This work was supported by NIH/NIDCD grant R03 DC 011361, SVM CORP grant LAV3202, NSF-LA EPSCoR grant PFUND86, and a publication grant from Leica Microsystems (CCL) and NIH/NIDCD grant R01 DC 008794 (SMS).

1. Surgery and Injections
<ol>
	<li>Anesthetize mice using an intraperitoneal injection (22-25 G needle recommended) of ketamine (100-120 mg/kg) and xylazine (7.5-16 mg/kg).
	<ol>
		<li>Assess sedation using strong toe pinches until no responses are elicited. For all animal procedures described, modify to follow local IACUC guidelines as necessary.</li>
	</ol>
	</li>
	<li>Place sedated mice in a stereotaxic device by securing the head and ears in their respective holders.
	<ol>
		<li>Lubricate the eyes prior to placing the animal in the stereotaxic device with a sterile ophthalmic ointment.</li>
	</ol>
	</li>
	<li>Shave the scalp with a razor and sterilize the area with Betadine swabs.</li>
	<li>Using a scalpel, cut a midline incision and reflect the skin and underlying muscles above the area to be injected.</li>
	<li>Rotate a scalpel blade as a trephine to perform a craniotomy above the region to be exposed. Alternatively, use a small drill bit attached to a Dremel to perform the craniotomy.
	<ol>
		<li>To perform the craniotomy, place scalpel blade or drill bit on surface of skull and slowly rotate until enough bone has been excavate to reveal the pia mater underneath.</li>
		<li>Once the initial excavation is made, enlarge the craniotomy using fine forceps to carefully remove bone from the edges.</li>
	</ol>
	</li>
	<li>Fill a 1 &#956;l syringe with the viral solution to be injected (UNC Vector Core).
	<ol>
		<li>To enhance transfection efficacy, prepare viral solution with 4 &#956;g/ml polybrene.</li>
	</ol>
	</li>
	<li>Lower the needle into the area to be injected and pressure infuse 200 nl of solution containing virus with a syringe pump.
	<ol>
		<li>Let syringe sit for 10 min before withdrawing the needle.</li>
	</ol>
	</li>
	<li>Suture the skin with nonabsorbable monofilament Nylon and cover the incision with antibiotic ointment.
	<ol>
		<li>Sutures should be removed after 7-10 days.</li>
	</ol>
	</li>
	<li>Inject buprenorphine (0.05-0.1 mg/kg) for post-operative analgesia.</li>
	<li>Allow animals to post-operatively recover for 14-21 days to allow for adequate expression of channel rhodopsin.</li>
</ol>
2. Slice Preparation
<ol>
	<li>Following the recovery period, deeply anesthetize the animal with isoflurane anesthesia, as verified by absence of withdrawal reflex when the hind foot and/or toes are firmly pinched.</li>
	<li>Dissect the brain by decapitation followed by midline and frontal cuts to the skull.</li>
	<li>Peel apart the skull from the midline to expose the brain.</li>
	<li>Carefully remove the brain using a fine scoop spatula to separate the cranial nerves from the base of the brain.</li>
	<li>Quickly place dissected brain in small 100 ml beaker containing chilled, ice-cold artificial cerebrospinal fluid (ACSF; in mM: 125 NaCl, 3 KCl, 1.25 NaH2PO4, 1 MgCl2, 2 CaCl2, 25 NaHCO3, 25 glucose).</li>
	<li>Remove brain from ACSF and place on a microscope slide covered with filter paper prewetted with ACSF.</li>
	<li>Make a blocking cut of the brain in the desired orientation using a clean razor blade.</li>
	<li>Affix the blocked surface of the brain to cutting stage using instant adhesive glue.</li>
	<li>Transfer stage to a vibratome cutting chamber (kept in freezer until ready for cutting).</li>
	<li>Pour ice cold ACSF into cutting chamber.</li>
	<li>Slice brain sections at 400-500 &#181;m using a vibrating tissue slicer.</li>
	<li>Transfer brain slices to a holding chamber containing oxygenated ACSF, and incubate at 30 &#176;C for at least 1 hr before experiment.</li>
</ol>
3. Physiological Recording
<ol>
	<li>After the incubation period, transfer a slice to a recording chamber on a modified microscope stage.</li>
