The authors thank Bryan Krause for technical support and guidance on this project.
This work was supported by the International Anesthesia Research Society (IMRA to AR), National Institutes of Health (R01 GM109086 to MIB), and the Department of Anesthesiology, School of Medicine and Public Health, University of Wisconsin, Madison, WI, USA.

All procedures involving animals described in this protocol were approved by the University of Wisconsin-Madison School of Medicine and Public Health Animal Care and Use Committee.
1. Breeding mice to express fluorescent reporter protein in interneuron subpopulations
<ol>
	<li>Pair homozyogous, Cre-dependent tdTomato male mouse with either homozygous SOM-Cre female or homozygous PV-Cre female mouse. 
	NOTE: Other specific neuronal populations may be targeted by using the appropriate Cre lines.</li>
	<li>Allow heterozygous offspring to mature to at least 3 weeks of age before proceeding. For experiments described here, genotyping is not necessary, as homozygous parents produce offspring that are all heterozygous for both cell type-specific Cre recombinase and Cre-dependent reporter alleles.</li>
</ol>
2. Performing unilateral stereotaxic injection of viral construct
<ol>
	<li>Adjust settings of the micropipette puller for injection pipettes as indicated in the instrument user manual (see Table 1 for recommended settings). Pull the glass micropipette.</li>
	<li>Break the tip of the sharp end of the pipette such that the tip diameter is approximately 30 &#181;m with minimal taper over several millimeters.</li>
	<li>Using previously documented procedures, decide on the appropriate titer and volume of virus to be injected. In the experiments described here, 1.0 &#956;L (titer: 3.1-5.7 TU/mL) injected unilaterally produced good results.</li>
	<li>Backfill the full volume of the pipette with mineral oil. Load the pipette onto the microsyringe and flow a small amount of mineral oil through the tip to ensure the tip is not clogged.</li>
	<li>Frontfill at least 1.0 &#956;L of viral construct. The recombinant adeno-associated viral vector used in these experiments was AAV2-hSyn-hChR2(H134R)-EYFP.</li>
	<li>Arrange sterile drape in a surgical area. Sterilize tools for stereotaxic procedure and place it on the drape.</li>
	<li>Anesthetize SOM-tdTomato or PV-tdTomato heterozygous animal using isoflurane (3% for induction, 1.5-2% for maintenance) and oxygen mixture. Periodically confirm surgical level of anesthesia with toe pinch throughout surgery. Ensure animal does not move beyond the surgical plan of anesthesia by monitoring respirations every 10-15 min.</li>
	<li>Shave the top of the animal&#8217;s head. Apply 70% isopropyl alcohol and iodine-based solution liberally to surgical area and ophthalmic ointment to eye sockets to prevent drying of the membrane. Administer bupivacaine/lidocaine (1:1 ratio, 1.0 mg/kg) subcutaneously to surgical site for local anesthetic.</li>
	<li>Fit the animal into stereotaxic frame.</li>
	<li>Use scalpel to make incision along sagittal axis of skin overlying the dorsal surface of the skull. Retract skin using forceps. Hydrate skull with 0.9% saline as necessary.</li>
	<li>On the surface of the skull, lightly mark the intersection of anterior and lateral coordinates with a cross in pencil. Drill a hole at the appropriate coordinates in the transverse plane (in mm relative to Bregma, for cingulate cortex (Cg) injection: anterior 0.2, lateral 0.3; for posterior thalamus (Po) injection: posterior 2.25, lateral 3.4). 
	NOTE: Markings should extend beyond the boundaries of the burr hole to provide guidance for accurate placement of pipette.</li>
	<li>Turn on and balance the air table.</li>
	<li>Reposition electrode manipulator of the stereotaxic frame at 0&#176; for injections into Cg, or 45&#176; in the coronal plane for injections into Po.</li>
	<li>Attach the syringe pump to the electrode manipulator. Attach the microsyringe pump controller to the syringe pump.</li>
	<li>Navigate pipette tip near (but not touching) the surface of the brain, at the intersection of markings created in Step 2.11. Advance the pipette at approximately 1 mm/s along its longitudinal axis into the brain either 0.9 mm (injection in Cg) or 3.1 mm (injection in Po). Wait for 10 min before proceeding.</li>
	<li>Inject 1.0 &#181;L of viral construct over a period of 10 min (100 nL/min). If welling of virus from pipette insertion site is observed, slow the injection rate to 50 nL/min.</li>
	<li>After injection, wait for 10 min before slowly retracting the injection pipette.</li>
	<li>Suture to close the scalp incision and administer 2-5 mg/kg meloxicam subcutaneously.</li>
	<li>Discontinue isoflurane and monitor animal during emergence from anesthesia. Allow to recover according to procedures described by Institution&#8217;s Animal Care and Use Committee, including further administration of analgesics.</li>
</ol>
3. Preparation of acute brain slices
<ol>
	<li>Allow at least 3 weeks for the expression of viral construct before harvesting tissue.</li>
	<li>Prepare 1 L of artificial cerebral spinal fluid for slicing procedure (slicing artificial cerebral spinal fluid, sACSF). See Table 2 for ingredients.</li>
	<li>Throughout the slicing procedure, supply sACSF with dissolved 95% O2/5% CO2 mixture, delivered via gas dispersion tube.</li>
	<li>Prepare ice cold bath for vibrating blade microtome. Mount ice-cold specimen stage onto microtome and fix sapphire blade in place for tissue sectioning.</li>
	<li>Anesthetize mouse with 3% isoflurane and oxygen until loss of righting reflex.</li>
	<li>Decapitate the mouse using guillotine and immediately submerge head in 4&#176;C sACSF. To preserve the health of the tissue, complete the following steps as swiftly as possible.</li>
	<li>Open skull cavity by making a small incision at the base of the skull and gently removing each skull plate. Gently remove underlying dura mater.</li>
	<li>While the brain is still in the skull cavity, use the razor blade to remove the cerebellum. Make a second vertical cut along the sagittal plane in the left hemisphere, just lateral to the midline.</li>
