This work was funded by the Whitehall Foundation and the NIH. We thank Clifford Dax (University of Oregon Technical Support Administration) for his help and expertise in designing a circuit for light delivery.

NOTE: The following protocol is in accordance with the National Institutes of Health guidelines as approved by the University of Oregon Animal Care and Use Committee.
1. Acute Surgery
<ol>
	<li>Anesthetize the animal with a ketamine-medetomidine cocktail, via intraperitoneal (i.p.) injection (Table 1). 
	NOTE: The mice used in these experiments are generated by crossing a cre-dependent ChR2-eYFP transgenic line10 to interneuron driver lines (Pvalb-iCre11, PV+; Sst-iCre12, SOM+; Cr-iCre12, CR+). Viral delivery of ChR2 or related opsins should work equally well, assuming similar expression levels are obtained.</li>
	<li>Before beginning surgery, ensure that the animal exhibits no response to a gentle toe pinch. Re-administer anesthetics throughout the experiment as necessary to maintain this depth of anesthesia. If using injectable anesthetics, optionally implant an i.p. catheter for maintenance injections.</li>
	<li>Keep the animal hydrated with saline or lactated Ringer&#8217;s solution throughout the experiment (approximately 3 ml/kg/hr), for example by using an appropriately diluted anesthetic cocktail for maintenance injections (Table 1).</li>
	<li>Place the animal in a stereotaxic or other head-holding apparatus. Ensure that the skull is well-secured. This is essential for maintaining stable single cell recordings.</li>
	<li>Apply opthalamic ointment to the eyes to prevent dryness. Maintain body temperature at 36.5-37 &#176;C.</li>
	<li>Perform a cisternal drain for additional recording stability. Using a scalpel blade, remove tissue from the posterior face of the skull to expose the cisterna magna. Make a small nick in the dura to drain the cerebrospinal fluid. Use only the very tip of the blade, and avoid contacting the cerebellum or brain stem.</li>
	<li>Using the scalpel, perform a small craniotomy (~2 mm2) over the cortical area of interest (for auditory cortex, roughly -2.3 mm posterior of bregma, 4.5 mm lateral of the midline).</li>
	<li>Remove the dura and cover the exposed cortex with a layer of warm agarose 0.5 - 1 mm thick (1.5% agarose in saline, 0.9% NaCl; apply at ~37 &#176;C). Keep the agarose moist throughout the experiment by periodically applying several drops of saline.</li>
</ol>
2. Recording Set-up
<ol>
	<li>Prepare a tungsten electrode (7-14 M&#937;, 127 &#956;m diameter, 12&#176; tapered tip, epoxy-coated). Superglue the electrode to a glass capillary tube and add heat shrink tubing for grip (Figure 1).</li>
	<li>Mount the electrode on a motorized (or hydraulic) micromanipulator, set to travel orthogonal to the cortical surface. Slide a wire ground under the skin, against the skull. Avoid contact with the muscles on the side of the head and the back of the neck as they can generate electromyographic artifacts.</li>
	<li>Amplify the electrical signal using an extracellular ampli&#64257;er suitable for single-unit recording, preferably equipped with an impedance check mode. Monitor ongoing spiking activity (band pass 300 - 5,000 Hz) with two oscilloscopes and a set of powered speakers. Here, the recorded data is digitized continuously at a sampling rate of 10 kHz; spikes are extracted offline.</li>
	<li>Advance the electrode through the agarose until the tip reaches the surface of the cortex. Observe this step through a microscope. &#8220;Zero&#8221; this position on the micromanipulator.</li>
	<li>To best approximate the depth of recording, visually confirm that the electrode tip exits the cortical surface at a depth of zero when withdrawing the electrode at the end of a penetration. 
	NOTE: A discrepancy between the manipulator reading and the observed electrode position indicates that it has drifted relative to the tissue. This can be caused by instability of the brain or animal, or manipulator drift.
	<ol>
		<li>As an additional check to ensure that this zero set-point corresponds to the surface of the cortex, monitor stimulus-evoked field potentials while advancing through the first several hundred micrometers of tissue. When monitoring field potentials, lower the lower band-pass filter cutoff of the extracellular amplifier to 10 Hz. Confirm that the polarity of the local field potential reverses around a depth of ~100 &#181;m, near the layer I/II border13. This is a reliable reference point for auditory cortex, but it may differ for other cortical areas.</li>
	</ol>
	</li>
	<li>Mount an optical fiber coupled to a blue light source (Figure 2) onto a manual micromanipulator. Using the microscope, position the tip of the fiber as close to the surface of the agarose as possible. Center the beam where the electrode will enter the tissue.</li>
	<li>Regulate the output of the light source with a control unit capable of delivering a TTL (transistor-transistor logic) pulse train of specified width and duration&#8211; e.g., an Arduino (Figure 3), a computer I/O card (as shown), or a commercially available pulse generator.
	<ol>
		<li>Monitor the signal on both oscilloscopes.</li>
	</ol>
	</li>
