The authors wish to thank J. Vazquez for illustrations, D. Kaplan and C. Preda for reagents and helpful guidance, and the labs of B. Sabatini and C. Harvey for in vivo imaging. Microscopy was made possible by M. Ocana and the Neurobiology Imaging Center, supported in part by the Neural Imaging Center as part of a National Institute of Neurological Disorders and Stroke (NINDS) P30 Core Center grant (NS072030). This work was supported by the GVR Khodadad Family foundation, the Nancy Lurie Marks foundation, and by NIH grants, NINDS R21NS093498, U01NS108177 and NINDS R35NS097284 to W.G.R, and by an NIH postdoctoral fellowship F32NS101889 to C.H.C.

All experiments involving animals were performed in accordance with protocols approved by the Harvard Standing Committee on Animal Care following guidelines described in the US NIH Guide for the Care and Use of Laboratory Animals. Adult C57BL/6 mice of either sex (6-15 weeks of age) were used for all experiments.
1. Obtain Aqueous Silk Fibroin
<ol>
	<li>Prepare or purchase aqueous silk fibroin (5-7.5% w/v).</li>
</ol>
2. Mix Aqueous Silk with AAV Expression Vectors
<ol>
	<li>Choose an AAV expression vector to drive the optogenetic protein or fluorescent indicator of choice. 
	Note: To minimize the volume of silk/AAV that must be applied to implants while still driving robust expression, stock-titer AAV (stock titers typically obtained from vector cores are around ~1013 gc/mL) is recommended.</li>
	<li>Immediately prior to coating implants, thaw an aliquot of AAV and combine with 5-7.5% aqueous silk fibroin (this mixture will be referred to as silk/AAV). In a 200 &#181;L PCR tube, mix aqueous fibroin and AAV in a 1:1 ratio (for cranial windows use 1:4) immediately prior to the application. Gently pipette the solution in and out several times to thoroughly mix the fibroin and AAV.</li>
	<li>Keep silk/AAV mixture on ice prior to use.</li>
</ol>
3. Prepare Equipment for Fabrication and Storage of Silk/AAV-Coated Devices
<ol>
	<li>Procure equipment for coating optical fibers and Gradient-Index (GRIN) lenses (Figures 1, 2).
	<ol>
		<li>Construct a stable ferrule holder. To hold ceramic ferrules, drill 1.25 mm holes in a block of &#188;&#8221; sheet acrylic. Tap holes to insert set screws from the side to hold ferrules in place. 
		&#8203;Note: Any clamp can be used for this purpose.</li>
		<li>Position a manipulator with sub-millimeter precision to move the optical fibers (stereotaxic apparatus or other precision micromanipulator).</li>
		<li>Assemble a stable holder to position the microinjector.</li>
		<li>Use a stereoscope to visualize optical fibers and silk droplet.</li>
		<li>Position a light source to illuminate the optical fibers.</li>
	</ol>
	</li>
	<li>Prepare equipment for coating cranial windows (Figure 3).
	<ol>
		<li>Choose any P10 pipettor.</li>
		<li>Obtain a container with lid. 
		Note: Any container with a silicone bottom is suggested&#8212;the soft bottom facilitates lifting up cranial windows.</li>
	</ol>
	</li>
	<li>Prepare equipment to store finished implants (Figure 4).
	<ol>
		<li>Obtain a small (1-5 L) vacuum chamber.</li>
		<li>Make sure that there is space to store implants in a 4 &#176;C refrigerator.</li>
	</ol>
	</li>
</ol>
4. Apply Silk/AAV Film to Devices
<ol>
	<li>Coating optical fibers to drive focal expression at the fiber tip
	<ol>
		<li>Prepare chronic fiber implants as previously described4.</li>
		<li>Prior to use, rinse implants with ethanol, then with ultrapure water to ensure that the optical fibers are clean. 
		Note: Silk films adhere more reliably to clean glass surfaces.</li>
		<li>Prepare a device to hold fiber ferrules. For typical 1.25 mm diameter ferrules, use a block &#188; inch clear acrylic, with ~1.3 mm holes, and tapped set screws entering from the side to hole implants firmly in place (Figure 1A).</li>
		<li>Mount the ferrule holder into a stereotaxic apparatus (or any manipulation solution with submillimeter precision) equipped with a microinjector. Place the ferrule holder above the microinjector and apply the silk/AAV mixture from below. 
		Note: This is because applications of large volumes from above resulted in silk/AAV that was not restricted to the tip. However, the application of many small sequential volumes from above or below can produce AAV/silk deposits that are confined to the tip (although we prefer to apply from below).</li>
		<li>Pull a standard intracranial injection pipette from borosilicate glass capillary.
		<ol>
			<li>To make it easier to</li>
			<li>To produce an injection tip with a clean flat tip of the desired diameter, hold one pipette in each hand and use the thicker part of the taper on one pipette to score the other pipette at the desired break location.</li>
