We are grateful to the Canadian Institutes of Health Research (grant MT-5877), the National Institutes of Health (grants NS058668 and NS066664), and the US-Israeli Binational Foundation (grant 2003222) for funding these studies.

Experimental protocols were carried out on female Long Evans rats (250 - 350 g) in accordance with federally prescribed animal care and use guidelines and were approved by the Institutional Animal Care and Use Committee at the University of California San Diego.
1. Acclimating the Rat to Body Restraint
NOTE: Place the rat on a restricted diet. Feed the rat once per day immediately after each daily handling session to acclimate the rat to the restraint (described below). Provide enough feed to maintain the animal at 80% of its initial weight. This amount is approximately 6 grams of feed per day for a 250 g female Long Evans rat.
<ol>
	<li>Acclimate the rat to being handled by humans. Gently restrain the rat by hand for periods of approximately 5 sec every 30 sec. Do this for 15 min a day on 2 consecutive days. Monitor the rats daily for signs of stress during the training period. Signs include struggling in response to restraint, vocalization, and tooth-chattering.</li>
	<li>On the 3rd day, place the rat in a body-constraining cloth sock (Figure 1) for 15 min a day on two consecutive days.</li>
	<li>On the 5th day, place the rat inside the sock, and place the sock inside a rigid tube on the experimental jig (Figure 4). Leave the rat in the tube for 15 min a day on 2 consecutive days.</li>
	<li>Feed the rat immediately after each training session. At the end of the last session (6th day) give unrestricted access to food. Select for implantation only those rats which do not show signs of excessive stress on the last training day. 
	NOTE: Though selection of rats for implantation is subjective, in our hands over 90% of rats were found to be responsive to habituation and able to undergo implantation.</li>
</ol>
2. Implanting the Recording Chamber and Head-restraint Mechanism
<ol>
	<li>Make a reference wire by soldering bare stainless steel wire to a pin connector. Cut the wire so that 3-5 mm remains uncovered by the pin. Autoclave the reference wire, along with all surgical tools.</li>
	<li>Administer ketamine/xylazine anesthesia. Administer 95 mg/kg ketamine mixed with 5 mg/kg xylazine intraperitoneally. The initial dose lasts for 1&#189; to 2 hr. Supplement the anesthesia every 30-45 min thereafter, as needed.&#160;Lubricate the animal&#8217;s eyes with an ophthalmic ointment.</li>
	<li>Check the toe withdrawal reflex to determine the plane of anesthesia, and maintain anesthesia as necessary.</li>
	<li>Place the rat in a stereotaxic holding frame with ear bars (Figure 3A). Shave the hair on the head and disinfect the wound site with povidone-iodine (10% solution). Take care to insert the ear bars appropriately so that they do not damage the ear drum. Ear bars with blunt tips are preferable.</li>
	<li>Make an incision with a scalpel approximately along the midline of the cranium from the rostro-caudal level of the eyes to the back of the ears. Use scissors to cut and remove a 2-3 mm strip of skin on either side of the incision.</li>
	<li>Scrape away the periosteum to expose the surface of the cranium out to the lateral ridges. Cover the exposed cranium with thin layer of superglue.</li>
	<li>Drill a small hole, with a slightly smaller diameter than 0-80 screws, using a &#189; mm diameter drill burr (see Materials section). Drill the hole immediately posterior to where the bregma suture meets the lateral ridge, and at a 30-45&#176; angle into the cranium, so that the screw can go in with the top more lateral than the bottom.</li>
	<li>Screw in a 0-80 flat-bottom machine screw to the hole (approximately 3 turns), at 30-45&#176; angle. Be careful not to screw in too deeply to avoid damaging the underlying brain tissue.</li>
	<li>Repeat this procedure for 6 additional screws in the configuration shown (Figure 3B). Apply superglue to the base of all screws.</li>
	<li>Place a syringe needle in the stereotax manipulator. Measure from the bregma suture, and make a dent in the cranium with the needle near, but outside of the desired craniotomy location, as a reference mark. 
	NOTE: In this demonstration, the stereotaxic coordinates of the mark are 3 mm posterior and 1 mm lateral to the bregma suture.</li>
	<li>Mark the dent with a permanent marker. Note the stereotaxic coordinates of the mark, as it will serve as a stereotaxic reference point (Figure 3B).</li>
	<li>Make a craniotomy centered on the desired coordinates (3 mm posterior and 3 mm lateral to bregma in this example). Leave the dura mater intact. Cover the craniotomy with modified artificial cerebral spinal fluid (125 mM NaCl, 10 mM glucose, 10 mM HEPES, 3.1 mM&#160;CaCl2, 1.3 mM MgCl2, pH 7.4)13 (Figure 3C).</li>
	<li>Cut a 0.2 ml centrifuge tube to 4 - 5 mm in length, and cut off the cap. Place the tube on the cranium and center it over the craniotomy.</li>
	<li>Apply dental cement around the bottom of the tube to seal the base of the tube to the cranium. Be careful not to leak cement into the exposed craniotomy (Figure 3D).</li>
	<li>Drill another small hole (about &#189; mm diameter) in the contralateral cranial plate and carefully insert the reference wire. Construct the reference wire by soldering a pin that interfaces with the recording amplifier to the end of the wire (see Equipment section). Do not move the reference wire once it is in the brain as this may cause damage.</li>
	<li>Apply superglue to the hole in which the wire was inserted. This will seal the pin and wire in place temporarily.</li>
	<li>Mix the two parts of the silicone gel kit (see Materials section) in approximately equal portions. Wait two minutes for mixing and fill the centrifuge tube approximately &#8531; full with gel.</li>
	<li>Attach a right angle post clamp (see Materials section) to the head restraint bar. Attach the head plate (Figure 2A) to the holding bar (Figure 2B) at a 45&#176; pitch angle so that the bottom of the plate is toward the nose of the animal. It is helpful to use an inclinometer to set the pitch angle to match that of the experimental jig (Figure 4).</li>
	<li>Attach the holding bar to the stereotax manipulator arm so that the bar is parallel to the ear bars. Lower the bar and plate so that the plate is posterior of the lambda suture and anterior to the caudal-most screw.</li>
	<li>Grasp the front head-bolt (an 8-32 stud or screw, see Materials section) at approximately 45&#176; angle with a helping hands arm and clamp, so that the head of the screw faces down and towards the tail of the animal. Lower the screw so that it touches the anterior 0-80 screws (Figure 3F, G).</li>
	<li>Secure the bolt and plate in place with dental acrylic. Apply dental acrylic around the bone screw heads, reference pin, and around the sides of the centrifuge tube. Wait approximately 10 min for the dental acrylic to dry (Figure 3H).</li>