	<li>Measure total light power (mW) at the tip of the optical fiber using a power meter. Use this value to calculate irradiance (mW/mm2) by dividing by the cross-sectional area of the fiber core. Begin with an intensity value in the range of 10 - 15 mW/mm2 and adjust downward if necessary, until artifacts are eliminated.</li>
	<li>Check for light artifacts with the electrode in the agarose, poised to enter the tissue. Start the search pulse train (e.g., 30 msec light pulses, 500 msec interstimulus interval (ISI)).
	<ol>
		<li>Eliminate transient light artifacts (Figure 4) by repositioning the optical fiber relative to the electrode to change the angle of incident light. If artifacts persist, try decreasing the light power. In the rare case that artifacts cannot be completely eliminated (only minimized), extend the duration of the search pulse. This can aid in identifying true neuronal spikes.</li>
	</ol>
	</li>
</ol>
3. Straight PINP-in&#8217;
<ol>
	<li>Advance the electrode slowly through the brain at a rate of approximately 1 &#181;m/sec. Use one oscilloscope to monitor ongoing activity. On the other, trigger off the laser pulses.</li>
	<li>Listen for the faint &#8216;hash&#8217; of light-evoked spikes on the audio monitor, which indicates that the electrode is approaching a ChR2+ cell and can often be perceived well before the electrical signal is apparent on the oscilloscope. Slow the rate of advance. 
	NOTE: If the cell lies directly in the path of the electrode, the light-evoked activity will grow larger (and louder). As the electrode approaches a single interneuron, the hash will resolve into small but well-defined spikes of uniform size and shape.</li>
	<li>As soon as light-evoked spikes are large enough to trigger off of individually on the oscilloscope, begin doing so. Adjust the scaling on the horizontal axis to observe the exact shape of the spike waveform.</li>
	<li>Stop and wait for the tissue to settle. Resist the temptation to move the electrode closer to the cell. This step is critical for a stable recording. If after several minutes the signal-to-noise ratio has not improved &#8211; that is, the tissue has settled, but it hasn&#39;t brought the cell any closer to the tip of the electrode &#8211; advance 5 &#181;m further and wait again.
	<ol>
		<li>Repeat this process until either the peak or the trough of the action potential can be reliably captured with a voltage threshold set well above the noise floor (e.g., +/-300 &#181;V or greater). 
		NOTE: There&#8217;s little risk of false-positives or ambiguity when PINPing inhibitory interneurons. Most cells can easily follow a train of pulses with duration of 30 msec and inter-pulse interval 500 msec, or faster, with a reliable first spike latency of 2-5 msec (Figure 5). The majority of PV+ cells, in particular, can sustain firing for a full 1 sec (Figure 6). Because these are inhibitory neurons, disynaptic (indirect) activation isn&#8217;t a major concern.</li>
	</ol>
	</li>
	<li>While recording, monitor ongoing activity on the first oscilloscope. Keep a close eye on the size and shape of spikes using the second oscilloscope. Note the quality of their sound on the speaker.
	<ol>
		<li>Listen attentively for abrupt changes, which indicate that the cell is either drawing too close to the electrode (where it risks being impaled or damaged), or is drifting away. If the spikes become large and distorted, back out; if they grow smaller, advance the electrode. Move slowly in 2 &#181;m steps.</li>
		<li>Confirm the quality of the recording post-hoc by superimposing all spike waveforms, aligned to peak or trough. Vary the voltage threshold. Across a wide range of threshold values, there should only ever be one consistent spike shape of uniform height (Figure 5a).</li>
	</ol>
	</li>
	<li>If at any point the signal becomes contaminated with spikes from a neighboring neuron, move on. Advance slowly, on the off-chance that the electrode will pass out of range of the neighboring cell, but remain in range of the target neuron. (This is usually unsuccessful.)</li>
	<li>Move the electrode through the entire depth of cortex (900+ &#181;m) in each penetration. If no light-responsive neurons are encountered after many penetrations or, conversely, recordings are routinely contaminated with spikes from neighboring ChR2- cells, try a new electrode. 
	NOTE: Even within a single batch, the impedance and tip geometry of individual electrodes can vary considerably. Both factors contribute to the effective &#8220;listening radius&#8221; of the electrode, which must be large enough to detect light-evoked spikes while searching, yet restricted enough to enable recording a single interneuron in isolation.</li>
	<li>Test the impedance of electrodes as desired. To avoid damaging the tissue, do this with the tip of the electrode in the upper part of the agarose layer, well outside the brain. Within the range of 7-14 M&#937;, the exact impedance is not a sure predictor of performance, or yield, for this application. That said, it&#8217;s a reasonable proxy for listening radius: lower-impedance electrodes pick up more units; higher-impedance, fewer.</li>
	<li>At the end of the experiment, euthanize the animal by institutionally-approved means, such as anesthetic overdose, or cervical dislocation under a deep plane of anesthesia.</li>
</ol>