			<li>Gently rub back and forth in a sawing motion (the glass-on-glass scoring method).</li>
			<li>After scoring the pipette, apply gentle pressure to the tip of the scored pipette with the body of the other pipette to achieve a clean break.</li>
		</ol>
		</li>
		<li>Position a stereoscope to give a clear view of the optical fiber faces. 
		Note: Magnification should be sufficient to accurately position the injection pipette above the face of optical fibers.</li>
		<li>Insert fiber implants into holder with the brain-side of the optical fiber facing downward.</li>
		<li>Load the injection pipette with silk/AAV solution, as for any standard intracranial injection5. Load the amount required for the number of implants being made, plus ~30% extra to accommodate losses due to pipettes clogging. For example, if 10 implants are being made, then load with 100 nL deposits and withdraw ~1.3 &#181;L. 
		Note: Silk/AAV may dry at the pipette tip in between ejections, which can clog the pipette. Large diameter pipettes (50-100 &#181;m) are less likely to clog. Clogs can be dislodged by gentle brushing down the tip of the pipette with a wet paper wipe or alcohol swab.</li>
		<li>Maneuver the injection pipette until it is touching or nearly touching the center of the optical fiber surface. Eject 10-20 nL of silk/AAV solution. Withdraw the pipette. 
		Note: The rate of delivery is not critical, but typical rates are 5-20 nL/s.</li>
		<li>Observe the bolus of silk/AAV on the flat surface which appears as a liquid dome that dries to a flat film within ~1 min (Figure 1B).</li>
		<li>Repeat steps 4.1.9-4.1.10 until the desired amount of silk/AAV is deposited (a total of 20-200 nL for most applications). When preparing multiple implants, apply silk/AAV to one implant and then move on to coat other implants before returning to the first.</li>
		<li>Allow 1 h for drying before moving implants.</li>
		<li>Vacuum desiccate overnight at ~125 Torr (-25 in. Hg), 4 &#176;C. Do this by placing the entire ferrule holder into a vacuum chamber.</li>
		<li>Evaluate the shape and position of the resulting silk film under a high-power microscope. Ensure that films are confined to the tip of the optical fiber surface, be relatively thin (&#62;100 &#181;m), and symmetrical (Figure 1C). 
		Note: Large or asymmetrical silk/AAV films may dislodge from the fiber during implantation (Figure 1D). The most common cause of problems arises from the application of single large volumes rather than the sequential application of many small volumes.</li>
	</ol>
	</li>
	<li>Coating tapered optical fibers to drive expression along the fiber axis
	<ol>
		<li>Obtain tapered optical fiber implants and perform steps 4.1.2-4.1.8, except that the tapered fiber is positioned laterally such that it is perpendicular to the injector (Figure 2A). Position the injector above the tapered fiber. 
		Note: Loading liquid droplets onto tapered fibers poses added challenges, because surface tension tends to cause droplets to jump back onto the injection pipette or migrate up the tapered fiber. Smaller injection pipettes (30-50 &#181;m diameter) help to overcome this problem but increase the risk that the injection pipette will clog. Due to surface tension, droplets tend to adhere to the area of largest surface area, so the optimal injection pipette size is dependent on the size of the tapered fiber and one&#8217;s tolerance for the occasional clog.</li>
		<li>Position the silk/AAV injection pipette against the side of the optical fiber at the beginning of the taper. Make sure the injection pipette is touching the optical fiber.</li>
		<li>Eject 20 nL of silk/AAV to start the coating process. Ensure that the droplet adheres to the optical fiber and remains at the interface of the fiber/pipette. Gently wick the droplet towards the end of the fiber tip as the silk/AAV dries (~45 s). Keep the injecting pipette in contact with the drying droplet to avoid clogging the pipette tip. 
		Note: Each deposit should coat approximately 400 &#181;m of the tapered fiber (Figure 2B).</li>
		<li>When the first bolus has dried almost completely, eject another 20 nL and continue wicking the droplet along the taper. 
		Note: The liquid silk will adhere to the dried silk, anchoring one end of the droplet as the pipette moves along the taper.</li>
		<li>Repeat step 4.2.4 by ejecting small amounts of silk/AAV, and gradually drawing the solution up the side of the taper. 5-6 ejections are sufficient to traverse the surface of a 2.5 mm taper.</li>
		<li>To drive more uniform expression around all sides of the fiber, rotate the fiber and repeat steps 4.2.2-4.2.5 until the desired amount of silk/AAV has been deposited.</li>