	<li>Apply a layer of dental cement around the edges of the dental acrylic, cementing the skin to the implant. Wait for the cement to dry.</li>
	<li>Poke several holes in the cap of the centrifuge tube, and place the cap on the tube.</li>
	<li>Remove the helping hands clamp. Then carefully remove the head-holding bar from the stereotax, and then remove head plate from the bar (Figure 3I).</li>
	<li>Remove the animal from the stereotax and administer post-operative care and monitoring in accordance with all applicable rules, regulations, and laws (e.g.,&#160;Buprenorphine 0.02 mg/kg, every 8-12 hr for a minimum of 24 hr).&#160;If inflammation develops around the edges of the implant, apply a thin layer of antibiotic ointment to the affected site daily until resolved.</li>
</ol>
3. Juxtacellular Monitoring of Neuronal Units
<ol>
	<li>Pull quartz capillary tubing on a carbon dioxide laser micropipette puller (see Equipment section) with tip diameters of less than 1 &#956;m (Figure 5A). 
	NOTE: the heating parameters will vary according to the particular instrument, and that the required neck diameter will vary depending on the brain structure of interest. For recordings in rat thalamus, micropipettes have necks ranging from 5-7 mm.</li>
	<li>Secure the pipette in place under a differential interference contrast (DIC) microscope equipped with long working distance objectives. Use modeling clay to hold the pipette in place on the microscope&#8217;s stage.</li>
	<li>Slowly move a glass block (0.5 - 1 cm thick; a piece of glass used for making ultra-microtome knives is convenient) into the field of view with the pipette tip. Using the stage micromanipulator, gently touch the pipette tip to the glass, causing it to break. Repeat as necessary until the pipette tip outer diameter is between 1-3 &#956;m (Figure 5B,C). Ensure that these micropipettes have impedances between approximately 5-15 M&#937;.</li>
	<li>Prepare extracellular saline (135 mM NaCl, 5.4 mM KCl, 1.8 mM CaCl2, 1 mM MgCl2, 5 mM HEPES, pH 7.2 with NaOH)8 and add 2% (w/v) Chicago Sky Blue. Using a syringe with a 30 G needle or smaller, fill the back end of the pipette with the solution. Alternatively, for single cell juxtacellular labeling add 2% (w/v) Neurobiotin or biotinylated dextran amine (BDA-3000 or BDA-10000) to the saline solution in place of Chicago Sky Blue.</li>
	<li>Place the rat inside the cloth sock inside the rigid tube on an appropriate experimental jig (Figure 4).Wait a minimum of 72 hr after surgery before placing the animal on the head-restraint jig. Tighten the drawstring around the upper sternum, rather than the neck, to avoid obstruction of the airway. If breathing appears to be labored or obstructed, loosen the drawstring or move it posteriorly. Carry out this procedure while the rat is alert.</li>
	<li>Attach the head-restraining plate on the rat to the corresponding piece on the jig. To do this, first pin the head-restraint plate in place, and then secure it using a 4-40 screw. Be sure to wait until the animal is calm in order to avoid applying excessive torque to the head. Next, place an 8-32 nut on the head-restraining bolt that is implanted on the rat. Then screw in a threaded stainless-steel rod to the head-bolt. Affix the rod to the experimental jig in such a way that it can be tightened in place (Figure 4). 
	NOTE: Securing the animal with head-bolt as well as the head-restraint plate minimizes bending of the apparatus and improves recording stability. In most cases recording can commence on the first day the animal is head-restrained. However, if the animal fidgets excessively or vocalizes, provide one day of head-restraint habituation training before commencing any recordings. In this case, leave the animal on the jig for 15 min and then place it back in its cage. Repeat this step the following day and continue with the protocol.</li>
	<li>Open the recording chamber and remove the silicone gel. Clean the craniotomy using fine forceps if tissue has re-grown in the craniotomy.</li>
	<li>Attach the pipette to the motorized micromanipulator, and plug in the headstage pre-amplifier. 
	NOTE: In the present demonstration, a small relay circuit on the headstage is used to switch between the amplifier lead wires and an external current source with high compliance (Figure 6A-C). Note that some amplifiers have an appropriate built-in high compliance source (see Discussion and Materials sections).</li>
	<li>Use the motorized micromanipulator to move the tip of the pipette to the stereotaxically identified mark in the recording chamber. Note the coordinates of this location. Be careful not to break the tip of the pipette on the cranium.</li>
	<li>Move the pipette to the desired recording location in the anterior-posterior and medio-lateral axes. Then advance the pipette ventrally through the dura until it is approximately 500 &#956;m dorsal to the intended recording location.</li>
	<li>Slowly advance the pipette while listening for spiking events on an audio monitor of the amplified voltage recorded between the pipette tip and the reference wire. Once spiking events are identified, continue to move the pipette 0-100 &#956;m until positive going voltage deflections greater than approximately 500 &#956;V are observed. 
	NOTE: The electrode resistance will increase by a factor of approximately 1.5-10 when this occurs.</li>
	<li>Begin recording once a unit has been isolated as according to step 3.11, along with all other behavioral and physiological measures of interest. In the present demonstration, self-generated vibrissa movements are monitored with high speed videography (see Materials section).</li>
	<li>To label the recording site, switch the electrode leads to connect to the current source. There are several ways to do this, using either a relay circuit (Figure 6A-C), a built-in current source on the amplifier, or manually (Figure 6D, see step 3.8 and Discussion). Pass -4 &#956;A with 2 sec pulses at 50% duty cycle for 4 min to iontophoretically inject Chicago Sky Blue through the pipette.</li>
	<li>Kill and perfuse the animal after several labeling procedures according to standard practice. Section the brain and counterstain as necessary to identify the anatomical location of the recording site. Counterstain the tissue for cytochrome oxidase reactivity as per14. Alternatively, identify the Chicago Sky Blue deposits using fluorescence microscopy. 
	NOTE: To accurately differentiate between different labeled units it is important that all labels are made with the same pipette on the same day, without unclamping the pipette between labels. In this case recording sites can be differentiated by their relative locations in post-hoc histology along with the noted manipulator coordinates of each site (see step 3.9). For unambiguous identification, mark the labels at least 200 &#956;m apart from one another, and no more than 3-5 labels per brain region.</li>
</ol>