<ol>
	<li>Lima, S. Q., Hromadka, T., Znamenskiy, P., Zador, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PINP:+a+new+method+of+tagging+neuronal+populations+for+identification+during+in+vivo+electrophysiological+recording.">PINP: a new method of tagging neuronal populations for identification during in vivo electrophysiological recording.</a> PLoS One. 4, (2009).</li><li>Moore, A. K., Wehr, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Parvalbumin-expressing+inhibitory+interneurons+in+auditory+cortex+are+well-tuned+for+frequency.">Parvalbumin-expressing inhibitory interneurons in auditory cortex are well-tuned for frequency.</a> J Neurosci. 33, 13713-13723 (2013).</li><li>Merchant, H., de Lafuente, V., Pena-Ortega, F., Larriva-Sahd, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+impact+of+interneuronal+inhibition+in+the+cerebral+cortex+of+behaving+animals.">Functional impact of interneuronal inhibition in the cerebral cortex of behaving animals.</a> Prog Neurobiol. 99, 163-178 (2012).</li><li>Markram, H., 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=Interneurons+of+the+neocortical+inhibitory+system.">Interneurons of the neocortical inhibitory system.</a> Nat Rev Neurosci. 5, 793-807 (2004).</li><li>Atallah, B. V., Bruns, W., Carandini, M., Scanziani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Parvalbumin-expressing+interneurons+linearly+transform+cortical+responses+to+visual+stimuli.">Parvalbumin-expressing interneurons linearly transform cortical responses to visual stimuli.</a> Neuron. 73, 159-170 (2012).</li><li>Wilson, N. R., Runyan, C. A., Wang, F. L., Sur, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Division+and+subtraction+by+distinct+cortical+inhibitory+networks+in+vivo.">Division and subtraction by distinct cortical inhibitory networks in vivo.</a> Nature. 488, 343-348 (2012).</li><li>Letzkus, J. J., 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=A+disinhibitory+microcircuit+for+associative+fear+learning+in+the+auditory+cortex.">A disinhibitory microcircuit for associative fear learning in the auditory cortex.</a> Nature. 480, 331-335 (2011).</li><li>Pi, H. J., 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=Cortical+interneurons+that+specialize+in+disinhibitory+control.">Cortical interneurons that specialize in disinhibitory control.</a> Nature. 503, 521-524 (2013).</li><li>Adesnik, H., Bruns, W., Taniguchi, H., Huang, Z. J., Scanziani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+neural+circuit+for+spatial+summation+in+visual+cortex.">A neural circuit for spatial summation in visual cortex.</a> Nature. 490, 226-231 (2012).</li><li>Madisen, L., 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=A+toolbox+of+Cre-dependent+optogenetic+transgenic+mice+for+light-induced+activation+and+silencing.">A toolbox of Cre-dependent optogenetic transgenic mice for light-induced activation and silencing.</a> Nat Neurosci. 15, 793-802 (2012).</li><li>Hippenmeyer, S., 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=A+developmental+switch+in+the+response+of+DRG+neurons+to+ETS+transcription+factor+signaling.">A developmental switch in the response of DRG neurons to ETS transcription factor signaling.</a> PLoS Biol. 3, 159 (2005).</li><li>Taniguchi, H., 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=A+resource+of+Cre+driver+lines+for+genetic+targeting+of+GABAergic+neurons+in+cerebral+cortex.">A resource of Cre driver lines for genetic targeting of GABAergic neurons in cerebral cortex.</a> Neuron. 71, 995-1013 (2011).</li><li>Christianson, G. B., Sahani, M., Linden, J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Depth-dependent+temporal+response+properties+in+core+auditory+cortex.">Depth-dependent temporal response properties in core auditory cortex.</a> J Neurosci. 31, 12837-12848 (2011).</li><li>Povysheva, N. V., Zaitsev, A. V., Gonzalez-Burgos, G., Lewis, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+heterogeneity+of+fast-spiking+interneurons:+chandelier+versus+basket+cells.">Electrophysiological heterogeneity of fast-spiking interneurons: chandelier versus basket cells.</a> PLoS One. 8, 70553 (2013).</li></ol>