		<li>If a hanging strand of dried silk/AAV extends beyond the fiber tip, carefully cut the strand with scissors, or use the ejection pipette to bend the strand back and adhere it to the taper of the fiber.</li>
		<li>Allow 1 h for drying before moving implants.</li>
		<li>Vacuum desiccate overnight in 4 &#176;C. The entire ferrule holder can be placed into a vacuum chamber.</li>
		<li>Evaluate the shape and position of the resulting silk film under a high-power microscope. 
		Note: Films need not be entirely uniform but should not have bumps that extend more than 100 &#181;m beyond the surface of the fiber to minimize damage to surrounding tissue during implantation (Figure 3C). To minimize film size, it is critical that each droplet is completely dry before subsequent deposits are made.</li>
	</ol>
	</li>
	<li>Coating GRIN lens implants
	<ol>
		<li>Obtain GRIN lenses6,7 and repeat steps 4.1.2-4.1.8. The injector can be mounted above.</li>
		<li>Deposit silk/AAV in a single ejection (1 &#181;L for a 1.0 mm diameter lens). 
		Note: This will yield a dome of liquid that adheres to the face of the lens, and dries to produce a uniform film (100-200 &#181;m thick). However, in the event that a single large ejection dries unevenly and produces a film that is thicker near the edges of the GRIN lens, try depositing multiple smaller droplets (100-200 nL) in the center of the lens surface (allowing each droplet to dry before depositing the next) to ensure that the film will drive expression at the center of the field of view.</li>
		<li>Allow 1 h for drying before moving implants.</li>
		<li>Evaluate the shape and position of the resulting silk film under a high-power microscope to ensure that the film covers the surface of the lens.</li>
	</ol>
	</li>
	<li>Coating glass cranial windows
	<ol>
		<li>Prepare glass cranial windows by adhering two 3 mm diameter round coverslips (no. 1 thickness) to one 5 mm diameter window with optical adhesive (for details, see Goldey et al. 20148).</li>
		<li>Mix silk:virus in a ratio of 1:4 to reduce the total amount of silk in the film. Excessive quantities of silk do not dissolve beneath cranial windows after implantation. Titration experiments may be required to determine the ratio and volume that gives the desired expression profile.</li>
		<li>Hand pipette a 5 &#181;L droplet onto the surface of the 3 mm (brain facing) coverslip. The droplet should spread out to cover the entire glass surface (Figure 3).</li>
		<li>Allow 2-3 h for drying before moving windows.</li>
	</ol>
	</li>
</ol>
5. Storing silk/AAV-coated implants
<ol>
	<li>Store silk/AAV-coated optical fibers in a cooled vacuum desiccator (~125 Torr, 4 &#176;C) prior to use (Figure 4A).</li>
	<li>Do not store cranial windows and GRIN lenses under vacuum, as large silk films stored under vacuum fail to fully dissolve after implantation. Implant cranial windows and GRIN lenses immediately after drying, or within a day of manufacture if stored at atmospheric pressure and 4 &#176;C.</li>
</ol>
6. Implanting the Devices
<ol>
	<li>Prepare animals for implant surgery as previously described4.
	<ol>
		<li>Briefly, anesthetize mice with an intraperitoneal injection of ketamine/xylazine (100/10 mg/kg) and check the depth of anesthesia using a gentle toe-pinch. Shave the skull in the area of the implant and clean the scalp with iodine and alcohol.</li>
		<li>Mount animals in a stereotaxic device and supplement anesthesia using a mixture of oxygen and isoflurane (1-2%). Make an incision in the scalp over the area of interest, and perform a craniotomy large enough to accommodate the implant.</li>
	</ol>
	</li>
	<li>Implant optical fibers9 and microendoscope lenses10 according to previously published procedures. Handle implants with care, as the silk/AAV deposit can be knocked off by an imperfect craniotomy, or by the implant catching on the edge of the skull. Lower the implant into the brain slowly (~2 mm/min).</li>
	<li>Implant cranial windows as describe previously8. Do not touch the coated side of the window and avoid rinsing the window with fluid if performing gavage, as this may wash away the virus. To achieve maximal expression, perform a durotomy.</li>
</ol>
7. Evaluating the Expression and Troubleshooting
<ol>
	<li>To evaluate expression of virally-expressed proteins, allow ~2-3 weeks for the virus to drive expression, then perform intracardial perfusion with 4% paraformaldehyde in phosphate buffered saline solution11 and process brain tissue for fluorescent microscopy12.</li>
	<li>Evaluate the expression by using fluorescent microscopy to image the expression pattern of fluorophore-tagged optogenetic proteins.</li>
	<li>If the level of expression is insufficient, increase the amount of virus in coatings by either increasing the total volume of the silk/AAV coating, or preferably by using a higher titer virus.</li>
</ol>