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The authors declare that they have no competing financial interests.

Construction of the experimental jig
The description of the mechanical parts used to build the experimental jig (Figure 4) is omitted from the protocol, as it can be constructed in a variety of ways. In this demonstration standard opto-mechanical parts and support clamps are used to mount the head restraint bar and the body restraint tube (see Materials section). Similar opto-mechanical parts can be used to mount the electrode ho...

Monitoring neuronal activity in an alert animal actively engaged in a behavioral task is critical for understanding the function and organization of the nervous system. Extracellular recording of the electrical activity from single neuronal units has long been a staple tool of systems neuroscience and is still widely in use at present. A variety of electrode types and configurations are available depending on the scientific and technical demands of a particular experiment. Chronically implanted microdrives or electrode arrays are often used in freely moving animals, including birds, rodents, and non-human primates1-4. Alternatively, acute penetrations with metal or glass microelectrodes via an external micromanipulator are often used to record from anesthetized or head-restrained animals. Glass micropipette electrodes have the advantage that they can be used in the juxtacellular or &#8220;cell attached&#8221; configuration to unambiguously isolate the activity of single neurons without the complications of post-hoc spike sorting5. These electrodes further permit recording from anatomically-identified cells or locations, as they can be used to inject small deposits of dye or neuroanatomical tracers, or even to fill the individual recorded cell. This configuration has been successfully applied in rats, mice and birds6-10. The presently described technique focuses on juxtacellular monitoring and extracellular dye deposits in alert, head-restrained rats. Note that unlike single cell juxtacellular fills, these dye deposits do not provide information about cell morphology or axonal projections11, but they enable exact anatomical localization to approximately 50 &#956;m and, critically, have a significantly higher yield in alert animals. Information regarding single-cell juxtacellular fills is nonetheless provided as an alternative strategy for anatomical labeling.
In brief, the protocol consists of three major phases. In the first phase, the rat is acclimated to body restraint in a cloth sock (Figure 1) over a period of 6 days. In the second phase, a head restraint apparatus (Figure 2) and recording chamber are surgically implanted such that the rat can be maintained in the stereotaxic plane during multiple subsequent recording sessions (Figure 3); this procedure enables the experimenter to target particular sub-cortical regions of the brain for electrophysiological study based on standard reference coordinates12. The third phase involves placing the rat in an appropriate jig for conducting the behavioral and electrophysiological experiments (Figure 4), constructing the electrode from a quartz capillary tube (Figure 5), making juxtacellular neuronal recordings that unambiguously isolate single units6-9, and marking the anatomical location of the recording site with Chicago Sky Blue dye (Figures 6 and 7). The recordings are performed with simultaneous behavioral monitoring; however, the technical details of the behavior will depend on the scientific goals of each experiment and are thus beyond the scope of a single protocol. After completion of the experimental procedure, which can be repeated on multiple days, the animal is euthanized. The brain is extracted and processed according to standard neuroanatomical techniques using either bright field or fluorescence microscopy.