The authors have no competing financial interests.

Although PINP is conceptually straightforward, it can be challenging in practice. A major determinant of success is the choice of electrode. The electrical listening radius is the critical parameter. It must be sufficiently large to detect light-evoked spikes when the tip is still some distance away from a ChR2+ cell, so that one can adjust the rate of advance accordingly. At the same time, it must be restricted enough to enable good single-unit isolation. That is, the electrode must not also pick up spikes from neighboring ChR2- units. Striking the right balance in terms of listening radius will be a challenge for any targeted cell type, but it is especially true for inhibitory interneurons, which are sparse, small, and often found in close proximity to pyramidal cells, which that have comparatively large extracellular spikes. Even using the strategies suggested here, the expected yield is not high. Our typical yield was 0-2 PV+ neurons/animal, given our criteria of excellent single-unit isolation and recordings lasting over half an hour.
When PINPing inhibitory neurons, disynaptic (indirect) activation isn&#8217;t a major concern, because inhibitory neurons are relatively unlikely to indirectly activate postsynaptic cells. For PINPing excitatory neurons, disynaptic activation is common; many neurons that fire in response to light do not express ChR2, but rather receive powerful synaptic excitation from those that do. Strategies for discriminating directly- from indirectly-activated neurons are discussed in detail in Lima et al.1.
Electrode Considerations
We systematically explored a variety of electrodes and recording techniques: glass patch pipettes with tips of different sizes, tungsten microelectrodes varying in impedance and tip geometry, and multi-channel arrays (e.g., tetrodes). Angular, high-impedence tungsten microelectrodes (as suggested here) gave, by far, the best yield. A discussion of the issues encountered with each of these techniques follows.
Using low-impedance tetrode arrays and multi-channel probes, we routinely detected light-evoked spikes; however, they were rarely large enough to sort reliably. In other words, the signal-to-noise ratio (SNR) was unsatisfactory. We did occasionally get excellent recordings with tetrodes, but the asymptotic yield was higher with single tungsten microelectrodes, and the quality of recordings was much better. Tetrode tips are comparatively blunt as well, which limited the total number of penetrations possible per animal. The issues we encountered with multi-channel probes were probably exacerbated by the use of broad surface illumination. Unlike targeted, low-power stimulation from an implanted fiber, surface illumination will cause synchronous firing in ChR2+ neurons beyond the recording site, leading to superimposed spike waveforms that are more difficult to sort.
Glass patch electrodes, like those used for standard loose cell-attached recordings, were also not well-suited to the task, but for different reasons. Low resistance (&#60;5 M&#937;) patch electrodes&#160;are strongly biased toward pyramidal cells,&#160;which&#160;makes it difficult to target inhibitory interneurons. While this can be&#160;corrected to some extent by using higher resistance (smaller tip) electrodes, the&#160;greater&#160;problem with patch electrodes is their extremely restricted listening radius. Although cell-attached patch recordings provide a very high SNR, light-responsive cells&#160;cannot&#160;be detected&#160;outside of&#160;cell-attached mode. The strategy with patch electrodes is, inevitably, to randomly and sequentially obtain cell-attached recordings,&#160;very few of which are ChR2-expressing&#160;inhibitory interneurons.
With high impedance tungsten electrodes one can both detect light-evoked spikes from a distance, and achieve good single-unit isolation. The geometry of the tip is probably the most important factor. We tried tungsten electrodes from two manufacturers, with impedances ranging from 2 - 14 M&#937;, and tip angles from 5-12&#176;. We found that 12&#176; tip angles were superior, but that the precise impedance was less important, within the range of 7 - 14 M&#937;. We found that glass-coated electrodes were more susceptible to light artifacts compared to epoxy-coated electrodes, so we recommend the latter. We did not attempt to make our own custom tungsten electrodes. As with any type of electrode, common modes of failure are to lose isolation or injure a neuron. Different tip parameters do not appear to have much influence on this. Rather, waiting for the tissue to settle is the more important factor in maintaining high-quality recordings.
Light Delivery
Because light is used only as a means of identifying ChR2-positive cells &#8211; rather than regulating their output in a specific manner &#8211; the range of usable intensities for this experiment is wide. A lower limit is set by the fact that the intensity must be sufficient to activate ChR2-positive cells at the maximum depth of your cells of interest. We found that tip irradiances &#62;5 mW/mm2 (measured at 470 nm) were sufficient to elicit robust responses from PV+ cells in deep Layer 6. Irradiance at specific depths within the brain can be estimated using a calculator based on direct measurements in mammalian brain tissue (e.g., <a href="http://www.stanford.edu/group/dlab/cgi-bin/graph/chart.php" target="_blank">http://www.stanford.edu/group/dlab/cgi-bin/graph/chart.php</a>), but we strongly recommend determining the minimum for your own experiments empirically, using a low impedance microelectrode (1 - 3 M&#937;, for multiunit activity). An upper limit is set by the fact that high intensities can cause light artifacts, which can complicate the detection of evoked spikes. In transgenic animals, high intensities can also cause electromyographic artifacts by activating ChR2 in exposed PV+ muscle on the neck and head.
In building our own light delivery system, we found that a coupling an LED to an optical fiber is ultimately more convenient than placing the LED directly over the brain. The latter is a very effective way to deliver light, but has some practical disadvantages. The current flowing through the device produces electrical artifacts that must be shielded, and the LED generates substantial heat that must be dissipated with a large heat sink. Moreover, light artifacts (both photovoltaic and electromyographic) are much more difficult to control with the broad illumination pattern of LEDs. Fiber coupling solves all of these issues.