	<li>Prepare tissue block for sectioning.
	<ol>
		<li>Gently lift the brain from the skull cavity. Place the brain on the filter paper with the flat, sagittal plane down. Guide the filter paper over the blocking template and align the brain to the underlying template outline (Supplementary Figure 1).</li>
		<li>Make two parallel cuts in the coronal plane as indicated by the lines on the template. Add a small drop of sACSF to keep filter paper wet, if necessary.</li>
		<li>Place the tissue block in 4&#176;C sACSF briefly while step 3.8.4 is conducted.</li>
		<li>Apply a small amount of super glue to the ice-cold specimen stage.</li>
		<li>Lift the tissue block from cold sACSF. Use the corner of an absorbent towel to wick away excess sACSF. Glue the posterior coronal plane of the tissue block to the specimen stage, with the dorsal surface of the brain facing the sapphire blade.</li>
	</ol>
	</li>
	<li>Collect 500 &#181;m thick coronal brain slices. Place slices of interest on nylon mesh (Supplementary Figure 2) in 34 &#176;C sACSF and allow the container to reach room temperature. 
	NOTE: For experiments described here, electrophysiological recordings were collected from a coronal section centered approximately 2.25 mm posterior to bregma to study a non-primary sensory area, medial secondary visual cortex (V2MM).</li>
</ol>
4. Preparation of experimental artificial cerebral spinal fluid (eACSF) bags containing dissolved volatile anesthetic isoflurane
<ol>
	<li>Prepare 300 mL of a stock mixture of 3.0% isoflurane.
	<ol>
		<li>In a sealed polytetrafluoroethylene gas bag, add ~100 mL of 95% O2/5% CO2 gas mixture to 20-30 mL of liquid isoflurane and a small amount of 0.9% saline. Wait at least for 30 min to allow equilibration of isoflurane between liquid and gas phases.</li>
		<li>Determine the amount of saturated isoflurane gas, Vsat, to add to the stock bag using the following equation: 
		<img alt="Equation 1" src="/files/ftp_upload/61333/61333equ01.jpg" /> 
		where P%stock is the target composition of the stock gas (3% in this case), Vstock is the final volume of the stock gas bag, Pisoflurane is the partial pressure of isoflurane at room temperature (~240 mmHg), and Ptotal is the atmospheric pressure (~760 mmHg).</li>
		<li>Add the calculated amount of saturated gas to an empty gas bag and fill the bag with a volume of 95% O2/5% CO2 gas mixture to bring the total volume of stock bag to 300 mL.</li>
	</ol>
	</li>
	<li>Prepare 2 L of artificial cerebral spinal fluid for perfusion of the slice during the experiment (experimental ACSF, eACSF). See Table 2 for ingredients. Dissolve 95% O2/5% CO2 gas mixture into solution.</li>
	<li>Prepare two separate bags of Control and Isoflurane solutions.
	<ol>
		<li>To an empty polytetrafluoroethylene gas bag, add 600 mL eACSF and 600 mL of 95% O2/5% CO2 gas mixture. Label this bag as Control.</li>
		<li>To another empty polytetrafluoroethylene gas bag, add 300 mL of eACSF. Label this bag as Isoflurane.</li>
		<li>Choose a physiologically relevant equilibrated gas phase concentration of isoflurane. Experiments were conducted using gas concentrations equivalent to 1.3% isoflurane. Mice lose righting reflex, and presumably consciousness, at 0.9% inhaled isoflurane.</li>
		<li>Use the following equation to calculate the equivalent gas phase concentration at room temperature, P%(Troom)31: 
		<img alt="Equation 2" src="/files/ftp_upload/61333/61333equ02.jpg" /> 
		where P%(Tbody) is the physiologically relevant gas phase concentration chosen in Step 4.3.3, Troom is 25 &#176;C, and Tbody is 37 &#176;C.</li>
		<li>Use the following equation to determine volume of gas from stock gas bag, Vstock, to add to the Isoflurane solution. 
		<img alt="Equation 3" src="/files/ftp_upload/61333/61333equ03.jpg" /> 
		where Vsolution is the volume of eACSF in ISOFLURANE bag (300 mL), P%(Troom) is entered from equation (2), &#955; is the saline/gas Ostwald partition coefficient of isoflurane (&#955; = 1.232), and P%stock is the gas phase concentration of the stock gas bag (P%stock = 3.0%).</li>
		<li>To the Isoflurane solution bag, add the volume of gas from the stock gas bag, Vstock, calculated in Step 4.3.5.</li>
		<li>To the Isoflurane solution bag, add a volume of 95% O2/5% CO2 gas mixture to bring the total volume of gas in the Isoflurane solution bag to 300 mL.</li>
	</ol>
	</li>
	<li>Shake both Control and Isoflurane bags on shaker for at least 1 h to allow isoflurane phase equilibration.</li>
	<li>After all data has been collected, the correct concentration may be verified by using an anesthetic gas monitor to measure equilibrated gas concentration of isoflurane above the remaining solution in bag.</li>
	<li>Report experimental concentrations of volatile gases in aqueous units, as millimolar concentrations are more robust to changes in temperature. Use the following equation to convert room temperature gas phase concentration, P%(Troom), to equivalent aqueous concentration (Caqueous, in mM)31: 
	<img alt="Equation 4" src="/files/ftp_upload/61333/61333equ04.jpg" /> 
	where &#945; is the saline/gas Bunsen partition coefficient for isoflurane at 25&#176;C32.</li>
</ol>
5. Preparation of hardware and software for multi-channel recordings
<ol>
	<li>Set up 16-channel data acquisition system according to manufacturer instructions. 
	NOTE: Several commercially available amplifiers and data acquisition systems can be used to collect multi-channel recordings. In the experiments described here, analog signals are delivered via an electrode reference panel to two amplifiers, where they are amplified (2000x) and filtered (0.1-10kHz). Analog inputs to the data acquisition system are digitized at 40kHz.</li>
	<li>Fasten the appropriate 16-channel headstage adaptor to a microscope micromanipulator. Orient the adaptor such that the female connector ports are facing downward.</li>
	<li>Adjust the angle of operation of this micromanipulator such that it is oriented downward toward the recording chamber, at an angle approximately 70&#176; relative to horizontal.</li>