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		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:191px;">Silicone Gel</td>
			<td style="width:179px;">Dow Corning</td>
			<td align="right" style="width:175px;">Mar-80</td>
			<td style="width:315px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:191px;">Jet denture repair acrylic powder</td>
			<td style="width:179px;">Lang Dental Manufacturing Co.</td>
			<td style="width:175px;">N/A</td>
			<td style="width:315px;">For securing the head restraint apparatus to the cranium</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:191px;">Ortho-Jet Fast curing orthodontic acrylic resin liquid</td>
			<td style="width:179px;">Lang Dental Manufacturing Co.</td>
			<td style="width:175px;">N/A</td>
			<td style="width:315px;">For securing the head restraint apparatus to the cranium</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:191px;">Chicago sky blue</td>
			<td style="width:179px;">Sigma</td>
			<td style="width:175px;">C8679</td>
			<td style="width:315px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:191px;">Paraformaldehyde</td>
			<td style="width:179px;">Sigma</td>
			<td align="right" style="width:175px;">158127</td>
			<td style="width:315px;">For perfusion and tissue fixation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:191px;">Phosphate-buffered saline</td>
			<td style="width:179px;">Sigma</td>
			<td style="width:175px;">P3813</td>
			<td style="width:315px;">For perfusion and tissue fixation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:191px;">Cytochrome C</td>
			<td style="width:179px;">Sigma</td>
			<td style="width:175px;">C2506</td>
			<td style="width:315px;">For cytochrome-oxidase staining (Figure 7)</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Diaminobenzidine</td>
			<td style="width:179px;">Sigma</td>
			<td style="width:175px;">D5905</td>
			<td style="width:315px;">For cytochrome-oxidase staining (Figure 7)</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Material Name</td>
			<td style="width:179px;">Company</td>
			<td style="width:175px;">Catalogue Number</td>
			<td style="width:315px;">Comments&#160;</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:191px;">Rat sock</td>
			<td style="width:179px;">Sew Elegant (San Diego, CA)</td>
			<td style="width:175px;">N/A</td>
			<td style="width:315px;">Custom made, Figures 1, 4</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">PVC tube 2 &#189;&#8221;</td>
			<td style="width:179px;">U.S. Plastic Co.</td>
			<td align="right" style="width:175px;">34108</td>
			<td style="width:315px;">Figure 4</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:191px;">Subminiature D pins &#38; sockets</td>
			<td style="width:179px;">TE Connectivity</td>
			<td style="width:175px;">205089-1</td>
			<td style="width:315px;">Figure 3</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:191px;">Stainless steel music wire 0.010&#8221; diameter</td>
			<td style="width:179px;">Precision Brand Products, Inc.</td>
			<td align="right" style="width:175px;">21010</td>
			<td style="width:315px;">Figure 3</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Stereotaxic holding frame</td>
			<td style="width:179px;">Kopf Instruments</td>
			<td style="width:175px;">Model 900</td>
			<td style="width:315px;">Figure 3</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Stereotaxic ear bars</td>
			<td style="width:179px;">Kopf Instruments</td>
			<td style="width:175px;">Model 957</td>
			<td style="width:315px;">Figure 3</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Stereotaxic manipulator</td>
			<td style="width:179px;">Kopf Instruments</td>
			<td style="width:175px;">Model 960</td>
			<td style="width:315px;">Figure 3</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">&#189; mm drill burr</td>
			<td style="width:179px;">Henry Schein</td>
			<td style="width:175px;">100-3995</td>
			<td style="width:315px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Quiet-Air dental drill&#160;</td>
			<td style="width:179px;">Midwest Dental</td>
			<td style="width:175px;">393-1600</td>
			<td style="width:315px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:191px;">Stainless steel 0-80 1/8&#8221; screw</td>
			<td style="width:179px;">Fastener superstore</td>
			<td align="right" style="width:175px;">247438</td>
			<td style="width:315px;">Figure 3</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">0.2mL centrifuge tube</td>
			<td style="width:179px;">Fisher Scientific</td>
			<td style="width:175px;">05-407-8A</td>
			<td style="width:315px;">Figure 3</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Custom head-holding bar</td>
			<td style="width:179px;">UCSD SIO Machine Shop</td>
			<td style="width:175px;">N/A</td>
			<td style="width:315px;">Custom made, Figures 2, 3, 4</td>
		</tr>
		<tr height="20">
			<td height="41" rowspan="2" style="height:41px;width:191px;">Custom head-holding plate</td>
			<td rowspan="2" style="width:179px;">UCSD SIO Machine Shop</td>
			<td rowspan="2" style="width:175px;">N/A</td>
			<td rowspan="2" style="width:315px;">Custom made, Figure 2, 3, 4</td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:191px;">Right angle post-clamp</td>
			<td style="width:179px;">Newport</td>
			<td style="width:175px;">MCA-1</td>
			<td style="width:315px;">Figure 3,4; standard opto-mechanical parts for the experimental jig (Figure 4) are also from Newport Corp.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">8-32 3/4&#8221; screw</td>
			<td style="width:179px;">Fastener Superstore</td>
			<td align="right" style="width:175px;">240181</td>
			<td style="width:315px;">For head-restraint, Figure 3</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">4-40 &#188;&#8221; screw</td>
			<td style="width:179px;">Fastener Superstore</td>
			<td align="right" style="width:175px;">239958</td>
			<td style="width:315px;">For head restraint, Figures 3, 4</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Quartz capillary tubing</td>
			<td style="width:179px;">Sutter Instruments</td>
			<td style="width:175px;">QF-100-60-10</td>
			<td style="width:315px;">Figure 5</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Carbon dioxide laser puller</td>
			<td style="width:179px;">Sutter instruments</td>
			<td style="width:175px;">P-2000</td>
			<td style="width:315px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:191px;">Motorized micromanipulator</td>
			<td style="width:179px;">Sutter Instruments</td>
			<td style="width:175px;">MP-285</td>
			<td style="width:315px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:191px;">Microelectrode amplifier</td>
			<td style="width:179px;">Molecular Devices</td>
			<td style="width:175px;">Multiclamp 700B</td>
			<td style="width:315px;">Alternate part: Molecular Devices Axoclamp 900A</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:191px;">Microelectrode amplifier head stage</td>
			<td style="width:179px;">Molecular Devices</td>
			<td style="width:175px;">CV-7B</td>
			<td style="width:315px;">Alternate part: HS-9Ax10 with Molecular Devices Axoclamp 900A&#160;</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:191px;">Isolated pulse stimulator</td>
			<td style="width:179px;">A-M Systems</td>
			<td style="width:175px;">Model 2100</td>
			<td style="width:315px;">Alternate part: HS-9Ax10 with Molecular Devices Axoclamp 900A</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Audio monitor</td>
			<td style="width:179px;">Radio Shack</td>
			<td style="width:175px;">32-2040</td>
			<td style="width:315px;"></td>
		</tr>
		<tr height="21">
			<td height="43" rowspan="2" style="height:43px;width:191px;">Pipette holder</td>
			<td rowspan="2" style="width:179px;">Warner Instruments</td>
			<td rowspan="2" style="width:175px;">#MEW-F10T</td>
			<td style="width:315px;">Alternate parts: see Discussion</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:315px;">Figure 6A</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Electrode lead wire</td>
			<td style="width:179px;">Cooner wire</td>
			<td style="width:175px;">NEF34-1646</td>
			<td style="width:315px;">(optional), Figure 6D</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:191px;">Relay for amplifier head-stage</td>
			<td style="width:179px;">COTO Technology</td>
			<td style="width:175px;">#2342-05-000</td>
			<td style="width:315px;">(optional) Used with a custom-made printed circuit board (UCSD Physics Electronics Shop), Figure 6A-C</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:191px;">Digital video camera</td>
			<td style="width:179px;">Basler</td>
			<td style="width:175px;">A602fm</td>
			<td style="width:315px;">(optional) For behavioral monitoring, Figure 7</td>
		</tr>
	</tbody>
</table>

