This study was supported by the National Institute on Drug Abuse grants DA 006886 (MOW) and DA 032270 (DJB).

The utmost care must be taken to maintain aseptic conditions (as described in the Guide for the Care and Use of Laboratory Animals9) while preparing for and conducting the following procedure. The following protocol is in compliance with the Guide for the Care and Use of Laboratory Animals and has been approved by the Institutional Animal Care and Use Committee, Rutgers University. It is estimated that the subsequent procedures will require 3-6 hr to complete.
Implanting the Microwire Array:
<ol>
	<li>Place animals under anesthesia using 50 mg/kg sodium pentobarbital (i.p) and administer 10 mg/kg of atropine methyl nitrate (IP; Glycopyrrolate may be substituted) and 0.25 mg penicillin (300,000 U/ml i.m.) to maintain respiratory function and prevent infection, respectively. 
	Note: With the 3-6 hr length of this implantation surgery, sodium pentobarbital is used because it is cost-effective and limits human exposure to anesthetics (as might occur with prolonged use of gas anesthetics) while still providing long-lasting anesthesia. Substitution of other anesthetics is acceptable.</li>
	<li>Verify that the anesthesia has taken effect using the tail pinch test before proceeding.</li>
	<li>As necessary, give alternating injections of ketamine hydrochloride (60 mg/kg IP) and sodium pentobarbital (5-10 mg/kg IP) to maintain anesthesia throughout the surgery.</li>
	<li>Shave the scalp using a #22 scalpel blade.</li>
	<li>Disinfect the shaved scalp with povidone iodine.</li>
	<li>Give subcutaneous injections of bupivacaine (~1 mg/kg SC spread over 4 injection sites) to locally anesthetize the scalp. Allow 5-10 min for the local anesthetic to take effect.</li>
	<li>Place an ophthalmic lubricant over the eyes to maintain moisture during anesthesia.</li>
	<li>Secure the animal into the ear bars and nose clamp of a stereotaxic apparatus.</li>
	<li>Use visual landmarks (e.g. a horizontal bar on the stereotaxic frame) to approximately level the animal&#39;s head. This step is only meant to approximately level the skull.</li>
	<li>Make an incision along the midline of the scalp using a #11 scalpel blade mounted on a scalpel holder. The incision must extend from just behind the ears to the posterior portion of the nasal bone.</li>
	<li>Using a dissection spatula, clear the skull of all remaining tissue until both lateral skull ridges and the posterior skull ridge have been reached.</li>
	<li>Pull back the skin around the incision using a number of hemostats (6x).</li>
	<li>Clean the skull of any blood and allow it to dry. If any remaining bleeding occurs, terminate the residual bleeding with a small cauterizing tool. Ensuring that the skull remains clean and dry allows the dental acrylic used in subsequent steps to bind to the skull permanently.</li>
	<li>Mark bregma and lambda (Figure 2D) by interpolating the intersection of the skull sutures. A dissecting microscope is necessary to accurately mark these positions.</li>
	<li>Attach a small pointed item (e.g.&#160;a pin or dental drill bit) to a stereotaxic arm and lower it to determine the dorsal/ventral (DV) coordinate of bregma and lambda.</li>
	<li>Adjust the nose clamp until the DV coordinates for bregma and lambda are within 100 &#181;m (0.1 mm) of each other.</li>
	<li>Once leveled, record the anterior/posterior (AP), medial/lateral (ML) and DV coordinates of bregma along with the position of the nose clamp. Double check the coordinates for accuracy, as the remainder of the surgery depends on the precision of these coordinates.</li>
	<li>Use the measured coordinates to calculate the ML and AP coordinates for the four corners of the &#34;skull window&#34; relative to bregma (i.e. craniotomy; Figure 1). The skull window is a precisely positioned rectangle through which the microwire array will pass.</li>
	<li>Use the calculated coordinates to mark the skull window coordinates on the skull using a pointed stereotaxic attachment and fountain pen ink. To obtain precise marks, apply only a small amount of ink to the tip of the marking tool using a cotton applicator.</li>
	<li>Drill out the skull window by removing the bone in a series of small layers.