	<li>Secure the slice in the bath during the experiment using a few threads of Nylon filaments attached to a platinum wire slice holder.</li>
	<li>Perfuse the slice with ACSF constantly.</li>
	<li>Pull recording pipettes to a tip resistance of 3-6 M&#937;&#160;when filled with intracellular solution containing the following (in mM): 127 K-gluconate, 3 KCl, 1 MgCl2, 0.07 CaCl2, 10 HEPES, 2 Na2-ATP, 0.3 Na-GTP, 0.1 EGTA. The intracellular solution should have pH 7.3, adjusted with KOH or gluconic acid and the osmolality of 280-290 mOsm.</li>
	<li>Using a standard visualized whole-cell recording setup, place recording pipette near neuron to be recorded while applying positive pressure22,23.</li>
	<li>Release positive pressure while applying suction to form Gigaohm seal.</li>
	<li>Apply brief suction to break membrane and form whole-cell configuration recording.</li>
	<li>Record the responses of neurons in either voltage or current clamp.</li>
</ol>
4. Laser-scanning Photostimulation
<ol>
	<li>Start data acquisition software and hardware, described in previous reports23-29. 
	Note: We have utilized both Tidelwave25 and, more recently, Ephus26 acquisition programs, written in Matlab and developed by the laboratory of Karel Svboda at Janelia Farms30. For a full description of laser-scanning photostimulation in Ephus, see Suter et al.26&#160;and the step-by-step guide30. 
	Note: The setup described in this article is derived from that described by Shepherd (2012)23. The laser beam is guided via an open beam&#160;system of mirrors to the top port of the microscope through the objective lens onto the slice (Figure&#160;2).</li>
	<li>Divert a small portion of the laser onto a photodiode with a microscope coverslip placed in the laser path and use the current output from this photodiode to monitor the laser intensity during experiments.
	<ol>
		<li>Calibrate the photodiode current output recorded in the software to the measured laser power at the back focal plane of the objective using a power meter during equipment setup23.</li>
	</ol>
	</li>
	<li>Control the laser power by rotating a variable neutral density wheel. The laser power used varied between 25-70 mW (at the back focal plane) in our experiments.</li>
	<li>Set the length and timing of the laser stimulating by modifying the pulse parameters for the laser and shutter. Laser stimulation is typically 2 msec long.</li>
	<li>Capture image of the region to be stimulated using the &#39;Grab Video&#39; function.</li>
	<li>Select the stimulation grid according to desired dimensions. We typically use a 16 x 16 stimulation array with 80 &#956;m spacing between adjacent rows and columns.</li>
	<li>Overlay the mapping grid with the acquired image and adjust the x, y, and degree offset to the desired location and orientation.</li>
	<li>Activate the &#39;Map&#39; function to start stimulation and acquisition of data.</li>
	<li>Following acquisition of photostimulation evoked responses, carry out analysis of mapped data using either the offline map analysis program in Tidelwave25 or the &#39;MapAnalysis&#39; program in Ephus26.</li>
</ol>
5. Fixed Slice Imaging
<ol>
	<li>After recordings, transfer brain slice to 4% paraformaldehyde (PFA) in 0.01 M PBS (pH7.4) overnight at 4 &#186;C.</li>
	<li>Transfer the slice in 30% sucrose in 4% PFA/0.01 M PBS overnight at 4 &#186;C.</li>
	<li>Cut 50 &#181;m sections using a cryostat and collect in 0.01 M PBS.</li>
	<li>Rinse sections in 0.01 M PBS twice at room temperature.</li>
	<li>Mount sections and coverslip in an appropriate antifade medium.</li>
	<li>Transfer sections to the confocal microscope.</li>
	<li>Focus into the ROI using white light.</li>
	<li>Choose the wavelength, imaging size, and repetition for image acquisition. For YFP, we use 514 nm (Ex)/527 nm (Em).</li>
</ol>