juxtacellular monitoring and localization of single neurons within sub-cortical brain structures of alert, head-restrained rats

There are a variety of techniques to monitor extracellular activity of single neuronal units. However, monitoring this activity from deep brain structures in behaving animals remains a technical challenge, especially if the structures must be targeted stereotaxically. This protocol describes convenient surgical and electrophysiological techniques that maintain the animal&#8217;s head in the stereotaxic plane and unambiguously isolate the spiking activity of single neurons. The protocol combines head restraint of alert rodents, juxtacellular monitoring with micropipette electrodes, and iontophoretic dye injection to identify the neuron location in post-hoc histology. While each of these techniques is in itself well-established, the protocol focuses on the specifics of their combined use in a single experiment. These neurophysiological and neuroanatomical techniques are combined with behavioral monitoring. In the present example, the combined techniques are used to determine how self-generated vibrissa movements are encoded in the activity of neurons within the somatosensory thalamus. More generally, it is straightforward to adapt this protocol to monitor neuronal activity in conjunction with a variety of behavioral tasks in rats, mice, and other animals. Critically, the combination of these methods allows the experimenter to directly relate anatomically-identified neurophysiological signals to behavior.

Neuronal units in ventral posterior medial (VPM) thalamus encode the phase of vibrissa movement during self-generated whisking15,16. Figure 7A shows sample spiking activity of a VPM thalamic unit as a rat is actively whisking. Figure 7B shows a histogram of spike times aligned to the instantaneous phase of vibrissa motion17. There are more spikes during the retraction phase of whisking. After the recording, the location of the unit was labeled via iontophoresis of C...

Watch this Scientific Journal Video about Juxtacellular Monitoring and Localization of Single Neurons within Sub-cortical Brain Structures of Alert, Head-restrained Rats at JoVE.com

Juxtacellular Monitoring and Localization of Single Neurons within Sub-cortical Brain Structures of Alert, Head-restrained Rats

This protocol describes the design and surgical implantation of a head-restraining mechanism to monitor neuronal activity in sub-cortical brain structures in alert rats. It delineates procedures to isolate single neurons in the juxtacellular configuration and to efficiently identify their anatomical locations.

There are a variety of techniques to monitor extracellular activity of single neuronal units. However, monitoring this activity from deep brain structures ...

juxtacellular-monitoring-localization-single-neurons-within-sub

Research

JoVE Journal

Neuroscience

11.3K vues.  University of California, San Diego. This protocol describes the design and surgical implantation of a head-restraining mechanism to monitor neuronal activity in sub-cortical brain structures in alert rats. It delineates procedures to isolate single neurons in the juxtacellular configuration and to efficiently identify their anatomical locations.