	<li>Connect the headstage input to a 16 x 1 probe for in vitro electrophysiology via the headstage adaptor anchored to the micromanipulator.</li>
	<li>Connect the headstage output connector to the data acquisition system.</li>
	<li>Install appropriate software for data acquisition. Configure 15 input channels to correspond to input signals from the first 15 multi-channel probe contacts. Configure the remaining channel to receive input from the intracellular electrode. 
	NOTE: Take care to consider electrode and adaptor maps when collecting and analyzing data, to ensure the appropriate signal corresponds to the electrode contact from which it was collected.</li>
</ol>
6. Configuration of light stimulation protocols
<ol>
	<li>Set up light delivery system and install the accompanying software.</li>
	<li>Open the software. Choose hardware wiring configuration in which a Trigger Source (Digital/TTL Out) provides Trigger In signal to the light delivery system, and the light delivery system provides Trigger Out signal to a 470 nm LED.</li>
	<li>Mount high-power objective lens. Using digital camera, calibrate high-power objective for use with light delivery system.</li>
	<li>Create new profile sequence of light stimulation profiles.
	<ol>
		<li>Create a pattern of choice. In the experiments described here, a circle of diameter 150 &#181;m is used to allow layer-specific activation of axon terminals.</li>
		<li>To construct a profile sequence, copy and paste this profile for each of any number of trials.</li>
		<li>Create a waveform list that contains waveforms of any light intensity, pulse duration, or pulse number.</li>
		<li>Randomly assign waveforms to each profile. Each profile with its assigned waveform corresponds to one trigger pulse from a Digital TTL input, or one trial.</li>
		<li>Save the profile sequence.</li>
	</ol>
	</li>
	<li>In the data acquisition software, create a new protocol.
	<ol>
		<li>Set the number of trials to equal the number of profiles in the profile sequence just created.</li>
		<li>Choose signal inputs to match those configured in Step 5.6. Configure a protocol that provides a single digital TTL output, recording from these 16 input channels for an appropriate amount of time before and after the digital trigger.</li>
	</ol>
	</li>
</ol>
7. Placing multi-channel probe in ex vivo brain tissue slice
<ol>
	<li>Perfuse bubbled eACSF (not in sealed bags) at 3-6 mL/min.</li>
	<li>Transfer the brain slice containing area of interest onto mesh grid in microscope perfusion chamber. Anchor with platinum harp (see Supplementary Figure 3).</li>
	<li>Rotate mesh grid such that the line of electrode contacts on the distal end of the multi-channel probe is approximately perpendicular to the pial surface.</li>
	<li>Under broadfield illumination and under fine control of the micromanipulator, lower the multi-channel probe toward the surface of the slice.</li>
	<li>Rotate the filter cube turret to engage the appropriate filter cube for visualization of the fluorescent reporter protein expressed in axon terminals of cortical afferents. If necessary, rotate the slice to more precisely align the probe with the pial surface.</li>
	<li>Position the probe just above the plane of the slice, ~200 &#181;m short of the final target position along the x-axis, leaving at least one channel outside the boundary of the area of tissue being recorded</li>
	<li>Slowly insert the probe into the slice by moving the manipulator along its longitudinal axis. To minimize damage to the tissue, only advance the probe to the extent that the sharp tips are just visible below the tissue surface. This will minimize damage to the tissue while still ensuring the electrode contacts are in contact with the tissue.</li>
</ol>
8. Patch clamping targeted neurons and obtaining whole-cell configuration
<ol>
	<li>Switch eACSF source to bagged Control solution.</li>
	<li>Identify fluorescently labeled cell for targeted patch clamp recording.
	<ol>
		<li>Restrict the aperture iris diaphragm to the smallest diameter. Engage a low-power objective lens and bring the tissue into focus.</li>
		<li>Center the light over an area of tissue adjacent to (but not overlapping) the multi-channel probe.</li>
		<li>Engage the high-power (40x or 60x) water immersion objective, using caution to avoid contact between the multi-channel probe and objective lens.</li>
		<li>Rotate the filter cube turret to engage the appropriate filter set to allow imaging of cells expressing Cre-dependent fluorescent marker.</li>
		<li>Identify a fluorescently labeled cell as a target for patch clamp recording. Raise the objective lens to create ample space to lower a patch pipette.</li>
	</ol>
	</li>
	<li>Load a patch pipette (see Table 1) with internal solution (Table 2) and mount pipette into electrode holder. Using 1 mL syringe, apply positive pressure corresponding to ~0.1mL air.</li>
	<li>Lower patch pipette into the solution. Bring the pipette tip into focus under visual guidance.</li>
	<li>Obtain whole-cell recording from the targeted cell using the steps previously demonstrated33.</li>
	<li>If planning to assess changes to intrinsic properties of the cell (e.g., input resistance, action potential firing rate in response to current steps), conduct these recordings. Otherwise, move to axon stimulation protocol in below.</li>
</ol>
9. Layer-specific optogenetic activation of axon terminals
<ol>
	<li>Manipulate field of view in the x-y plane to align light stimulation profile with desired location on slice.</li>
	<li>Load light stimulus protocol and prepare the light delivery system to receive a digital TTL pulse.</li>
	<li>Optogenetically activate axon terminals while simultaneously recording extracellular field potentials and intracellular membrane fluctuations.</li>
	<li>Switch eACSF source to Isoflurane solution and wash drug in for 15 min. If necessary, collect spontaneous recordings during the wash-in.</li>
	<li>Repeat step 9.2-9.3.</li>
	<li>Switch eACSF source to Control solution and wash drug out for 20 min. If necessary, collect spontaneous recordings during wash-out.</li>
	<li>Repeat step 9.2-9.3.</li>
</ol>