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J., Mogri, M., Deisseroth, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optogenetics+in+neural+systems.">Optogenetics in neural systems.</a> Neuron. 71, 9-34 (2011).</li><li>Nagel, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Channelrhodopsin-2,+a+directly+light-gated+cation-selective+membrane+channel.">Channelrhodopsin-2, a directly light-gated cation-selective membrane channel.</a> Proc. Natl. Acad. Sci. U.S.A. 100, 13940-13945 (2003).</li><li>Zemelman, B. V., Lee, G. 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The authors declare they have no competing interests.
The author (CCL) received a publication grant from Leica Microsystems, which produces an instrument used in this filming of this protocol.

The described protocol details an approach for the in vitro assessment of functional topography and physiology of neuronal circuits, which is not limited to the investigation of forebrain circuits, i.e. those of the thalamus and cortex. Several viral constructs for optogenetic transfection have been made available to the scientific community and are readily obtainable from the UNC Vector Core. For a description of viral constructs readily available, see the UNC Vector Core31, the optogenetics resource page at the Deisseroth Lab at Stanford University32 and the Boyden synthetic neurobiology lab at MIT33. Variants of these opsins have been developed by these labs and enable wide-ranging modes for controlling neuronal excitability1,2. Obtaining these vectors from the UNC Vector Core requires a Material Transfer Agreement from the originating labs. In this protocol, we note that the addition of polybrene enhances transfection efficacy. In addition, another variable influencing expression is the post-surgical recovery period, which we note that a period of at least 14 days is sufficient to obtain robust expression in the forebrain pathways of mice.
For the targeted transfection of specific neuronal groups, several Cre-transgenic mouse lines have been developed, which can be used in combination with &#39;floxed&#39; reversed open reading frame viral constructs that have been developed11. Several strains pertinent to neuroscience research are available through the Mutant Mouse Regional Resource Centers34. Atlases of Cre-recombinase expression patterns in these transgenic lines are offered through the GENSAT database35. In addition, Jackson Labs offers several Cre-transgenic strains, as well as others that constitutively express opsins in specific neuronal groups. A critical step to utilizing these mouse lines is to factor in the time necessary for obtaining and establishing a breeding colony, which may vary based on the particular strain of interest.
For photostimulation, we have utilized and adapted the setup developed by the Shepherd Lab at Northwestern University, described in a recent protocol29. Although optimized for glutamate uncaging, this setup is amenable also for photostimulation of channelrhodopsin without modification. This setup enables precise spatial control and positioning of the loci of photostimulation, which enables an assessment of the functional topography of channelrhodopsin innervation of recorded neurons. Although spatial control is enhanced with this method of photostimulation, it is more cost prohibitive than methods utilizing LED based field illumination20. In addition, the nonuniformity of brain tissue is a limitation of optogenetics, or any techniques involving manipulation of light. In practice though, it is rarely a factor since results are interpreted mostly in a qualitative manner. In case more quantitative comparison between brain regions is needed, one can always use maximum light intensity to achieve &#34;saturated&#34; responses.
In addition, we have used both Tidelwave and Ephus software programs written in Matlab for controlling the laser-scanning photostimulation and data acquisition25,26. However, it is recommended that those setting up a similar system employ the newer Ephus software26, which has support available through the Janelia Farms development site30. A key step is choosing which of these software packages to use, since their hardware requirements differ slightly. In addition, the map analysis programs are tailored to their respective data acquisition programs.
Several of these steps can be modified to suit the practitioner&#39;s particular setup. Injections of viral solutions can be performed with delivery mechanisms besides a syringe, e.g. capillary glass pipettes pulled to tip diameters of ~40 &#956;m also work well. For surgeries, a nonstereotaxic headholder can be substituted if such coordinates are not required for guiding injections. In addition, ACSF and intracellular solution compositions may be altered depending on the particular experimental requirements, e.g. substituting a cesium or high chloride intracellular solution to isolate inhibitory currents. Also, an alternative to imaging of expression patterns in fixed slices may also be done in the live slice on a suitable imaging rig with a bath perfusion system.
In summary, this protocol will enable the practitioner to characterize the functional topography and synaptic physiology of projections in vitro, which may not be possible using standard methods, e.g. characterizing long-range projections that are unable to be preserved in a slice. The transfection protocol described can be applied to in vivo&#160;optogenetic experiments1-3,12,14, since the delivery mechanism is the same. Finally, although this protocol describes transfection of mice, many of the viral vectors available can be used to theoretically transfect and express these opsins in a wide range of mammalian species, thus enabling a myriad of comparative and species-specific studies1-3.