	<ol>
		<li>Start by drilling the marked corners of the window where the marks have been placed.</li>
		<li>Next, connect the corners and outline the window.</li>
		<li>Finally, clear out the area within the outline down to the depth of dura mater. The hole must be wider (i.e. beveled) below the superficial layers of the skull to ensure that the window maintains an appropriate width from top to bottom.</li>
	</ol>
	</li>
	<li>Use microforceps to carefully remove any remaining bone chips, debris, or dura mater inside the skull window. This step is extremely important, as these bits of material can compromise the integrity of the array during lowering. Once cleared, one must keep the window moist with bacteriostatic saline for the remainder of the surgery.</li>
	<li>Place markings on the skull for 5 skull screws and 1 ground wire (Figure 1). These positions will vary depending on the targeted brain region. The headstage will be most secure if one screw is placed on each of the 5 skull bones. Place both the skull screws and ground wire in locations that will not interfere with the microwire array placement.</li>
	<li>Drill holes for the skull screws and secure the screws in place. Screws must be lowered until only 3-4 threads are showing (or, if using screws other than those recommended, deep enough to traverse the thickness of the skull to promote array stability but not too deep so as to damage cortex). Clean the threads of the screws after placement.</li>
	<li>Drill a hole for the ground wire. Lower the wire slowly (over 1-2 min) to the target DV coordinate and fix the wire in place using dental acrylic.</li>
	<li>Add acrylic around the threads of the skull screws. Before the acrylic dries, remove any excess cement that flows away from the ground wire or skull screws. Allow 15 min for the dental acrylic to dry/harden.</li>
	<li>Attach the array to the stereotaxic arm. Level the array and orient it so that it will squarely pass through the confines of the skull window.</li>
	<li>Place saline in the skull window so that it is level with the skull. Lower the array until it creates a dimple in the saline. This coordinate is used as skull level for the array. Use this value to calculate the final DV coordinate of the array.</li>
	<li>Lower the array slowly until it reaches its final DV coordinate. Stop every 1mm of lowering and wait for several minutes in order to allow brain tissue to recover from dimpling and to dissolve the bottom portion of the polyethylene glycol (PEG) on the array (PEG is used to temporarily keep the wires in their conformation while lowering, and dissolves slowly in saline).</li>
	<li>When the array is 1mm away from the target, lower it more slowly until the final target is reached. Lowering at 0.1-0.2 mm at a time before allowing the tissue to rest for a period of 5 min is intended to preserve the tissue at the target site as well as proximal synaptic connections. If the equipment is available, precision lowering of the array can be assisted using a motorized manipulator.</li>
	<li>Dissolve the remaining PEG and use dental acrylic to cement the microwires in place. Add multiple layers of cement to ensure that the wires are secure and then allow 15-20 min for the cement to harden before proceeding. This ensures that the wires will not be jostled from their final placement.</li>
	<li>Build the remainder of the animal&#39;s headstage using dental acrylic. Use the acrylic to encase the skull screws, ground wire, and connector for the microwires. Allow the acrylic hat to harden sufficiently before proceeding.</li>
	<li>Suture the scalp incision using absorbable sutures and administer 2 ml of bacteriostatic saline (s.c.) to restore hydration from surgery.</li>
	<li>Remove the animal from the ear bars and place the subject in a clean area. Observe the animal frequently during the post-operative recovery until thermoregulation and locomotion have recovered. Following recovery from anesthesia, move the animal into single-housing for the remainder of post-surgical recovery.</li>
	<li>Give animals daily postoperative monitoring and care in the days following the procedure. Recovery from this procedure is optimal when seven or more days are allowed.</li>
	<li>Give animals injections of Carprofen (5mg/kg) and Enrofloxacin (5-10 mg/kg) or their equivalents during recovery per the schedule recommended by the attending veterinarian.</li>
</ol>

<ol>