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Partition+coefficients+of+volatile+anesthetics+in+aqueous+electrolyte+solutions+at+various+temperatures.">Partition coefficients of volatile anesthetics in aqueous electrolyte solutions at various temperatures.</a> Anesthesiology. 89 (4), 1032-1035 (1998).</li><li>Au - Segev, A., Au - Garcia-Oscos, F., Au - Kourrich, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole-cell+Patch-clamp+Recordings+in+Brain+Slices.">Whole-cell Patch-clamp Recordings in Brain Slices.</a> Journal of Visualized Experiments. (112), e54024 (2016).</li><li>Lin, J. Y., Lin, M. Z., Steinbach, P., Tsien, R. 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The authors have nothing to disclose.

In this manuscript, a protocol for evaluating intra- and extracellular responses to selectively activated afferent pathways in ex vivo brain slices is described.
The use of optogenetic tools and parallel recording schemes allows investigators to probe responses of local populations to afferent inputs from distant brain regions, while recording simultaneously from targeted populations of interneurons. The use of optogenetic technology allows for axon terminals of afferent projections to be pres...

Volatile anesthetics have been used ubiquitously in a variety of clinical and academic settings for more than a century. Distinct classes of anesthetics have unique, often non-overlapping molecular targets1,2,3, yet nearly all of them produce unconsciousness. While their behavioral effects are quite predictable, the mechanisms by which anesthetics induce loss of consciousness are largely unknown. Anesthetics may ultimately influence both the level and contents of consciousness via actions on corticothalamic circuits, disrupting integration of information throughout the cortical hierarchy4,5,6,7,8,9. More broadly, modulation of corticothalamic circuits may play a role in experimentally10 or pharmacologically11 altered states of consciousness, and may also be implicated in sleep12 and in pathophysiological disorders of consciousness13,14.
The elusiveness of the mechanisms underlying loss and return of consciousness during anesthesia may be attributed partially to non-linear, synergistic actions of anesthetics at the cellular, network, and systems levels15. Isoflurane, for example, suppresses activity within the selected brain regions16,17,18, impairs connectivity between distant brain regions19,20,21,22,23, and diminishes synaptic responses in a pathway-specific manner24,25. Which effects of anesthetics, from the molecular to the systems level, are necessary or sufficient to effect loss of consciousness remains unclear. In addition to substantive clinical investigations of consciousness using non-invasive techniques19,20,26, it is important that experimentalists seek to disentangle the distinct cellular and network interactions that subserve the conscious experience.
By simplifying the complex interactions found in the intact brain, ex vivo brain slices allow the study of isolated components of the brain&#8217;s dynamic systems9. A reduced slice preparation combines the benefits of relatively intact anatomical structures of local neural circuits with the versatility of in vitro manipulations. However, until recently, methodological constraints have precluded the study of synaptic and circuit properties of long-range inputs in brain slices27,28; the tortuous path of corticothalamic fiber tracts made activation of independent afferent pathways all but impossible by electrical stimulation.
Investigating the effects of anesthetic agents on the brain slice preparations presents additional challenges. Absent an intact respiratory and circulatory system, anesthetic agents must be bath-applied, and concentrations carefully matched to estimated effect site concentrations. For many intravenous anesthetic agents, the slow rate of equilibration in the tissue renders traditional pharmacological investigations laborious29,30. Investigating the effects volatile gas anesthetics in ex vivo preparations is more tractable, but also presents challenges. These include converting inhaled partial pressure doses to aqueous concentrations, and the need for a modified delivery system of the drug to the tissue via artificial cerebral spinal fluid31.
Here, methods are described by which investigators may capitalize on the well-documented physicochemical properties of the volatile anesthetic isoflurane for drug delivery to ex vivo brain slices, activate pathway- and layer-specific inputs to a cortical area of interest with high spatiotemporal resolution, and conduct simultaneous laminar recordings and targeted patch clamp recordings from select populations of neurons. Combined, these procedures allow investigators to measure volatile anesthetic-induced changes in several observable electrophysiological response properties, from the synaptic to local network level.