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The authors have nothing to disclose.

The use of silk/AAV to target the expression of optogentic proteins overcomes limitations of approaches that are currently in use. Although many studies successfully use AAV injections to express optogenetic proteins, it is challenging to align expression to the tip of optical fibers, to regions around the length of tapered fibers, and to the viewing region of a GRIN lens. Because of misalignment between optical components and optogenetic expression, stereotaxic injections can be unreliable, and many experiments fail. The silk/AAV labelling method we describe here solves this problem. It also simplifies the procedure by eliminating a second surgical step and in some cases eliminating the need for a second surgery. It can also be difficult to use viruses to obtain widespread expression beneath cranial windows, and experimenter typically perform long surgeries to inject virus in multiple locations. The ability to obtain widespread expression over large cortical regions by simply coating cranial windows with silk/AAV is a simplification that eliminates the need for the many invasive injections.
Another potential advantage of the silk/AAV method is that it might induce less inflammation in the neural tissues compared to viral injections. Injecting high-titer AAV into the brain can cause inflammatory responses such as reactive astrocytosis that have the potential to alter cellular and circuit properties14,15 (although such potential complications are usually ignored). Silk films induce little immunogenic response on their own13 and silk/AAV films are expected to release virus over the course of many hours or days16, which may lower the viral load in the surrounding tissue and reduce immunogenic responses. With conventional approaches in which implantation of a device is preceded by an AAV injection, inflammatory responses can arise from both the implantation and the injection. In the future it will be desirable to systematically compare conventional approaches and the silk/AAV method to determine whether silk/AAV films reduces overall inflammatory responses.
Several steps are critical to the successful use of silk/AAV films. Most importantly, the coating of optical fibers must be done carefully as described in the Methods and the location of the dried films should be assessed carefully by visual inspection under a microscope to ensure that the films are compact, in the correct location, and adhere to the face of the optical fiber. Any silk/AAV on the sides of the optical fiber will lead to expression outside the region of interest, and misshaped films that protrude beyond the face of the fiber may break off during implantation and lead to unreliable or no expression. The techniques we describe for applying silk/AAV to implantable devices can be adapted to use of any materials are that are readily available and allow the precise deposition of small volumes of silk/AAV.
A bit of practice is required to achieve accurate, reproducible results. If expression is observed along the track of the fiber, it is likely that the silk film dried on the side of the fiber rather than the fiber face. Repeat the manufacturing process and closely inspect dried implants for signs that films are drying on the side of the fiber. Because silk/AAV films are optically transparent, it may help to practice applying silk mixed with dye (Fast Green or a similar dye) to better visualize the shape of resulting films (Figure 2C). If there is no expression, it is probable that the silk film dislodged from the fiber tip during implantation. We suggest the use of stock-titer virus when making implants. For optical fibers, this reduces the total volume that must be applied to small diameter fibers. If the size of the coating is a concern consider waiting longer between each 10 nL application to allow complete drying of the deposited droplet. Silk/AAV droplets dry faster under a warm lamp. For cranial windows, high-titer virus may be necessary to supply adequate viral load across the pia or dura. Certain types of implants may dissolve silk and release AAV more readily than others. We have found that cranial windows implanted over the surface of the brain require a lower ratio of silk/virus to achieve reliable expression, perhaps owing to different cerebral spinal fluid dynamics or protease activity. If expression cannot be increased by increasing effective AAV concentrations, decreasing the volume of aqueous silk is a plausible alternative.
Finally, it is important to store the optical components properly and implant them quite soon after they are prepared. We have shown that coated fibers that are refrigerated under vacuum can be stored for many days prior to use. Vacuum storage removes residual moisture17 which may reduce the solubility of silk films, and also help maintain viral efficacy. Ideally, optical fibers should be implanted within 24 hours of fabrication. However, we find that silk/AAV-coated fibers stored under vacuum drive similar levels of expression when implanted 7 days after fabrication (Figure 4B). In contrast, coated cranial windows and GRIN lenses drove the most reliable expression when they were dried at room temperature and used within hours of preparation. The reason for this disparity remains unclear. Further studies may be needed to refine preparation and storage conditions to further extend the storage time.
Silk/AAV-coated cranial windows have considerable potential because they drastically shorten surgical times and are extremely simple to manufacture, but at present this method has limitations. Coated cranial windows uniformly label large areas of cortex and drive sufficient expression in layer 2/3 for GCaMP imaging, with somewhat less expression in deeper layers. However, stereotaxic injections drive more robust expression and provide more control over the layers targeted for expression. Reliable expression was only achieved when the dura was removed. Although the dura is often removed for many 2-photon imaging experiments to improve image quality8, for many experiments it is desirable to obtain labelling in a less invasive manner. We have therefore explored our ability to use silk/AAV to label cortical regions without removing the dura. We obtained some labelling, but it is possible that this was a consequence of damaging the dura in the process of preparing the craniotomy. Further study is needed for coated cranial windows to be used to reliably label the cortex without removing the dura.
The preparation of aqueous silk fibroin from cocoons of Bombyx Mori is described in detail in Rockwood et al. (2011)18. Aqueous silk fibroin is now commercially available (5% w/v). Although most of our experiments were performed using aqueous silk fibroin stocks prepared in our lab (5-7.5% w/v), we have obtained similar results using commercial aqueous fibroin. Aqueous fibroin is stable at 4 &#176;C for up to 3 months, after which it spontaneously transitions from liquid to hydrogel18. We recommend that fibroin stocks be divided into ~1 mL aliquot and stored at -80 &#176;C. A 1 mL working aliquot (sufficient for coating hundreds of implants) can be stored at 4 &#176;C and used until it begins to gel. Be careful not shake, vortex, agitate or aggressively pipette aqueous fibroin, as shear forces can lead to gelation19,20.
Silk/AAV films permit a wide range of expression patterns, from widespread cortical expression under cranial windows, to precise subcortical expression at the tip of a small-diameter optical fibers. These techniques were developed to take advantage of common AAV expression vectors but could likely be used to disperse other expression vectors like Lentiviruses or rabies viruses into the brain. Silk films could also be manufactured into three-dimensional shapes to improve viral release into tissue. For example, in order to drive strong expression beneath cortical windows without the use of a durotomy, cranial windows could be coated with arrays of silk microneedles that would pierce the dura and release virus in deeper cortical layers21. Further refinement will likely lead to improved properties of virus release, and new applications for silk/AAV films.