Vidéo: Juxtacellular Monitoring and Localization of Single Neurons within Sub-cortical Brain Structures of Alert, Head-restrained Rats

Single-cell Profiling of Developing and Mature Retinal Neurons

Highly specialized, but exceedingly small populations of cells play important roles in many tissues. The identification of cell-type specific markers and gene expression programs for extremely rare cell subsets has been a challenge using standard whole-tissue approaches. Gene expression profiling of individual cells allows for unprecedented access to cell types that comprise only a small percentage of the total tissue1-7. In addition, this technique can be used to examine the gene expression programs that are transiently expressed in small numbers of cells during dynamic developmental transitions8.
This issue of cellular diversity arises repeatedly in the central nervous system (CNS) where neuronal connections can occur between quite diverse cells9. The exact number of distinct cell types is not precisely known, but it has been estimated that there may be as many as 1000 different types in the cortex itself10. The function(s) of complex neural circuits may rely on some of the rare neuronal types and the genes they express. By identifying new markers and helping to molecularly classify different neurons, the single-cell approach is particularly useful in the analysis of cell types in the nervous system. It may also help to elucidate mechanisms of neural development by identifying differentially expressed genes and gene pathways during early stages of neuronal progenitor development.
As a simple, easily accessed tissue with considerable neuronal diversity, the vertebrate retina is an excellent model system for studying the processes of cellular development, neuronal differentiation and neuronal diversification. However, as in other parts of the CNS, this cellular diversity can present a problem for determining the genetic pathways that drive retinal progenitors to adopt a specific cell fate, especially given that rod photoreceptors make up the majority of the total retinal cell population11. Here we report a method for the identification of the transcripts expressed in single retinal cells (Figure 1). The single-cell profiling technique allows for the assessment of the amount of heterogeneity present within different cellular populations of the retina2,4,5,12. In addition, this method has revealed a host of new candidate genes that may play role(s) in the cell fate decision-making processes that occur in subsets of retinal progenitor cells8. With some simple adjustments to the protocol, this technique can be utilized for many different tissues and cell types.

A method for the isolation of single retinal cells and subsequent amplification of their cDNAs is described. Single-cell transcriptomics reveals the degree of cellular heterogeneity present in a tissue and uncovers new marker genes for rare cell populations. The accompanying protocol can be adjusted to suit many different cell types.

Highly specialized, but exceedingly small populations of cells play important roles in many tissues. The identification of cell-type specific markers and ...

Laser Capture Microdissection of Enriched Populations of Neurons or Single Neurons for Gene Expression Analysis After Traumatic Brain Injury

Long-term cognitive disability after TBI is associated with injury-induced neurodegeneration in the hippocampus-a region in the medial temporal lobe that is critical for learning, memory and executive function.1,2 Hence our studies focus on gene expression analysis of specific neuronal populations in distinct subregions of the hippocampus. The technique of laser capture microdissection (LCM), introduced in 1996 by Emmert-Buck, et al.,3 has allowed for significant advances in gene expression analysis of single cells and enriched populations of cells from heterogeneous tissues such as the mammalian brain that contains thousands of functional cell types.4 We use LCM and a well established rat model of traumatic brain injury (TBI) to investigate the molecular mechanisms that underlie the pathogenesis of TBI. Following fluid-percussion TBI, brains are removed at pre-determined times post-injury, immediately frozen on dry ice, and prepared for sectioning in a cryostat. The rat brains can be embedded in OCT and sectioned immediately, or stored several months at -80 &deg;C before sectioning for laser capture microdissection. Additionally, we use LCM to study the effects of TBI on circadian rhythms. For this, we capture neurons from the suprachiasmatic nuclei that contain the master clock of the mammalian brain. Here, we demonstrate the use of LCM to obtain single identified neurons (injured and degenerating, Fluoro-Jade-positive, or uninjured, Fluoro-Jade-negative) and enriched populations of hippocampal neurons for subsequent gene expression analysis by real time PCR and/or whole-genome microarrays. These LCM-enabled studies have revealed that the selective vulnerability of anatomically distinct regions of the rat hippocampus are reflected in the different gene expression profiles of different populations of neurons obtained by LCM from these distinct regions. The results from our single-cell studies, where we compare the transcriptional profiles of dying and adjacent surviving hippocampal neurons, suggest the existence of a cell survival rheostat that regulates cell death and survival after TBI.

We describe how to use laser capture microdissection (LCM) to obtain enriched populations of hippocampal neurons or single neurons from frozen sections of the injured rat brain for subsequent gene expression analysis using quantitative real time PCR and/or whole-genome microarrays.

Long-term cognitive disability after TBI is associated with injury-induced neurodegeneration in the hippocampus-a region in the medial temporal lobe that ...

Imaging Neurons within Thick Brain Sections Using the Golgi-Cox Method

The Golgi-Cox method of neuron staining has been employed for more than two hundred years to advance our understanding of neuron morphology within histological brain samples. While it is preferable from a practical perspective to prepare brain sections at the greatest thickness possible, in order to increase the probability of identifying stained neurons that are fully contained within single sections, this approach is limited from a technical perspective by the working distance of high-magnification microscope objectives. We report here a protocol to stain neurons using the Golgi-Cox method in mouse brain sections that are cut at 500 &#956;m thickness, and to visualize neurons throughout the depth of these sections using an upright microscope fitted with a high-resolution 30X 1.05 N.A. silicone oil-immersion objective that has an 800 &#956;m working distance. We also report two useful variants of this protocol that may be employed to counterstain the surface of mounted brain sections with the cresyl violet Nissl stain, or to freeze whole brains for long-term storage prior to sectioning and final processing. The main protocol and its two variants produce stained thick brain sections, throughout which full neuron dendritic&#160;trees and dendrite spines may be reliably visualized and quantified.