<table><tbody><tr><td>2.5x broadfield objective lens</td><td>Olympus</td><td>MPLFLN2.5X</td><td></td></tr><tr><td>40x water immersion objective lens</td><td>Olympus</td><td>LUMPLFLN40XW</td><td></td></tr><tr><td>95% O2/5% CO2 mixture</td><td>Airgas</td><td>Z02OX95R2003045</td><td></td></tr><tr><td>A16 probe</td><td>NeuroNexus</td><td>A16x1-2mm-100-177-A16</td><td>16-channel probe</td></tr><tr><td>AAV2-hSyn-hChR2(H134R)-EYFP</td><td>Karl Deisseroth Lab, UNC Vector Core</td><td></td><td></td></tr><tr><td>Anesthetic gas monitor (POET II)</td><td>Criticare</td><td>602-3A</td><td></td></tr><tr><td>ATP, Magnesium Salt</td><td>Sigma Aldrich</td><td>A9187</td><td>intracellular solution</td></tr><tr><td>B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J</td><td>The Jackson Laboratory</td><td>007914</td><td>Cre-dependent tdTomato mouse</td></tr><tr><td>B6;129P2-Pvalbtm1(cre)Arbr/J</td><td>The Jackson Laboratory</td><td>008069</td><td>PV-Cre mouse</td></tr><tr><td>Belly Dancer Shaker</td><td>Thomas Scientific</td><td>1210H86-TS</td><td>for equilibration of sealed gas bags</td></tr><tr><td>Betadine solution</td><td>Generic brand</td><td></td><td></td></tr><tr><td>Bleach</td><td>Generic brand</td><td></td><td>for silver chloriding patch clamp electrode</td></tr><tr><td>Bupivicaine</td><td></td><td></td><td></td></tr><tr><td>Calcium Chloride (CaCl2)</td><td>Dot Scientific</td><td>DSC20010</td><td>ACSF</td></tr><tr><td>Capillary glass (patch clamp recordings)</td><td>King Precision Glass, Inc.</td><td>KG-33</td><td>Borosilicate, ID: 1.1mm, OD: 1.7mm, Length: 90.0mm</td></tr><tr><td>Capillary glass (viral injections)</td><td>Drummond Scientific Company</td><td>3-000-203-G/X</td><td>3.5&quot;</td></tr><tr><td>Control of junior micromanipulator</td><td>Luigs and Neumann</td><td>SM8</td><td>for control of junior micromanipulator</td></tr><tr><td>Control of manipulators and shifting table</td><td>Luigs and Neumann</td><td>SM7</td><td>for control of multichannel electrode and shifting table</td></tr><tr><td>Digidata 1440A + Clampex 10</td><td>Molecular Devices</td><td>1440A</td><td>Digitizer and software</td></tr><tr><td>E-3603 tubing</td><td>Fisher Scientific</td><td>14171208</td><td>for delivery of 95% O2/5% CO2 gas mixture to incubation chamber + application of pressure during patch clamping</td></tr><tr><td>EGTA</td><td>Dot Scientific</td><td>DSE57060</td><td>intracellular solution</td></tr><tr><td>ERP-27 EEG Reference/Patch Panel</td><td>Neuralynx</td><td>Retired</td><td></td></tr><tr><td>Filling needle</td><td>World Precision Instruments</td><td>50821912</td><td>for filling patch clamp pipettes</td></tr><tr><td>Filter cube for imaging EYFP</td><td>Olympus</td><td>U-MRFPHQ</td><td></td></tr><tr><td>Filter paper</td><td>Fisher Scientific</td><td>09801E</td><td>lay over slice template during preparation of tissue block</td></tr><tr><td>Flaming/Brown micropipette puller</td><td>Sutter Instrument</td><td>P-1000</td><td>2.5x2.5 Box filament</td></tr><tr><td>Gas dispersion tube</td><td>Sigma Aldrich</td><td>CLS3953312C</td><td></td></tr><tr><td>Glass syringe (100 mL)</td><td>Sigma Aldrich</td><td>Z314390</td><td>for filling gas-sealed bags</td></tr><tr><td>Gluconic Acid, Potassium Salt (K-gluconate)</td><td>Dot Scientific</td><td>DSG37020</td><td>intracellular solution</td></tr><tr><td>Glucose</td><td>Dot Scientific</td><td>DSG32040</td><td>ACSF</td></tr><tr><td>GTP, Sodium Salt</td><td>Sigma Aldrich</td><td>G8877</td><td>intracellular solution</td></tr><tr><td>Headstage-probe adaptor</td><td>NeuroNexus</td><td>A16-OM16</td><td>adaptor to connect 16-channel probe to headstage input</td></tr><tr><td>Hemostatic Forceps</td><td>VWR International</td><td>76192-096</td><td></td></tr><tr><td>HEPES</td><td>Dot Scientific</td><td>DSH75030</td><td>ACSF,intracellular solution</td></tr><tr><td>HS-16 Headstage</td><td>Neuralynx</td><td>Retired</td><td></td></tr><tr><td>Isoflurane</td><td>Patterson Veterinary</td><td>07-893-1389</td><td></td></tr><tr><td>Isopropyl alcohol (70%)</td><td>VWR International</td><td>101223-746</td><td></td></tr><tr><td>Junior micromanipulator</td><td>Luigs and Neumann</td><td>210-100 000 0090-R</td><td>for manipulation of patch clamp electrode</td></tr><tr><td>LED Light Source Control Module</td><td>Mightex</td><td>BLS-PL02_US</td><td>optogenetic light source control</td></tr><tr><td>Lidocaine</td><td></td><td></td><td></td></tr><tr><td>Lynx-8 Amplifier</td><td>Neuralynx</td><td>Retired</td><td></td></tr><tr><td>Lynx-8 Power Supply</td><td>Neuralynx</td><td>Retired</td><td></td></tr><tr><td>Magnesium Sulfate (MgSO4)</td><td>Dot Scientific</td><td>DSM24300</td><td>ACSF</td></tr><tr><td>mCherry, Texas Red filter cube</td><td>Chroma</td><td>49008</td><td>for imaging tdTomato fluorescent reporter</td></tr><tr><td>Meloxicam</td><td></td><td></td><td></td></tr><tr><td>Micropipette holder</td><td>Fisher Scientific</td><td>NC9044962</td><td></td></tr><tr><td>Microsyringe pump</td><td>World Precision Instruments</td><td>UMP3-4</td><td></td></tr><tr><td>Mineral oil</td><td>Generic brand</td><td></td><td></td></tr><tr><td>MultiClamp 700A</td><td>Molecular Devices/Axon Instruments</td><td>700A</td><td>Amplifier</td></tr><tr><td>Nitrogen (for air table)</td><td>Airgas</td><td>NI200</td><td></td></tr><tr><td>Nylon mesh</td><td>Fisher Scientific</td><td>501460083</td><td>stretched over horseshoe of flattened platinum wire, slice rest on top of this during recordings</td></tr><tr><td>Nylon, cut from pantyhose</td><td>Generic brand</td><td></td><td>small piece to create slice platform in incubation chamber, single fibers to create platinum harp</td></tr><tr><td>Ophthalmic ointment</td><td>Fisher Scientific</td><td>NC1697520</td><td></td></tr><tr><td>Pipette</td><td>Dot Scientific</td><td>307</td><td>For transferring tissue to rig</td></tr><tr><td>Platinum wire</td><td>VWR International</td><td>BT124000</td><td>2 cm, flattened, to make platinum harp</td></tr><tr><td>Polygon400</td><td>Mightex</td><td>DSI-E-0470-0617-000</td><td>optogenetic light delivery system, comes with PolyScan2 software</td></tr><tr><td>Potassium Chloride (KCl)</td><td>Dot Scientific</td><td>DSP41000</td><td>ACSF</td></tr><tr><td>Potassium Phosphate (KH2PO4)</td><td>Dot Scientific</td><td>DSP41200</td><td>ACSF</td></tr><tr><td>Razor blade</td><td>Fisher Scientific</td><td>12-640</td><td></td></tr><tr><td>Sapphire blade (for vibratome)</td><td>VWR International</td><td>100492-502</td><td></td></tr><tr><td>Scalpel blade</td><td>Santa Cruz Biotechnology, Inc.</td><td>sc-361445</td><td></td></tr><tr><td>Sealed gas bag</td><td>Fisher Scientific</td><td>109236</td><td></td></tr><tr><td>Shifting table for microscope</td><td>Luigs and Neumann</td><td>380FMU</td><td></td></tr><tr><td>Sodium Bicarbonate (HCO3-)</td><td>Dot Scientific</td><td>DSS22060</td><td>ACSF</td></tr><tr><td>Sodium Chloride (NaCl)</td><td>Dot Scientific</td><td>DSS23020</td><td>ACSF, intracellular solution</td></tr><tr><td>Ssttm2.1(cre)Zjh/J (SOM-IRES-Cre)</td><td>The Jackson Laboratory</td><td>013044</td><td>SOM-Cre mouse</td></tr><tr><td>Stereotaxic instrument</td><td>Kopf</td><td>Model 902</td><td>Dual Small Animal</td></tr><tr><td>Super glue</td><td>Staples</td><td>886833</td><td>to fix tissue block to specimen stage during slice preparation</td></tr><tr><td>Surgical drill</td><td>RAM Products Inc.</td><td>DIGITALMICROTORQUE</td><td>Microtorque II</td></tr><tr><td>Syringe (1 mL) with LuerLock tip</td><td>Fisher Scientific</td><td>309628</td><td>for application of pressure during patch clamping</td></tr><tr><td>Syringe (1 mL) with slip tip</td><td>WW Grainger, Inc.</td><td>19G384</td><td>for filling patch clamp pipettes</td></tr><tr><td>Syringe Filters</td><td>VWR International</td><td>66064-414</td><td></td></tr><tr><td>Upright microscope</td><td>Olympus</td><td>BX51</td><td></td></tr><tr><td>Vibrating microtome</td><td>Leica Biosystems</td><td>VT1000S</td><td></td></tr><tr><td>Wypall towels</td><td>Fisher Scientific</td><td>19-042-427</td><td></td></tr></tbody></table>