Characterizing the myriad cell types that comprise the mammalian brain has been a central, but long-elusive goal of neurophysiology. For instance, the properties and function of different inhibitory cell types in the cerebral cortex are topics of great interest but are still relatively unknown. This is in part because conventional blind in vivo recording techniques are limited in their ability to distinguish between different cell types. Extracellular spike width can be used to separate putative parvalbumin-positive inhibitory neurons from excitatory pyramidal cells, but this method is subject to both type I and type II errors2,3. Alternatively, recorded neurons can be filled, recovered, and stained to later confirm their morphological and molecular identity, but this is a pain-staking and time-consuming process. Recently, genetically identified populations of inhibitory interneurons have become accessible by means of calcium imaging or visually guided patch recordings. In these approaches, viral or transgenic expression of a calcium reporter (such as GCaMP) or fluorescent protein (such as GFP) allows identification and characterization of cell types defined by promoter expression. These approaches use 2-photon microscopy, which requires expensive equipment, and are also limited to superficial cortical layers due to the light scattering properties of brain tissue. 
Recently, Lima and colleagues1 developed a novel application of optogenetics to target electrophysiological recordings to genetically identified neuronal types in vivo, termed &#8220;PINP&#8221; &#8211; or Photostimulation-assisted Identification of Neuronal Populations. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. Unlike many other optogenetic applications, the goal is not to manipulate circuit function but simply to identify neurons belonging to a genetically-defined class, which can then be characterized during normal brain function. The technique can be implemented with standard extracellular recording equipment and can therefore serve as an accessible and inexpensive alternative to calcium imaging or visually-guided patching. Here we describe an approach to PINPing specific cell types in the anesthetized auditory cortex, with the expectation that the more general points can be usefully applied in other preparations and brain regions.
In cortex, PINP holds particular promise for investigating the in vivo response properties of inhibitory interneurons. GABAergic interneurons comprise a small, heterogeneous subset of cortical neurons4. Different subtypes, marked by the expression of particular molecular markers, have recently been shown to perform different computational roles in cortical circuits5-9. As genetic tools improve it may eventually be possible to distinguish morphologically- and physiologically-separable types that fall within these broad classes. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. This strategy was developed specifically for targeting parvalbumin-positive (PV+) cells, but we have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons. Although PINPing is conceptually straightforward, it can be surprisingly unyielding in practice. We learned a number of tips and tricks through trial-and-error that may be useful to others attempting the method.