We present a protocol for using the Golgi-Cox staining method in thick brain sections, in order to visualize neurons with long dendritic&#160;trees contained within single tissue samples. Two variants of this protocol are also presented that involve cresyl violet counterstaining, and the freezing of unprocessed brains for long-term storage.

The Golgi-Cox method of neuron staining has been employed for more than two hundred years to advance our understanding of neuron morphology within ...

Modeling Amyloid-&#946;42 Toxicity and Neurodegeneration in Adult Zebrafish Brain

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to synaptic degeneration, inflammation, neuronal death, and learning deficits. Humans cannot regenerate lost neurons in the case of AD in part due to impaired proliferative capacity of the neural stem/progenitor cells (NSPCs) and reduced neurogenesis. Therefore, efficient regenerative therapies should also enhance the proliferation and neurogenic capacity of NSPCs. Zebrafish (Danio rerio) is a regenerative organism, and we can learn the basic molecular programs with which we could design therapeutic approaches to tackle AD. For this reason, the generation of an AD-like model in zebrafish was necessary. Using our methodology, we can introduce synthetic derivatives of A&#946;42 peptide with tissue penetrating capability into the adult zebrafish brain, and analyze the disease pathology and the regenerative response. The advantage over the existing methods or animal models is that zebrafish can teach us how a vertebrate brain can naturally regenerate, and thus help us to treat human neurodegenerative diseases better by targeting endogenous NSPCs. Therefore, the amyloid-toxicity model established in the adult zebrafish brain may open new avenues for research in the field of neuroscience and clinical medicine. Additionally, the simple execution of this method allows for cost-effective and efficient experimental assessment. This manuscript describes the synthesis and injection of A&#946;42 peptides into zebrafish brain.

Modeling Amyloid-&amp;#946;42 Toxicity and Neurodegeneration in Adult Zebrafish Brain

This protocol describes the synthesis, characterization, and injection of monomeric amyloid-&#946;42 peptides for generating amyloid toxicity in adult zebrafish to establish an Alzheimer&#39;s disease model, followed by histological analyses and detection of aggregations.

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to ...

Transcranial Electrical Brain Stimulation in Alert Rodents

Transcranial electrical brain stimulation can modulate cortical excitability and plasticity in humans and rodents. The most common form of stimulation in humans is transcranial direct current stimulation (tDCS). Less frequently, transcranial alternating current stimulation (tACS) or transcranial random noise stimulation (tRNS), a specific form of tACS using an electrical current applied randomly within a pre-defined frequency range, is used. The increase of noninvasive electrical brain stimulation research in humans, both for experimental and clinical purposes, has yielded an increased need for basic, mechanistic, safety studies in animals. This article describes a model for transcranial electrical brain stimulation (tES) through the intact skull targeting the motor system in alert rodents. The protocol provides step-by-step instructions for the surgical set-up of a permanent epicranial electrode socket combined with an implanted counter electrode on the chest. By placing a stimulation electrode into the epicranial socket, different electrical stimulation types, comparable to tDCS, tACS, and tRNS in humans, can be delivered. Moreover, the practical steps for tES in alert rodents are introduced. The applied current density, stimulation duration, and stimulation type may be chosen depending on the experimental needs. The caveats, advantages, and disadvantages of this set-up are discussed, as well as safety and tolerability aspects.

This protocol describes a surgical set-up for a permanent epicranial electrode socket and an implanted chest electrode in rodents. By placing a second electrode into the socket, different types of transcranial electrical brain stimulation can be delivered to the motor system in alert animals through the intact skull.

Transcranial electrical brain stimulation can modulate cortical excitability and plasticity in humans and rodents. The most common form of stimulation in ...

Isolation and RNA Extraction of Neurons, Macrophages and Microglia from Larval Zebrafish Brains

To gain a detailed understanding of the role of different CNS cells during development or the establishment and progression of brain pathologies, it is important to isolate these cells without changing their gene expression profile. The zebrafish model provides a large number of transgenic fish lines in which specific cell types are labelled; for example neurons in the NBT:DsRed line or macrophages/microglia in the mpeg1:eGFP line. Furthermore, antibodies have been developed to stain specific cells, such as microglia with the 4C4 antibody.
Here, we describe the isolation of neurons, macrophages and microglia from larval zebrafish brains. Central to this protocol is the avoidance of an enzymatic tissue digestion at 37 &#176;C, which could modify cellular profiles. Instead a mechanical system of tissue homogenization at 4 &#176;C is used. This protocol entails homogenization of brains into cell suspension, their immuno-staining and the isolation of neurons, macrophages and microglia by FACS. Afterwards, we extracted RNA from those cells and evaluated their quality/quantity. We managed to obtain RNA of high quality (RNA Integrity Number (RIN) &#62; 7) to perform qPCR on macrophages/microglia and neurons, and transcriptomic analysis on microglia. This approach enables a better characterization of these cells, as well as a clearer understanding about their role in development and pathologies.