optogenetic activation of afferent pathways in brain slices and modulation of responses by volatile anesthetics

Anesthetics influence consciousness in part via their actions on thalamocortical circuits. However, the extent to which volatile anesthetics affect distinct cellular and network components of these circuits remains unclear. Ex vivo brain slices provide a means by which investigators may probe discrete components of complex networks and disentangle potential mechanisms underlying the effects of volatile anesthetics on evoked responses. To isolate potential cell type- and pathway-specific drug effects in brain slices, investigators must be able to independently activate afferent fiber pathways, identify non-overlapping populations of cells, and apply volatile anesthetics to the tissue in aqueous solution. In this protocol, methods to measure optogenetically-evoked responses to two independent afferent pathways to neocortex in ex vivo brain slices are described. Extracellular responses are recorded to assay network activity and targeted whole-cell patch clamp recordings are conducted in somatostatin- and parvalbumin-positive interneurons. Delivery of physiologically relevant concentrations of isoflurane via artificial cerebral spinal fluid to modulate cellular and network responses is described.

A timeline of steps described in the protocol is shown in Figure 1. Cortical inputs arriving from higher order cortical areas or from non-primary thalamic nuclei have partially overlapping terminal fields in layer 1 of non-primary visual cortex24. To isolate independent thalamocortical or corticocortical afferent pathways, a viral vector containing ChR2 and an eYFP fluorescent reporter into either Po or Cg was injected. Cells within the injection radius take up the vi...

Watch this Scientific Journal Video about Optogenetic Activation of Afferent Pathways in Brain Slices and Modulation of Responses by Volatile Anesthetics at JoVE.com

Optogenetic Activation of Afferent Pathways in Brain Slices and Modulation of Responses by Volatile Anesthetics

Ex vivo brain slices can be used to study the effects of volatile anesthetics on evoked responses to afferent inputs. Optogenetics are employed to independently activate thalamocortical and corticocortical afferents to non-primary neocortex, and synaptic and network responses are modulated with isoflurane.

Anesthetics influence consciousness in part via their actions on thalamocortical circuits. However, the extent to which volatile anesthetics affect ...

optogenetic-activation-afferent-pathways-brain-slices-modulation

Research

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Neuroscience

2.1K Views.  University of Wisconsin. Ex vivo brain slices can be used to study the effects of volatile anesthetics on evoked responses to afferent inputs. Optogenetics are employed to independently activate thalamocortical and corticocortical afferents to non-primary neocortex, and synaptic and network responses are modulated with isoflurane.

Video: Optogenetic Activation of Afferent Pathways in Brain Slices and Modulation of Responses by Volatile Anesthetics

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the interactions of these forebrain circuits has been aided recently by the development of optogenetic methods for light-mediated modulation of neuronal activity. Here, we describe a protocol for examining the functional organization of forebrain circuits&#160;in vitro using laser-scanning photostimulation of channelrhodopsin, expressed optogenetically via viral-mediated transfection. This approach also exploits the utility of cre-lox recombination in transgenic mice to target expression in specific neuronal cell types. Following transfection, neurons are physiologically recorded in slice preparations using whole-cell patch clamp to measure their evoked responses to laser-scanning photostimulation of channelrhodopsin expressing fibers. This approach enables an assessment of functional topography and synaptic properties. Morphological correlates can be obtained by imaging the neuroanatomical expression of channelrhodopsin expressing fibers using confocal microscopy of the live slice or post-fixed tissue. These methods enable functional investigations of forebrain circuits that expand upon more conventional approaches.