The elaborate circuitry of the mammalian forebrain poses an experimental challenge to dissecting the functional roles of its intertwined components. Untangling these relationships has been aided recently by the development of optogenetic methods for specifically exciting and inhibiting neuronal populations and their projections1-3. These approaches make use of the genetically controlled expression of membrane channels responsive to light4-6. Of particular note, channelrhodopsin, a light-gated ion channel, depolarizes neuronal membranes following optical stimulation, leading to excitation of the expressing neuron and its projections4,7. Conversely, halorhodopin, a light-gated chloride pump, hyperpolarizes the membrane, thus inhibiting the targeted projection6,8,9. Several variants of these opsins have been engineered, multiple vectors produced for transfection and expression in neuronal populations of interest, and transgenic mouse lines engineered that constitutively express these opsins or that enable cre-lox mediated control of their expression1,2,10,11.
The utility of these optogenetic methods and materials has revolutionized modern experimental neuroscience, enabling the dissection of the role of specific neural circuits in the physiology and behavior of the organism12-15. In particular, these methods can be powerfully applied to the functional dissection of brain circuits using in vitro slice preparations16-20. In this context, one of the main advantages of this method is the ability to characterize the physiology of long range projections that are difficult, if not impossible, to preserve in a thick slice, such as the commissural connections between cerebral cortical hemispheres16. Furthermore, this technology enables manipulations not possible with other methods, such as the selective inhibition of projections with fine temporal precision using halorhodopsin6,8,9. Moreover, the cell-type specific expression of these opsins allows isolated components in a neural circuit to be selectively stimulated or inhibited1,2,11,15,21. As such, these novel optogenetic tools enable the unprecedented analysis of the neuronal contributions to physiology and behavior.
Here, we describe a protocol for expressing channelrhodopsin in the mouse brain using adeno-associated viral (AAV) vectors1,2. Specific transfection of targeted neuronal populations can be accomplished using this protocol, with the appropriate choice of promoter constructs or transgenic mice that express Cre-recombinase in the targeted neurons of interest and a &#39;floxed&#39; viral construct1,11,15,21 (Figure&#160;1). As examples here, we examine the corticocortical projections between the primary visual cortex (V1) and the secondary visual cortex (V2) and we also investigate the layer 6 corticothalamic projection from the primary somatorsensory cortex (S1) to the ventroposterior medial nucleus (VPm). We combine this specific expression with laser-scanning photostimulation&#160;in vitro&#160;to assess synaptic physiology and functional topography. The setup presented here enables the precise spatial positioning and control of the loci of photostimulation. Finally, we describe the visualization of expression patterns using confocal microscopy.