The past decade has produced an explosion of engineered light-sensitive proteins for monitoring and manipulating neural activity1. Viruses offer unparalleled flexibility for expressing these optogenetic tools in the brain. Compared to transgenic animals, viruses are far easier to produce, transport, and store, allowing fast implementation of the newest optogenetic tools. Expression can be targeted genetically to distinct neuronal populations, and viruses designed for the retrograde transport can even be used to target expression based on neuronal connectivity2.
Viruses are usually introduced with stereotaxic injections, which can be time-consuming and challenging. Precisely targeting small regions can be difficult, while driving expression over broad areas often requires many injections. Moreover, when an optical device is subsequently implanted into the brain to deliver light in vivo, the implant must be properly aligned with the viral injection. Here, we describe an easily-implemented method for delivering viral vectors to the tissue around an implanted device using silk fibroin films3. Silk fibroin is commercially available, well-tolerated by neural tissues, and can be used to produce materials with varied properties. Silk films can be applied to implants using common laboratory equipment like microinjection pipettes or hand pipettes. Silk/AAV films eliminate the requirement for two surgical procedures and ensure that virus-mediated expression is properly aligned to the optical implant. The resulting expression is constrained to the tip of fibers, and results in less unwanted expression along the fiber track than stereotaxic injections.
In addition to producing targeted expression at the tip of small fibers, silk/AAV films can be used to drive widespread (&#62;3 mm diameter) cortical expression beneath cranial windows. In vivo 2-photon imaging of fluorescent activity sensors has become an indispensable tool for evaluating the role of neuronal activity in driving sensory and cognitive processing. However, to drive uniform expression over the broad cortical areas, experimenters often perform multiple injections. These injections can be extremely time-consuming and can lead to inconsistent expression across the field of view. In contrast, silk/AAV-coated cranial windows are extremely easy to manufacture, greatly reduce the time required for surgeries, and most remarkably drive expression hundreds of microns below the cortical surface.