We describe a method for characterizing the functional topography and synaptic properties of forebrain circuits using an optogenetic approach to photostimulate neuronal populations&#160;in vitro.

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the ...

Optogenetic Manipulation of Neuronal Activity to Modulate Behavior in Freely Moving Mice

Optogenetic modulation of neuronal circuits in freely moving mice affects acute and long-term behavior. This method is able to perform manipulations of single neurons and region-specific transmitter release, up to whole neuronal circuitries in the central nervous system, and allows the direct measurement of behavioral outcomes. Neurons express optogenetic tools via an injection of viral vectors carrying the DNA of choice, such as Channelrhodopsin2 (ChR2). Light is brought into specific brain regions via chronic optical implants that terminate directly above the target region. After two weeks of recovery and proper tool-expression, mice can be repeatedly used for behavioral tests with optogenetic stimulation of the neurons of interest.
Optogenetic modulation has a high temporal and spatial resolution that can be accomplished with high cell specificity, compared to the commonly used methods such as chemical or electrical stimulation. The light does not harm neuronal tissue and can therefore be used for long-term experiments as well as for multiple behavioral experiments in one mouse. The possibilities of optogenetic tools are nearly unlimited and enable the activation or silencing of whole neurons, or even the manipulation of a specific receptor type by light.
The results of such behavioral experiments with integrated optogenetic stimulation directly visualizes changes in behavior caused by the manipulation. The behavior of the same animal without light stimulation as a baseline is a good control for induced changes. This allows a detailed overview of neuronal types or neurotransmitter systems involved in specific behaviors, such as anxiety. The plasticity of neuronal networks can also be investigated in great detail through long-term stimulation or behavioral observations after optical stimulation. Optogenetics will help to enlighten neuronal signaling in several kinds of neurological diseases.

With optogenetic manipulation of specific neuronal populations or brain regions, behavior can be modified with high temporal and spatial resolution in freely moving animals. By using different optogenetic tools in combination with chronically implanted optical fibers, a variety of neuronal modulations and behavioral testing can be performed.

Optogenetic modulation of neuronal circuits in freely moving mice affects acute and long-term behavior. This method is able to perform manipulations of ...

Behavior

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices

Knowledge of cell-type specific synaptic connectivity is a crucial prerequisite for understanding brain-wide neuronal circuits. The functional investigation of long-range connections requires targeted recordings of single neurons combined with the specific stimulation of identified distant inputs. This is often difficult to achieve with conventional and electrical stimulation techniques, because axons from converging upstream brain areas may intermingle in the target region. The stereotaxic targeting of a specific brain region for virus-mediated expression of light-sensitive ion channels allows selective stimulation of axons originating from that region with light. Intracerebral stereotaxic injections can be used in well-delimited structures, such as the anterior thalamic nuclei, in addition to other subcortical or cortical areas throughout the brain.
Described here is a set of techniques for precise stereotaxic injection of viral vectors expressing channelrhodopsin in the mouse brain, followed by photostimulation of axon terminals in the brain slice preparation. These protocols are simple and widely applicable. In combination with whole-cell patch clamp recording from a postsynaptically connected neuron, photostimulation of axons allows the detection of functional synaptic connections, pharmacological characterization, and evaluation of their strength. In addition, biocytin filling of the recorded neuron can be used for post-hoc morphological identification of the postsynaptic neuron.

This protocol describes a set of methods to identify the cell-type specific functional connectivity of long-range inputs from distant brain regions using optogenetic stimulations in ex vivo brain slices.

Knowledge of cell-type specific synaptic connectivity is a crucial prerequisite for understanding brain-wide neuronal circuits. The functional ...

Combined In Vivo Anatomical and Functional Tracing of Ventral Tegmental Area Glutamate Terminals in the Hippocampus

Optogenetic modulation of neuron sub-populations in the brain has allowed researchers to dissect neural circuits in vivo and ex vivo. This provides a premise for determining the role of neuron types within a neural circuit, and their significance in information encoding relative to learning. Likewise, the method can be used to test the physiological significance of two or more connected brain regions in awake and anesthetized animals. The current study demonstrates how VTA glutamate neurons modulate the firing rate of putative pyramidal neurons in the CA1 (hippocampus) of anesthetized mice. This protocol employs adeno-associated virus (AAV)-dependent labeling of VTA glutamate neurons for the tracing of VTA presynaptic glutamate terminals in the layers of the hippocampus. Expression of light-controlled opsin (channelrhodopsin; hChR2) and fluorescence protein (eYFP) harbored by the AAV vector permitted anterograde tracing of VTA glutamate terminals, and photostimulation of VTA glutamate neuron cell bodies (in the VTA). High-impedance acute silicon electrodes were positioned in the CA1 to detect multi-unit and single-unit responses to VTA photostimulation in vivo. The results of this study demonstrate the&#160;layer-dependent distribution of presynaptic VTA glutamate terminals in the hippocampus (CA1, CA3, and DG). Also, the photostimulation of VTA glutamate neurons increased the firing and burst rate of putative CA1 pyramidal units in vivo.

The current protocol demonstrates a simple method for tracing of ventral tegmental area (VTA) glutamate projections to the hippocampus. Photostimulation of VTA glutamate neurons was combined with CA1 recording to demonstrate how VTA glutamate terminals modulate CA1 putative pyramidal firing rate in vivo.

Optogenetic modulation of neuron sub-populations in the brain has allowed researchers to dissect neural circuits in vivo and ex vivo. This provides a ...