<table border="0" cellpadding="0" cellspacing="0" width="708">
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		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Polybrene</td>
			<td style="width:175px;">Santa Cruz Biotech</td>
			<td style="width:119px;">SC134220</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Paraformaldehyde 32% solution</td>
			<td style="width:175px;">EMS</td>
			<td align="right" style="width:119px;">15714</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">VECTASHIELD</td>
			<td style="width:175px;">Vector Labs</td>
			<td style="width:119px;">H-1400</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">AAV viral constructs</td>
			<td style="width:175px;">UNC Vector Core</td>
			<td style="width:119px;">NA</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Mouse stereotactic headholder</td>
			<td style="width:175px;">Stoelting</td>
			<td align="right" style="width:119px;">51730</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Nanocool injector</td>
			<td style="width:175px;">Harvard Apparatus</td>
			<td align="right" style="width:119px;">703040</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">UV laser: 355 nm, Nd:YVO4; 1 W;</td>
			<td style="width:175px;">DPSS Lasers, Inc</td>
			<td align="right" style="width:119px;">3500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Galvonometers</td>
			<td style="width:175px;">Cambridge Scanning</td>
			<td align="right" style="width:119px;">6210</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">mirrors</td>
			<td style="width:175px;">Cambridge Scanning</td>
			<td style="width:119px;">6m2003S-A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">mount</td>
			<td style="width:175px;">Cambridge Scanning</td>
			<td style="width:119px;">6102103L</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">cable</td>
			<td style="width:175px;">Cambridge Scanning</td>
			<td style="width:119px;">6010-19-120</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">driver/controller</td>
			<td style="width:175px;">Cambridge Scanning</td>
			<td style="width:119px;">67121-1</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">Shutter - ZM coated (UV-compatible)</td>
			<td style="width:175px;">Uniblitz</td>
			<td style="width:119px;">L23-ZM2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Shutter Controller</td>
			<td style="width:175px;">Uniblitz</td>
			<td style="width:119px;">VMM-D1</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">Current Amplifier</td>
			<td style="width:175px;">Stanford Research Systems</td>
			<td style="width:119px;">SR570</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">6713 board</td>
			<td style="width:175px;">National Instruments</td>
			<td style="width:119px;">777741-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">2110 break-out box</td>
			<td style="width:175px;">National Instruments</td>
			<td style="width:119px;">777643-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">PCI-MIO-16E-4</td>
			<td style="width:175px;">National Instruments</td>
			<td style="width:119px;">777383-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">BNC-2090</td>
			<td style="width:175px;">National Instruments</td>
			<td style="width:119px;">777270-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Cables</td>
			<td style="width:175px;">National Instruments</td>
			<td style="width:119px;">184749-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">MATLAB</td>
			<td style="width:175px;">Mathworks</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">TCS SP2</td>
			<td style="width:175px;">Leica Microsystems</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Laser-scanning photostimulation of projections between the primary visual area (V1) and the secondary visual area (V2) in the mouse was accomplished following injections of pAAV-CaMKIIa-hChR2(H134R)-EYFP into V1 of a Balb/C mouse (Figure 3). The CaMKIIa promoter in this viral construct drives specific expression of channelrhodopsin-EYFP in excitatory glutamatergic neurons1,15. After preparation of slices for physiological recording, expression of channelrhodopsin-EYFP fibers, i.e. epifluorescence of the conjugated EYFP, was observed traversing from V1 to V2 (Figure 3A). In addition, the corticothalamic fibers in the lateral geniculate nucleus (LGN) were also observed (Figure 3A). Whole-cell patch clamp recordings from a neuron in V2 revealed EPSCs elicited following laser-scanning photostimulation of channelrhodopsin-EYFP expressing fibers in the lower layers of V2 (Figure 3B).