<table><tbody><tr><td>ChR2-EYFP Line</td><td>Jackson Colonies</td><td>12569</td><td></td></tr><tr><td>Pvalb-iCre (PV) Line</td><td>Jackson Colonies</td><td>8069</td><td></td></tr><tr><td>Sst-iCre (SOM) Line</td><td>Jackson Colonies</td><td>13044</td><td></td></tr><tr><td>Cr-iCre (CR) Line</td><td>Jackson Colonies</td><td>10774</td><td></td></tr><tr><td>Agarose</td><td>Sigma-Aldrich</td><td>A9793</td><td>Type III-A, High EEO</td></tr><tr><td>Micro Point (dural hook)</td><td>FST</td><td>10066-15</td><td></td></tr><tr><td>Surgical Scissors</td><td>FST</td><td>14084-09</td><td></td></tr><tr><td>Scalpel</td><td>FST</td><td>10003-12 (handle), 10011-00 (blades)</td><td></td></tr><tr><td>Puralube Ophthalmic Ointment</td><td>Foster &amp;amp; Smith</td><td>9N-76855</td><td></td></tr><tr><td>Homeothermic Blanket</td><td>Harvard Apparatus</td><td>&lt;a name=&quot;RANGE!C11&quot;&gt;507220F&lt;/a&gt;</td><td></td></tr><tr><td>Tungsten Microelectrodes</td><td>A-M Systems</td><td>577200</td><td>12 M&amp;Omega; AC resistance, 127 &amp;mu;m diameter, 12&amp;deg; tapered tip, epoxy-coated</td></tr><tr><td>Capillary Glass Tubing</td><td>Warner Instruments</td><td>G150TF-3</td><td></td></tr><tr><td>Heat Shrink Tubing</td><td>DigiKey</td><td>A332B-4-ND</td><td></td></tr><tr><td>Zapit Accelerator</td><td>DVA</td><td>SKU ZA/ZAA</td><td>Use with standard Super Glue.&amp;nbsp;</td></tr><tr><td>Microelectrode AC Amplifier 1800</td><td>AM Systems</td><td>700000</td><td></td></tr><tr><td>MP-285 Motorized Micromanipulator</td><td>Sutter</td><td>MP-285</td><td></td></tr><tr><td>4-channel Digital Oscilloscopes</td><td>Tektronix</td><td>TDS2000C</td><td></td></tr><tr><td>Powered Speakers</td><td>Harman</td><td>Model JBL&amp;nbsp;Duet</td><td></td></tr><tr><td>Manual Manipulator</td><td>Scientifica</td><td>LBM-7</td><td></td></tr><tr><td>800 &amp;micro;m Fiber Optic Patch Cable</td><td>ThorLabs</td><td>FC/PC BFL37-800</td><td></td></tr><tr><td>Power Meter</td><td>ThorLabs</td><td>PM100D (Power Meter), S121C (Standard Power Sensor)</td><td></td></tr><tr><td>475 nm Cree XLamp XP-E</td><td>DigiKey</td><td>XPEBLU-L1-R250-00Y01DKR-ND</td><td>LED power and efficiency are continually increasing, so we recommend checking for the latest products (www.cree.com).</td></tr><tr><td>Arduino UNO</td><td>DigiKey</td><td>1050-1024-ND</td><td></td></tr></tbody></table>

a guide to in vivo single-unit recording from optogenetically identified cortical inhibitory interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