<table>
	<tbody>
		<tr>
			<td>Aqueous silk fibroin</td>
			<td>Sigma</td>
			<td>5154-20ML</td>
			<td>Aqueous Silk Fibroin (5% w/v) for making films</td>
		</tr>
		<tr>
			<td>Microinjector to deposit silk/AAV</td>
			<td>Drummond</td>
			<td>3-000-207</td>
			<td>Nanoject III nanoliter injector</td>
		</tr>
		<tr>
			<td>Manipulator to hold implants</td>
			<td>Narashige</td>
			<td>MM-33</td>
			<td>Micromanipulator</td>
		</tr>
		<tr>
			<td>Stereoscope to visualize silk deposits</td>
			<td>AmScope</td>
			<td>SM-6TX-FRL</td>
			<td>3.5X-45X Trinocular articulating zoom microscope with ring light</td>
		</tr>
		<tr>
			<td>Vacuum chamber to store implants</td>
			<td>Ablaze</td>
			<td>N/A</td>
			<td>3.5 Quart Vacuum Vac Degassing Chamber</td>
		</tr>
		<tr>
			<td>Optional, implant holder for storage</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>To store premade optical fibers, drill a grid of ~4 mm-deep holes with a diameter just larger than the ferrule diameter into a plastic block.</td>
		</tr>
		<tr>
			<td>Optical fiber</td>
			<td>Thorlabs</td>
			<td>FT200EMT</td>
			<td>&#216;200 &#181;m Core Multimode Optical Fiber for fiber implants</td>
		</tr>
		<tr>
			<td>Ferrules</td>
			<td>Kientec</td>
			<td>FZI-LC-230</td>
			<td>LC Zirconia Ferrule for fiber implants</td>
		</tr>
		<tr>
			<td>Various materials for manufacturing chronic fiber implants</td>
			<td>Various</td>
			<td>N/A</td>
			<td>For detailed procedure, see Ung K, Arenkiel BR. Fiber-optic implantation for chronic optogenetic stimulation of brain tissue. Journal of visualized experiments: JoVE. 2012(68).</td>
		</tr>
		<tr>
			<td>Tapered fiber implants</td>
			<td>Optogenix</td>
			<td>Lambda-B</td>
			<td>Tapered fiber implants</td>
		</tr>
		<tr>
			<td>GRIN lenses</td>
			<td>GoFoton</td>
			<td>CLH-100-WD002-002-SSI-GF3</td>
			<td>GRIN lenses</td>
		</tr>
		<tr>
			<td>Small glass cranial windows</td>
			<td>Warner</td>
			<td>64-0726 (CS-3R-0)</td>
			<td>Small round cover glass, #0 thickness</td>
		</tr>
		<tr>
			<td>Large glass cranial windows</td>
			<td>Warner</td>
			<td>64-0731 (CS-5R-0)</td>
			<td>Small round cover glass, #0 thickness</td>
		</tr>
		<tr>
			<td>Various materials for manufacturing cranial windows</td>
			<td>Various</td>
			<td>N/A</td>
			<td>For detailed procedure, see Goldey GJ et al. Removable cranial windows for long-term imaging in awake mice. Nature protocols. 2014 Nov;9(11):2515.</td>
		</tr>
	</tbody>
</table>