In another example, laser-scanning photostimulation of the corticothalamic projections from layer 6 of the primary somatosensory cortex (S1) to the ventroposterior nucleus (VP) was accomplished using this protocol. Here, we used a Cre-Lox approach to specifically target channelrhodopsin expression in the layer 6 neurons and corticothalamic projections of interest. Explicitly, we injected pAAV-Ef1a-DIO-hChR2(H134R)-EYFP-WPRE-pA into S1 of B6.Cg-Tg(Ntsr1-cre)GN220Gsat/Mmcd mice (Figures 1 and&#160;4). This transgenic mouse strain expresses Cre-recombinase specifically in layer 6 corticothalamic neurons21, and therefore results in ChR-YFP expression only in those neurons following transfection by the &#39;floxed&#39; AAV construct. After preparation of slices, channelrhodopsin-EYFP fibers were observed in the layer 6 neurons and fibers projecting to layer 4 and VP (Figures 4A and 4B). Physiological recording from a VPm neuron using whole-cell patch clamp revealed that EPSCs could be elicited following laser-scanning photostimulation near the recorded neuron (Figures 4C and 4D).
The morphology of labeled fibers was assessed more readily following physiological recordings. Here, we resectioned (50 &#956;m) slice preparations with channelrhodopsin-EYFP expression in layer 6 corticothalamic neurons (Figure 5). This revealed a fine pattern of labeled fibers traversing from layer 6 to layer 4, where they terminated and arborized extensively (Figure 5A). In addition, corticothalamic fibers were observed entering the white matter traversing towards the thalamus (Figures 5A and 5B).
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/50915/50915fig1.jpg" width="500px" /> 
Figure 1. Schematic illustration of targeted expression of channelrhodopsin via a Cre-Lox approach. Transgenic mice expressing Cre-recombinase in the neuronal populations of interest are injected with a &#39;floxed&#39; virus encoding channelrhodopsin fused to a fluorescent protein, such as YFP. Following transfection, only neurons with Cre-recombinase will express channelrhodopsin-EYFP. <a href="https://www.jove.com/files/ftp_upload/50915/50915fig1highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/50915/50915fig2.jpg" width="500px" /> 
Figure 2. Schematic illustration of photostimulation setup. (A) Diagram of the laser-scanning photostimulation setup. The laser beam is guided successively through an open beam&#160;system to a shutter and a neutral density (N.D.) filter. A coverslip reflects part of the beam to a photodiode connected to an amplifier for laser power measurements. Mirror galvanometers are used to position the laser beam through a scan lens, which guides the beam through a side port on the top of the microscope. The beam is then reflected down through the objective lens to the slice preparation using a dichroic mirror. For further details see Lam and Sherman23&#160;and Shepherd29. (B) Example of the photostimulation grid pattern for the representative experiment shown in Figure 3. The 16 x 16 stimulation grid is overlaid on a photomicrograph of the slice preparation. A red star indicates the location of a neuron recorded using whole-cell patch clamp. The laser-beam position is guided to each of the 256 positions in the stimulation grid by computerized positioning of the mirror galvanometers. The laser positions of the 1st 5 stimulation sites are depicted in the figure. The responses from the recorded neuron to photostimulation at each of the sites is depicted in Figure 3. <a href="https://www.jove.com/files/ftp_upload/50915/50915fig2highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/50915/50915fig3.jpg" width="500px" /> 
Figure 3. Laser-scanning photostimulation of corticocortical projections from primary visual cortex (V1) to secondary visual cortex (V2) in the mouse. (A) Epifluorescent image of the slice preparation showing channelrhodopsin-EYFP fibers from V1 to V2 and in the lateral geniculate nucleus (LGN). Specific expression of channelrhodopsin-EYFP in excitatory glutamatergic neurons is driven by a viral construct containing the CaMKIIa promoter. (B) Excitatory postsynaptic currents (EPSCs) elicited from a neuron in V2 following laser-scanning photostimulation. Red star indicates recording site. Scale bars: (A) 500 &#181;m, (B) 200 pA, 100 msec. <a href="https://www.jove.com/files/ftp_upload/50915/50915fig3highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/50915/50915fig4.jpg" width="500px" /> 
Figure 4. Laser-scanning photostimulation of corticothalamic projections from primary somatosensory cortex (S1) to the ventroposterior nucleus (VP) in the mouse. (A) Epifluorescent image of the slice preparation showing channelrhodopsin-EYFP fibers in S1. (B) Epifluorescent image of labeled-fibers in VP. (C) Excitatory postsynaptic currents (EPSCs) elicited from a VP neuron following laser-scanning photostimulation. (D) Plot of mean EPSCs. Red star indicates recording site. Boxed traces in (C) correspond with boxed values in (D). Scale bars: (A) 250 &#181;m, (B) 250 &#181;m, (C) 100 pA, 100 msec. <a href="https://www.jove.com/files/ftp_upload/50915/50915fig4highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/50915/50915fig5.jpg" width="500px" /> 
Figure 5.&#160;Imaging of layer 6 neurons expressing channelrhodopsin-EYFP in the primary somatosensory cortex (S1) of the mouse. (A-B) Labeled neurons in layer 6 and fibers projecting to layer 4 and the white matter. Scale bars: (A) 250 &#181;m, (B) 250 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50915/50915fig5highres.jpg" target="_blank">Click here to view larger image</a>.