We present a protocol to isolate neurons, macrophages and microglia from larval zebrafish brains under physiological and pathological conditions. Upon isolation, RNA is extracted from these cells to analyze their gene expression profile. This protocol allows for the collection of high-quality RNA for performing downstream analysis like qPCR and transcriptomics.

To gain a detailed understanding of the role of different CNS cells during development or the establishment and progression of brain pathologies, it is ...

Localization of the Locus Coeruleus in the Mouse Brain

The locus coeruleus (LC) is a major hub of norepinephrine producing neurons that modulate a number of physiological functions. Structural or functional abnormalities of LC impact several brain regions including cortex, hippocampus, and cerebellum and may contribute to depression, bipolar disorder, anxiety, as well as Parkinson disease and Alzheimer disease. These disorders are often associated with metal misbalance, but the role of metals in LC is only partially understood. Morphologic and functional studies of LC are needed to better understand the human pathologies and contribution of metals. Mice are a widely used experimental model, but the mouse LC is small (~0.3 mm diameter) and hard to identify for a non-expert. Here, we describe a step-by-step immunohistochemistry-based protocol to localize the LC in the mouse brain. Dopamine-&#946;-hydroxylase (DBH), and alternatively, tyrosine hydroxylase (TH), both enzymes highly expressed in the LC, are used as immunohistochemical markers in brain slices. Sections adjacent to LC-containing sections can be used for further analysis, including histology for morphological studies, metabolic testing, as well as metal imaging by X-ray fluorescence microscopy (XFM).

The locus coeruleus is a small cluster of neurons involved in a variety of physiological processes. Here, we describe a protocol to prepare mouse brain sections for studies of proteins and metals in this nucleus.

The locus coeruleus (LC) is a major hub of norepinephrine producing neurons that modulate a number of physiological functions. Structural or functional ...

A Bedside, Single Burr Hole Approach to Multimodality Monitoring in Severe Brain Injury

Intracranial pressure (ICP) monitoring is a cornerstone of the intensive care management of patients with severe acute brain injuries, including traumatic brain injury. While elevations in ICP are common, data regarding the measurement and treatment of these ICP elevations are conflicting. There is increasing recognition that changes in the balance between supply and demand of brain tissue are critically important and therefore the measurement of multiple modalities is required. Approaches are not standard, and therefore this article provides a description of a bedside, single burr hole approach to multimodality monitoring that allows the passage of probes designed to measure not only ICP but brain tissue oxygen, blood flow, and intracranial electroencephalography. Patient selection criteria, operative procedures, and practical considerations for securing probes during critical care are described. This method is readily performed, safe, secure, and flexible for the adoption of a variety of multimodality monitoring approaches aimed at detecting or preventing secondary brain injuries.

A method of recording multimodality monitoring signals in patients with severe brain injuries using a bedside, single burr hole technique is described.

Intracranial pressure (ICP) monitoring is a cornerstone of the intensive care management of patients with severe acute brain injuries, including traumatic ...

Three-Dimensional Shape Modeling and Analysis of Brain Structures

Statistical shape analysis of brain structures has been used to investigate the association between their structural changes and pathological processes. We have developed a software package for accurate and robust shape modeling and group-wise analysis. Here, we introduce an pipeline for the shape analysis, from individual 3D shape modeling to quantitative group shape analysis. We also describe the pre-processing and segmentation steps using open software packages. This practical guide would help researchers save time and effort in 3D shape analysis on brain structures.

We introduce a semi-automatic protocol for shape analysis on brain structures, including image segmentation using open software, and further group-wise shape analysis using an automated modeling package. Here, we demonstrate each step of the 3D shape analysis protocol with hippocampal segmentation from brain MR images.

Statistical shape analysis of brain structures has been used to investigate the association between their structural changes and pathological processes. ...

Focused Ultrasound Induced Blood-Brain Barrier Opening for Targeting Brain Structures and Evaluating Chemogenetic Neuromodulation

Acoustically Targeted Chemogenetics (ATAC) allows for the noninvasive control of specific neural circuits. ATAC achieves such control through a combination of focused ultrasound (FUS) induced blood-brain barrier opening (FUS-BBBO), gene delivery with adeno-associated viral (AAV) vectors, and activation of cellular signaling with engineered, chemogenetic, protein receptors and their cognate ligands. With ATAC, it is possible to transduce both large and small brain regions with millimeter precision using a single noninvasive ultrasound application. This transduction can later allow for a long-term, noninvasive, device-free neuromodulation in freely moving animals using a drug. Since FUS-BBBO, AAVs, and chemogenetics have been used in multiple animals, ATAC should also be scalable for the use in other animal species. This paper expands upon a previously published protocol and outlines how to optimize the gene delivery with FUS-BBBO to small brain regions with MRI-guidance but without a need for a complicated MRI-compatible FUS device. The protocol, also, describes the design of mouse targeting and restraint components that can be 3D-printed by any lab and can be easily modified for different species or custom equipment. To aid reproducibility, the protocol describes in detail how the microbubbles, AAVs, and venipuncture were used in ATAC development. Finally, an example data is shown to guide the preliminary investigations of studies utilizing ATAC.

This protocol delineates steps necessary for the gene delivery through focused ultrasound blood brain barrier (BBB) opening, evaluation of the resulting gene expression, and measurement of neuromodulation activity of chemogenetic receptors through histological tests.