in vivo targeted expression of optogenetic proteins using silk/aav films

The quest to understand how neural circuits process information in order to drive behavioral output has been greatly aided by recently-developed optical methods for manipulating and monitoring the activity of neurons in vivo. These types of experiments rely on two main components: 1) implantable devices that provide optical access to the brain, and 2) light-sensitive proteins that change neuronal excitability or provide a readout of neuronal activity. There are a number of ways to express light-sensitive proteins, but stereotaxic injection of viral vectors is currently the most flexible approach because expression can be controlled with genetic, anatomical, and temporal precision. Despite the great utility of viral vectors, delivering the virus to the site of optical implants poses numerous challenges. Stereotaxic virus injections are demanding surgeries that increase surgical time, increase the cost of studies, and pose a risk to the animal's health. The surrounding tissue can be physically damaged by the injection syringe, and by immunogenic inflammation caused by the abrupt delivery of a bolus of high-titer virus. Aligning injections with optical implants is especially difficult when targeting small regions deep in the brain. To overcome these challenges, we describe a method for coating multiple types of optical implants with films composed of silk fibroin and Adeno-associated viral (AAV) vectors. Fibroin, a polymer derived from the cocoon of Bombyx mori, can encapsulate and protect biomolecules and can be processed into forms ranging from soluble films to ceramics. When implanted into the brain, silk/AAV coatings release virus at the interface between optical elements and the surrounding brain, driving expression precisely where it is needed. This method is easily implemented and promises to greatly facilitate in vivo studies of neural circuit function.

To assess the success of silk/AAV films in driving expression, we perfused animals 2-3 weeks after implantation and prepared brain slices from the region of interest. Fluorescence images of fluorophore-tagged optogenetic proteins (ChR2-YFP) provided a measure of the extent of expression (Figure 1D). Typical optical fibers (230 &#181;m diameter) can readily accommodate 200 nL of silk/AAV. With practice, experimenters can achieve highly reliable expression around the tip of implanted fibers (Figure 5).
To assess expression driven by silk/AAV-coated cranial windows, begin imaging beginning 7-10 days after implantation. We have used two-photon imaging for visualization, but other methods such as fluorescence imaging with a CCD could also be used. Two possible issues with coated cranial windows are insufficient expression, and silk films that fail to dissolve and obscure the field of view. To increase expression, we suggest performing a durectomy prior to implanting the window, and/or increasing the amount of virus in the film. We achieved the best expression using a 1:4 mixture of silk and stock-titer AAV, respectively. While this represents a substantially larger number of viral particles than are usually used in stereotaxic injections, the decreased surgical time counters the marginal additional cost of virus. Meanwhile, if silk films fail to dissolve beneath the window, further reduce the amount of silk used to coat the window. The total amount of silk in coated windows is 10-100 times more than on fiber implants, and the film is less embedded in tissue and thus may not be exposed to the same levels of proteolytic activity than can dissolve silk films13. However, the presence of some silk is essential to achieving expression beneath windows3, likely because a film made of virus alone is washed away by interstitial fluid during surgery.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58728/58728fig1.jpg" /> 
Figure 1: Applying silk/AAV films to optical fibers. (A) Chronic fiber implants are placed fiber side-down into a holder (inset) mounted on a XYZ translator. A fixed microinjector below the fibers dispenses silk/AAV onto the fiber tips. A stereoscope allows visualization of the process. (B) Apply silk/AAV to fiber tips in small volumes (10-20 nL). After ejecting a bolus, retract the pipette and allow ~60 s for the droplet to dry to a flat film. Repeat the process until the required volume has been applied to the fiber tip. (C) Inspect silk coatings. Optimal coatings should be centered on the fiber tip (left), while improper coatings extend outward from the fiber face making them more prone to dislodge from the fiber (right). (D) Representative fibers coated with 200 nL of silk/AAV, and the resulting AAV-driven ChR2-YFP expression 2 weeks after implantation. The compact silk/AAV coating on the left resulted in robust expression, while the coating on the right protruded past the face of the fiber and resulted almost no expression, likely because the silk/AAV did not adhere to the optical fiber during implantation. Scale bars 0.2 mm (fibers) and 1.0 mm (brain slices). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58728/58728fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58728/58728fig2.jpg" /> 
Figure 2: Setup for coating tapered fiber implants. (A) The microinjector is mounted above the tapered fiber holder, and tapered fibers are positioned orthogonally to the ejection syringe. (B) Begin at the widest point (inset) and eject small volumes while moving the ejection syringe towards the point of the taper. This results in a continuous coating along the length of the taper. (C) Representative tapered fiber coated with silk mixed with Fast Green to aid in visualization. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58728/58728fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58728/58728fig3.jpg" /> 
Figure 3: Coating cranial windows. Silk/AAV can be applied cranial windows using a hand pipette. A standard 3 mm diameter window can be coated with a 5 &#181;L droplet, which will slowly dry to a flat film. Inset: GCaMP6f expression resulting from silk/AAV-coated cranial windows implanted with and without durectomies. This figure has been adapted from Jackman et al. (2018)3. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58728/58728fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58728/58728fig4.jpg" /> 
Figure 4: Storing silk/AAV coated implants. (A) To remove residual moisture and preserve viral efficacy, implants should be stored under vacuum at 4 &#176;C until used. Implants stored this way remain viable for at least 7 days. (B) Expression resulting from 4 silk/AAV coated fibers implanted after 7 days of storage. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58728/58728fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58728/58728fig5.jpg" /> 
Figure 5: Silk/AAV-GFP coated optical fibers reliably drive expression. Fluorescent images of slices from 24 consecutive striatal implants. Each implant was coated with 100-400 nL of 1:1 silk/AAV-GFP. This cohort of implants indicates the ability of silk to restrict expression to the implant site (in this case the dorsal striatum). GFP fluorescence is indicated in green; DAPI staining is shown in blue. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58728/58728fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Vivo Targeted Expression of Optogenetic Proteins Using Silk/AAV Films at JoVE.com