Watch this Scientific Journal Video about Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits at JoVE.com

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

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 ...

laser-scanning-photostimulation-optogenetically-targeted-forebrain

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Neuroscience

9.1K Views.  Louisiana State University. We describe a method for characterizing the functional topography and synaptic properties of forebrain circuits using an optogenetic approach to photostimulate neuronal populations in vitro.

Video: Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

Ultrasound-Guided Microinjection into the Mouse Forebrain In Utero at E9.5

In utero survival surgery in mice permits the molecular manipulation of gene expression during development. However, because the uterine wall is opaque during early embryogenesis, the ability to target specific parts of the embryo for microinjection is greatly limited. Fortunately, high-frequency ultrasound imaging permits the generation of images that can be used in real time to guide a microinjection needle into the embryonic region of interest. Here we describe the use of such imaging to guide the injection of retroviral vectors into the ventricular system of the mouse forebrain at embryonic day (E) 9.5. This method uses a laparotomy to permit access to the uterine horns, and a specially designed plate that permits host embryos to be bathed in saline while they are imaged and injected. Successful surgeries often result in most or all of the injected embryos surviving to any subsequent time point of interest (embryonically or postnatally). The principles described here can be used with slight modifications to perform injections into the amnionic fluid of E8.5 embryos (thereby permitting infection along the anterior posterior extent of the neural tube, which has not yet closed), or into the ventricular system of the brain at E10.5/11.5. Furthermore, at mid-neurogenic ages (~E13.5), ultrasound imaging can be used direct injection into specific brain regions for viral infection or cell transplantation. The use of ultrasound imaging to guide in utero injections in mice is a very powerful technique that permits the molecular and cellular manipulation of mouse embryos in ways that would otherwise be exceptionally difficult if not impossible.

Ultrasound-Guided Microinjection into the Mouse Forebrain In Utero at E9.5

In utero survival surgery in mice permits the molecular manipulation of gene expression during development. Here we describe the use of high-frequency ultrasound imaging to guide the injection of retroviral vectors into the mouse brain at embryonic day (E) 9.5.

In utero survival surgery in mice permits the molecular manipulation of gene expression during development.  However, because the uterine wall is opaque ...

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

Targeted Training of Ultrasonic Vocalizations in Aged and Parkinsonian Rats

Voice deficits are a common complication of both Parkinson disease (PD) and aging; they can significantly diminish quality of life by impacting communication abilities. 1, 2 Targeted training (speech/voice therapy) can improve specific voice deficits,3, 4 although the underlying mechanisms of behavioral interventions are not well understood. Systematic investigation of voice deficits and therapy should consider many factors that are difficult to control in humans, such as age, home environment, age post-onset of disease, severity of disease, and medications. The method presented here uses an animal model of vocalization that allows for systematic study of how underlying sensorimotor mechanisms change with targeted voice training. The ultrasonic recording and analysis procedures outlined in this protocol are applicable to any investigation of rodent ultrasonic vocalizations.
The ultrasonic vocalizations of rodents are emerging as a valuable model to investigate the neural substrates of behavior.5-8 Both rodent and human vocalizations carry semiotic value and are produced by modifying an egressive airflow with a laryngeal constriction.9, 10 Thus, rodent vocalizations may be a useful model to study voice deficits in a sensorimotor context. Further, rat models allow us to study the neurobiological underpinnings of recovery from deficits with targeted training.
To model PD we use Long-Evans rats (Charles River Laboratories International, Inc.) and induce parkinsonism by a unilateral infusion of 7 &mu;g of 6-hydroxydopamine (6-OHDA) into the medial forebrain bundle which causes moderate to severe degeneration of presynaptic striatal neurons (for details see Ciucci, 2010).11, 12 For our aging model we use the Fischer 344/Brown Norway F1 (National Institute on Aging).
Our primary method for eliciting vocalizations is to expose sexually-experienced male rats to sexually receptive female rats. When the male becomes interested in the female, the female is removed and the male continues to vocalize. By rewarding complex vocalizations with food or water, both the number of complex vocalizations and the rate of vocalizations can be increased (Figure 1).
An ultrasonic microphone mounted above the male's home cage records the vocalizations. Recording begins after the female rat is removed to isolate the male calls. Vocalizations can be viewed in real time for training or recorded and analyzed offline. By recording and acoustically analyzing vocalizations before and after vocal training, the effects of disease and restoration of normal function with training can be assessed. This model also allows us to relate the observed behavioral (vocal) improvements to changes in the brain and neuromuscular system.