We here share our strategy for obtaining single-unit recordings from genetically-classified inhibitory interneurons in the anesthetized mouse cortex, using an optogenetic method developed by Lima et al.1. Table 1 details the suggested anesthetic cocktail, Ketamine-Medetomidine-Acepromazine (&#8220;KMA&#8221;). Figure 1 depicts a tungsten microelectrode, prepared for recording. Figure 2 contains a circuit diagram for a simple LED control unit. Figure 3 contains the configuration and code for gating light output with an Arduino microcontroller. Figure 4 shows an example of the light-induced electrical artifacts discussed in the Protocol. Artifacts are occasionally encountered (left panel), but easily corrected (right panel). Figure 5 shows example recordings from three types of optically-identified interneurons (PV+, CR+, and SOM+). The left panel shows raw voltage traces and aligned waveforms, representative of the kind of signal-to-noise ratio one can expect to obtain with this strategy. Spikes are reliably captured with a fixed voltage threshold of several hundred microvolts. A raster plot (right panel) shows the short latency and reliability of light-evoked responses, which allow for unambiguous identification of ChR2-positive interneurons. Figure 6 shows voltage traces from three PV+ interneurons, responding to a 1 sec light pulse. The majority of PV+ cells can sustain high-frequency firing for hundreds of milliseconds, making them particularly easy to identify.
<img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/51757/51757fig1highres.jpg" width="600" /> 
Figure 1. Preparing an electrode.&#160;(A) Diagram of a prepared electrode. The electrode is affixed to a glass capillary tube using two small drops of Super Glue and a minimal amount of accelerator (such as Zap-It). Heat shrink tubing is added for grip, using very low heat. Electrodes typically ship with a connector pin pre-attached. (B) Photograph of a prepared tungsten microelectrode mounted in an electrode holder.
<img alt="Figure 2" fo:content-width="6in" src="/files/ftp_upload/51757/51757fig2highres.jpg" width="600" /> 
Figure 2. Circuit diagram for an LED control unit. A simple and inexpensive circuit for light delivery. A super-bright LED (475 nm) is attached to a heat sink and coupled to the other end of a fiber optic cable (800 &#181;m diameter). The output of the LED is gated with a TTL pulse. The control unit incorporates a voltage-mode power supply for adjustable light intensity. 2K POT: potentiometer to control light intensity. P/S: DC power supply (e.g., laptop charger). TIP 122: Darlington transistor. 4N35: Optical isolator. The parts listed here cost &#60;$10.
<img alt="Figure 3" fo:content-width="6in" src="/files/ftp_upload/51757/51757fig3highres.jpg" width="600" /> 
Figure 3. Arduino code and configuration. A TTL pulse train for gating light output can be generated using an Arduino microcontroller. The program specifies a pulse duration and ISI for a looping pulse train. Instructions for loading the program onto the board can be found on Arduino&#8217;s &#8220;Getting Started&#8221; page (<a href="http://arduino.cc/en/Guide/HomePage" target="_blank">http://arduino.cc/en/Guide/HomePage</a>). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51757/51757fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" fo:content-width="6in" src="/files/ftp_upload/51757/51757fig4highres.jpg" width="600" /> 
Figure 4. Light artifact. Metal electrodes are susceptible to light artifacts, present at the onset and the offset of pulses (left panel; filtered signal, 300 - 5,000 Hz). They can usually be eliminated completely (right panel) by repositioning the optical fiber relative to the electrode to change the angle of incident light, and/or decreasing the light power.
<img alt="Figure 5" fo:content-width="6in" src="/files/ftp_upload/51757/51757fig5highres.jpg" width="600" /> 
Figure 5. Example cells. (A) Typical example recordings from optically-identified interneurons, PV+ (purple), CR+ (green), SOM+ (red). Cells respond to the search pulse train (pulse duration 30 ms, ISI 500 ms) with a reliable burst of spikes. The panel to the right shows 100 trough-aligned spikes and the average interpolated waveform (10x oversampled from a sampling rate of 10 kHz). (B) Raster plots for the same files show the short latency (~2 - 5 msec) and consistent timing of light-evoked spikes. (C) Mean and standard deviation of first-spike latencies (5 - 10 pulses) for a sample of 44 PV+ interneurons (average mean and standard deviation: 3.4 +/- 2.6 msec). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51757/51757fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" fo:content-width="6in" src="/files/ftp_upload/51757/51757fig6highres.jpg" width="600" /> 
Figure 6. Examples of sustained responses. One can be especially confident in identifying PV+ interneurons as the majority of them can sustain high-frequency firing for hundreds of milliseconds &#8211; reminiscent of their responses to current injection in vitro14. Three ChR2/PV cells responding to 1 sec light pulses: two that fire regularly throughout the entire duration of the pulse, and one unusual example of a cell that does not (&#60;10% of encountered cells).
<table border="0" cellpadding="0" cellspacing="0" fo:keep-together.within-page="always">
	<tbody>
		<tr>
			<td>KMA (20.0 ml)</td>
		</tr>
		<tr>
			<td>2 ml of 100 mg/ml Ketamine (10 mg/ml, once diluted)&#160;</td>
		</tr>
		<tr>
			<td>0.4 ml of 1 mg/ml Dexdomitor (medetomidine; 0.02 mg/ml, once diluted)&#160;</td>
		</tr>
		<tr>
			<td>0.05 ml of 100 mg/ml Acepromazine (0.25 mg/ml, once diluted)&#160;</td>
		</tr>
		<tr>
			<td>17.55 ml of 0.9% saline</td>
		</tr>
		<tr>
			<td>Dosage, adult mouse (15 - 40 g)</td>
		</tr>
		<tr>
			<td>Knock-down: 0.012 ml/g * body mass (Ketamine = 120 mg/kg)</td>
		</tr>
		<tr>
			<td>Maintenance: 0.0025 ml/g * body mass (Ketamine = 25 mg/kg) &#160;</td>
		</tr>
	</tbody>
</table>

Table 1. Ketamine-Medetomidine-Acepromazine (&#8220;KMA&#8221;). Knock-down and maintenance dose for an adult mouse, diluted to maintain hydration. The cocktail is re-administered every 40 - 90 min, as necessary.

