461">Evaluation of synaptic multiplicity using whole-cell patch-clamp electrophysiology.</a> J. Vis. Exp. (2019)
This video demonstrates positioning a fine-pointed pipette near brain slices, forming a seal with a neuronal cell. Suction breaks the cell membrane, thereby creating a whole-cell patch clamp that enables current measurements for electrophysiology experiments.

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

Samenvatting

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