Injections of AAV Vectors for Optogenetics in Anesthetized and Awake Behaving Non-Human Primate Brain

Optogenetic techniques have revolutionized neuroscience research and are poised to do the same for neurological gene therapy. The clinical use of optogenetics, however, requires that safety and efficacy be demonstrated in animal models, ideally in non-human primates (NHPs), because of their neurological similarity to humans. The number of candidate vectors that are potentially useful for neuroscience and medicine is vast, and no high-throughput means to test these vectors yet exists. Thus, there is a need for techniques to make multiple spatially and volumetrically accurate injections of viral vectors into NHP brain that can be identified unambiguously through postmortem histology. Described herein is such a method. Injection cannulas are constructed from coupled polytetrafluoroethylene and stainless-steel tubes. These cannulas are autoclavable, disposable, and have low minimal-loading volumes, making them ideal for the injection of expensive, highly concentrated viral vector solutions. An inert, red-dyed mineral oil fills the dead space and forms a visible meniscus with the vector solution, allowing instantaneous and accurate measurement of injection rates and volumes. The oil is loaded into the rear of the cannula, reducing the risk of co-injection with the vector. Cannulas can be loaded in 10 min, and injections can be made in 20 min. This procedure is well suited for injections into awake or anesthetized animals. When used to deliver high-quality viral vectors, this procedure can produce robust expression of optogenetic proteins, allowing optical control of neural activity and behavior in NHPs.

As currently implemented, optogenetics in non-human primates requires injection of viral vectors into the brain. An optimal injection method should be reliable and, for many applications, capable of targeting individual sites of arbitrary depth that are readily and unambiguously identified in postmortem histology. An injection method with these properties is presented.

Optogenetic techniques have revolutionized neuroscience research and are poised to do the same for neurological gene therapy. The clinical use of ...

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. Traditional in vitro electrophysiological techniques use electrical stimulation to evoke synaptic transmission. A major drawback of this method is its nonspecific nature; all axons in the region of the stimulating electrode will be activated, making it difficult to attribute an effect to a particular afferent connection. This issue can be overcome by replacing electrical stimulation with optogenetic-based stimulation. We describe a method for combining optogenetics with in vitro patch-clamp recordings. This is a powerful tool for the study of both basal synaptic transmission and synaptic plasticity of precise anatomically defined synaptic connections and is applicable to almost any pathway in the brain. Here, we describe the preparation and handling of a viral vector encoding channelrhodopsin protein for surgical injection into a pre-synaptic region of interest (medial prefrontal cortex) in the rodent brain and making of acute slices of downstream target regions (lateral entorhinal cortex). A detailed procedure for combining patch-clamp recordings with synaptic activation by light stimulation to study short- and long-term synaptic plasticity is also presented. We discuss examples of experiments that achieve pathway- and cell-specificity by combining optogenetics and Cre-dependent cell labeling. Finally, histological confirmation of the pre-synaptic region of interest is described along with biocytin labeling of the post-synaptic cell, to allow further identification of the precise location and cell type.

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Here we present a protocol describing viral transduction of discrete brain regions with optogenetic constructs to permit synapse-specific electrophysiological characterization in acute rodent brain slices.

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. ...

Photostimulation and Whole-Cell Patch Clamp Recordings of Neurons in Mouse Hippocampal Slices

Source: Richevaux, L., et al. <a href="https://app.jove.com/v/59534">In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices.</a> J. Vis. Exp. (2019)
The video demonstrates a protocol to study synaptic interactions and neuronal responses in a transfected mouse hippocampal brain slice. It involves illuminating long-range sensory neurons to trigger action potentials and recording the resulting excitatory post-synaptic potentials in the recorded neuron using the whole-cell patch-clamp technique.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Solutions for acute slice ...

JoVE Encyclopedia of Experiments

Neurophysiology

Electrophysiology Techniques

Simultaneous Optical and Electrophysiological Monitoring of Neuronal Cells in Brain Slices

Source:&#160; Murphy, C. A., et al. <a href="https://app.jove.com/v/61333">Optogenetic activation of afferent pathways in brain slices and modulation of responses by volatile anesthetics.</a> J. Vis. Exp. (2020)
This video outlines a method for simultaneously imaging and recording electrical activity from transgenic brain slices that contain neuron-specific fluorescent proteins. The procedure involves maintaining the brain slice in a perfusion chamber with aerated artificial cerebrospinal fluid, using a light source to excite fluorescent-labeled cells, and recording the electrical signals with a multichannel probe and patch clamp.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.&#160;&#160;&#160;&#160;&#160;
...

Optogenetic Stimulation and Electrophysiological Recording of Synaptic Events in Rat Brain Slices

Source: Kinnavane, L., et al. <a href="https://app.jove.com/v/63077">Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex.</a> J. Vis. Exp. (2022)
This video demonstrates the procedure for studying synaptic efficiency in the lateral entorhinal cortex (LEC) by optogenetically stimulating medial prefrontal cortex (mPFC) axons and recording the responses of pyramidal neurons in the LEC. This method combines patch-clamp recordings with synaptic activation by light stimulation to study synaptic plasticity.

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

Optogenetic Activation of Afferent Pathways in Brain Slices and Modulation of Responses by Volatile Anesthetics

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Chapters in this video

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

Optogenetic Manipulation of Neuronal Activity to Modulate Behavior in Freely Moving Mice

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices

Combined In Vivo Anatomical and Functional Tracing of Ventral Tegmental Area Glutamate Terminals in the Hippocampus

Injections of AAV Vectors for Optogenetics in Anesthetized and Awake Behaving Non-Human Primate Brain

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Photostimulation and Whole-Cell Patch Clamp Recordings of Neurons in Mouse Hippocampal Slices

Simultaneous Optical and Electrophysiological Monitoring of Neuronal Cells in Brain Slices

Optogenetic Stimulation and Electrophysiological Recording of Synaptic Events in Rat Brain Slices