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A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

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19.1K Views.  University of Oregon. Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching. 

Video: A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Single-unit In vivo Recordings from the Optic Chiasm of Rat

Information about the visual world is transmitted to the brain in sequences of action potentials in retinal ganglion cell axons that make up the optic nerve. In vivo recordings of ganglion cell spike trains in several animal models have revealed much of what is known about how the early visual system processes and encodes visual information. However, such recordings have been rare in one of the most common animal models, the rat, possibly owing to difficulty in detecting spikes fired by small diameter axons. The many retinal disease models involving rats motivate a need for characterizing the functional properties of ganglion cells without disturbing the eye, as with intraocular or in vitro recordings. Here, we demonstrate a method for recording ganglion cell spike trains from the optic chiasm of the anesthetized rat. We first show how to fabricate tungsten-in-glass electrodes that can pick up electrical activity from single ganglion cell axons in rat. The electrodes outperform all commercial ones that we have tried. We then illustrate our custom-designed stereotaxic system for in vivo visual neurophysiology experiments and our procedures for animal preparation and reliable and stable electrode placement in the optic chiasm.

Single-unit In vivo Recordings from the Optic Chiasm of Rat

Retinal ganglion cells transmit visual information from the eye to the brain with sequences of action potentials. Here, we demonstrate how to record the action potentials of single ganglion cells in vivo from anesthetized rats.

Information about the visual world is transmitted to the brain in sequences of action potentials in retinal ganglion cell axons that make up the optic ...

Mapping Inhibitory Neuronal Circuits by Laser Scanning Photostimulation

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into diverse subtypes based on their immunochemical, morphological, and physiological properties1-4. Although previous research has revealed much about intrinsic properties of individual types of inhibitory neurons, knowledge about their local circuit connections is still relatively limited3,5,6. Given that each individual neuron's function is shaped by its excitatory and inhibitory synaptic input within cortical circuits, we have been using laser scanning photostimulation (LSPS) to map local circuit connections to specific inhibitory cell types. Compared to conventional electrical stimulation or glutamate puff stimulation, LSPS has unique advantages allowing for extensive mapping and quantitative analysis of local functional inputs to individually recorded neurons3,7-9. Laser photostimulation via glutamate uncaging selectively activates neurons perisomatically, without activating axons of passage or distal dendrites, which ensures a sub-laminar mapping resolution. The sensitivity and efficiency of LSPS for mapping inputs from many stimulation sites over a large region are well suited for cortical circuit analysis.
Here we introduce the technique of LSPS combined with whole-cell patch clamping for local inhibitory circuit mapping. Targeted recordings of specific inhibitory cell types are facilitated by use of transgenic mice expressing green fluorescent proteins (GFP) in limited inhibitory neuron populations in the cortex3,10, which enables consistent sampling of the targeted cell types and unambiguous identification of the cell types recorded. As for LSPS mapping, we outline the system instrumentation, describe the experimental procedure and data acquisition, and present examples of circuit mapping in mouse primary somatosensory cortex. As illustrated in our experiments, caged glutamate is activated in a spatially restricted region of the brain slice by UV laser photolysis; simultaneous voltage-clamp recordings allow detection of photostimulation-evoked synaptic responses. Maps of either excitatory or inhibitory synaptic input to the targeted neuron are generated by scanning the laser beam to stimulate hundreds of potential presynaptic sites. Thus, LSPS enables the construction of detailed maps of synaptic inputs impinging onto specific types of inhibitory neurons through repeated experiments. Taken together, the photostimulation-based technique offers neuroscientists a powerful tool for determining the functional organization of local cortical circuits.

This paper introduces an approach of combining laser scanning photostimulation with whole cell recordings in transgenic mice expressing GFP in limited inhibitory neuron populations. The technique allows for extensive mapping and quantitative analysis of local synaptic circuits of specific inhibitory cortical neurons.

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into ...

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

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 ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

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 ...

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.

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 ...

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. ...

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

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