	<li>Carter, M., Shieh, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Electrophysiology In: Guide to research techniques in neuroscience. , (2009).</li><li>Aston-Jones, G., Siggins, G. R., Kupfer, D., Bloom, F. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Electrophysiology. In: Psychopharmacology: The Fourth Generation of Progress. , (1995).</li><li>Root, D. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+roles+of+ventral+pallidum+subregions+during+cocaine+self-administration+behaviors.">Differential roles of ventral pallidum subregions during cocaine self-administration behaviors.</a> J. Comp. Neurol. 521 (3), 558-588 (2013).</li><li>Cardin, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissecting+local+circuits+in+vivo:+integrated+optogenetic+and+electrophysiology+approaches+for+exploring+inhibitory+regulation+of+cortical+activity.">Dissecting local circuits in vivo: integrated optogenetic and electrophysiology approaches for exploring inhibitory regulation of cortical activity.</a> (3-4), 106-103 (2012).</li><li>Ma, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amphetamine's+dose-dependent+effects+on+dorsolateral+striatum+sensorimotor+neuron+firing.">Amphetamine's dose-dependent effects on dorsolateral striatum sensorimotor neuron firing.</a> Behav. Brain Res. , (2013).</li><li>Ghitza, U. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Persistent+cue-evoked+activity+of+accumbens+neurons+after+prolonged+abstinence+from+self-administered+cocaine.">Persistent cue-evoked activity of accumbens neurons after prolonged abstinence from self-administered cocaine.</a> J. Neurosci. 23 (19), 7239-7245 (2003).</li><li>Tang, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Changes+in+activity+of+the+striatum+during+formation+of+a+motor+habit.">Changes in activity of the striatum during formation of a motor habit.</a> Eur. J. Neurosci. 25 (4), 1212-1227 (2007).</li><li>Tang, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dose+and+rate-dependent+effects+of+cocaine+on+striatal+firing+related+to+licking.">Dose and rate-dependent effects of cocaine on striatal firing related to licking.</a> J. Pharmacol. Exp. Ther. 324 (2), 701-713 (2008).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=National+Research+Council.">National Research Council.</a> Guide for the Care and Use of Laboratory Animals: Eighth Edition. , (2011).</li><li>Fabbricatore, A. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+evidence+of+mediolateral+functional+dichotomy+in+the+rat+accumbens+during+cocaine+self-administration:+tonic+firing+patterns.">Electrophysiological evidence of mediolateral functional dichotomy in the rat accumbens during cocaine self-administration: tonic firing patterns.</a> Eur. J. Neurosci. 30 (12), 2387-2400 (2009).</li><li>Root, D. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Slow+phasic+and+tonic+activity+of+ventral+pallidal+neurons+during+cocaine+self-administration.">Slow phasic and tonic activity of ventral pallidal neurons during cocaine self-administration.</a> Synapse. 66 (2), 106-127 (2012).</li><li>Root, D. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid-phasic+activity+of+ventral+pallidal+neurons+during+cocaine+self-administration.">Rapid-phasic activity of ventral pallidal neurons during cocaine self-administration.</a> Synapse. 64 (9), 704-713 (2010).</li><li>Tang, C. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Decreased+firing+of+striatal+neurons+related+to+licking+during+acquisition+and+overtraining+of+a+licking+task.">Decreased firing of striatal neurons related to licking during acquisition and overtraining of a licking task.</a> J. Neurosci. 29 (44), 12952-12961 .</li><li>Paxinos, G., Watson, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Rat Brain in Stereotaxic Coordinates. , (1997).</li></ol>

The authors have no competing financial interests to disclose.

Extracellular recordings represent a powerful experimental technique that can be incorporated into nearly any experimental preparation in neuroscience. Wires that have been implanted in organized arrays can be tracked as their shafts pass through the brain and into their target region (Figure 5A). When a small, post-experimental lesion is created at the noninsulated microwire tip to create a small iron deposit from the stainless steel wire, one can precisely mark the location of the uninsulated microwire...

Electrophysiological recordings allow scientists to examine the electrical properties of biological cells. In the central nervous system, where electrical impulses serve as a signaling mechanism, these recordings are of particular importance for understanding neural function1-2. During single-unit recordings in behaving animals, a microelectrode that has been inserted into the brain is able to record changes in a neuron&#39;s generation of action potentials over time.