In Vivo Targeted Expression of Optogenetic Proteins Using Silk/AAV Films

Here, we present a method for delivering viral expression vectors into the brain using silk fibroin films. This method allows targeted delivery of expression vectors using silk/AAV coated optical fibers, tapered optical fibers, and cranial windows.

In Vivo Targeted Expression of Optogenetic Proteins Using Silk/AAV Films

The quest to understand how neural circuits process information in order to drive behavioral output has been greatly aided by recently-developed optical ...

in-vivo-targeted-expression-optogenetic-proteins-using-silkaav

Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.4K Views.  Oregon Health and Science University. Here, we present a method for delivering viral expression vectors into the brain using silk fibroin films. This method allows targeted delivery of expression vectors using silk/AAV coated optical fibers, tapered optical fibers, and cranial windows.

Video: In Vivo Targeted Expression of Optogenetic Proteins Using Silk/AAV Films

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato

Larval zebrafish are emerging as a model for describing the development and function of simple neural circuits. Due to their external fertilization, rapid development, and translucency, zebrafish are particularly well suited for optogenetic approaches to investigate neural circuit function. In this approach, light-sensitive ion channels are expressed in specific neurons, enabling the experimenter to activate or inhibit them at will and thus assess their contribution to specific behaviors. Applying these methods in larval zebrafish is conceptually simple but requires the optimization of technical details. Here we demonstrate a procedure for expressing a channelrhodopsin variant in larval zebrafish somatosensory neurons, photo-activating single cells, and recording the resulting behaviors. By introducing a few modifications to previously established methods, this approach could be used to elicit behavioral responses from single neurons activated up to at least 4 days post-fertilization (dpf). Specifically, we created a transgene using a somatosensory neuron enhancer, CREST3, to drive the expression of the tagged channelrhodopsin variant, ChEF-tdTomato. Injecting this transgene into 1-cell stage embryos results in mosaic expression in somatosensory neurons, which can be imaged with confocal microscopy. Illuminating identified cells in these animals with light from a 473 nm DPSS laser, guided through a fiber optic cable, elicits behaviors that can be recorded with a high-speed video camera and analyzed quantitatively. This technique could be adapted to study behaviors elicited by activating any zebrafish neuron. Combining this approach with genetic or pharmacological perturbations will be a powerful way to investigate circuit formation and function.

Optogenetic techniques have made it possible to study the contribution of specific neurons to behavior. We describe a method in larval zebrafish for activating single somatosensory neurons expressing a channelrhodopsin variant (ChEF) with a diode-pumped solid state (DPSS) laser and recording the elicited behaviors with a high-speed video camera.

Larval  zebrafish are emerging as a model for describing the development and function  of simple neural circuits. Due to their  external fertilization, ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

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 ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

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 ...