Voice disorders are debilitating in aging and Parkinson disease. The ultrasonic vocalizations of rats, also affected by these conditions, can be used to study these voice disorders, their neural substrates, and the nature of functional recovery with behavioral intervention.

Voice deficits are a common complication of both Parkinson disease (PD) and aging; they can significantly diminish quality of life by impacting ...

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 ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Isolation and Culture of Endothelial Cells from the Embryonic Forebrain

Embryonic brain endothelial cells can serve as an important tool in the study of angiogenesis and neurovascular development and interactions. The two vascular networks of the embryonic forebrain, pial and periventricular, are spatially distinctive and have different origins and growth patterns. Endothelial cells from the pial and periventricular vascular networks have unique gene expression profiles and functions. Here we present a step-by-step protocol for isolation, culture, and verification of pure populations of endothelial cells from the periventricular vascular network (PVECs) of the embryonic forebrain (telencephalon). In this approach, telencephalon devoid of pial membrane obtained from embryonic day 15 mice is minced, digested with collagenase/dispase, and dispersed mechanically into a single cell suspension. PVECs are purified from cell suspension using positive selection with anti-CD-31/PECAM-1 antibody conjugated to MicroBeads using a strong magnetic separation method. Purified cells are cultured on collagen 1&#160;coated culture dishes in endothelial cell culture medium until they become confluent and further subcultured. PVECs obtained with this protocol exhibit cobblestone and spindle shaped phenotypes, as visualized by phase-contrast light microscopy and fluorescence microscopy. Purity of PVEC cultures was established with endothelial cell markers. In our hands, this method reliably and consistently yields pure populations of PVECs. This protocol will benefit studies aimed at gaining mechanistic insights into forebrain angiogenesis, understanding PVEC interactions, and cross-talks with neuronal cell types and holds tremendous potential for therapeutic angiogenesis.

This video demonstrates an easy and reliable strategy for preparation of pure cultures of endothelial cells from the embryonic forebrain within 10-12 days and will be useful for research focused on many aspects of cerebral angiogenesis.

Embryonic brain endothelial cells can serve as an important tool in the study of angiogenesis and neurovascular development and interactions. The two ...

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.

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

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 ...

Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an increasingly detailed view of the circuits underpinning spinal pain processing. However, the effects of these connections for broader network activity in the DH remain less well understood because most studies focus on the activity of single neurons and small microcircuits. Alternatively, the use of microelectrode arrays (MEAs), which can monitor electrical activity across many cells, provides high spatial and temporal resolution of neural activity. Here, the use of MEAs with mouse spinal cord slices to study DH activity induced by chemically stimulating DH circuits with 4-aminopyridine (4-AP) is described. The resulting rhythmic activity is restricted to the superficial DH, stable over time, blocked by tetrodotoxin, and can be investigated in different slice orientations. Together, this preparation provides a platform to investigate DH circuit activity in tissue from na&#239;ve animals, animal models of chronic pain, and mice with genetically altered nociceptive function. Furthermore, MEA recordings in 4-AP-stimulated spinal cord slices can be used as a rapid screening tool to assess the capacity of novel antinociceptive compounds to disrupt activity in the spinal cord DH.

The combined use of microelectrode array technology and 4-aminopyridine-induced chemical stimulation for investigating network-level nociceptive activity in the spinal cord dorsal horn is outlined.