While many techniques allow one to record brain activity, single-unit electrophysiology is one of the most precise methods by allowing resolution at the single neuron level. When a high degree of spatial specificity is desired, microwires can be used to target discrete sub-nuclei or ensembles of cells within the brain3. Single-unit recordings also benefit from high temporal resolution as recordings are accurate at the microsecond level. And, in vivo awake recordings allow intact circuit interactions, with the natural milieu of afferent and efferent projections, systemic chemical and hormonal influences, and physiological parameters. Neural signals are derived from sensory input, motor behaviors, cognitive processing, neurochemistry/pharmacology, or some combination. Accordingly, the segregation of sensory, motor, cognitive, and chemical influences necessitates well-conceived experiments with effective contingencies and controls that may allow for the assessment of each of the aforementioned influences. All in all, recordings in behaving animals allow experimenters to observe the integration of multiple sources of information within a functioning circuit and to derive a more comprehensive model of circuit function.
Single-unit recordings also suffer from a number of disadvantages of which any experimenter should be aware. First and foremost, recordings can be difficult to conduct. Indeed, properties of the headstage amplifiers and the implanted microwires that allow for spatial and temporal specificity in these recordings also makes recordings susceptible to the influence of extraneous electrical signals (i.e. electrical &#34;noise&#34;). Accordingly, the ability to troubleshoot problems in an electrophysiological system necessitates a well-developed technical understanding of electrophysiological principles and apparatus. It is also important to note that, under certain circumstances, recorded electrical signals in extracellular recordings can represent the summation of multiple neural signals. Moreover, the generalizability of single-unit activity to population activity within a target region can often be limited by the degree of cellular heterogeneity within the target region (but see Cardin4). For example, electrodes might be biased towards recording high amplitude output neurons in lieu of other cells. The interpretability of single-unit recordings is increased by combining recordings with other techniques including, but not limited to, electrical (orthodromic or antidromic), chemical (e.g. iontophoretic or designer receptor) or optogenetic stimulation4, temporary neural inactivations, sensorimotor examinations5, disconnection procedures, or immunohistochemistry3.
In the protocol that follows we will enumerate the materials and steps necessary to implant an organized microwire array in the rat (although the protocol can be adapted for use in other species). The procedure and style of fixed arrays used in our laboratory have proven reliable for longitudinal recordings and can sustain recordings of the same neuron for over one month&#39;s time 6-8. This makes this procedure ideal for examining phasic responses to experimental stimuli, plastic changes in neural responses, or mechanisms of learning and motivation.

<table>
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr>
			<td>Table 1. List of Surgical Materials</td>
		</tr>
		<tr>
			<td>Gauze</td>
			<td>Fisher (MooreBrand)</td>
			<td>19-898-144</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton Swabs</td>
			<td>Fisher (Puritan)</td>
			<td>S304659</td>
			<td></td>
		</tr>
		<tr>
			<td>Nembutal (Pentobarbital)</td>
			<td>Sigma Aldrich</td>
			<td>P3761</td>
			<td></td>
		</tr>
		<tr>
			<td>Atropine Methyl Nitrate</td>
			<td>Sigma Aldrich</td>
			<td>A0382</td>
			<td></td>
		</tr>
		<tr>
			<td>Baytril (Enrofloxacin)</td>
			<td>Butler Shein (Bayer)</td>
			<td>1040007</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine Hydrochloride</td>
			<td>Butler Shein</td>
			<td>SKU# 023061</td>
			<td></td>
		</tr>
		<tr>
			<td>Betadine (Povidone-Iodine)</td>
			<td>Fisher (Perdue)</td>
			<td>19-066452&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotax</td>
			<td>Kopf</td>
			<td>Model 900</td>
			<td></td>
		</tr>
		<tr>
			<td>Cauterizing Tool</td>
			<td>Stoelting</td>
			<td>59017</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting Microscope</td>
			<td>Nikon</td>
			<td>SMZ445</td>
			<td></td>
		</tr>
		<tr>
			<td>Dental Drill</td>
			<td>Buffalo</td>
			<td>37800</td>
			<td></td>
		</tr>
		<tr>
			<td>Bacteriostatic Saline</td>
			<td>Bulter Schein</td>
			<td>8973</td>
			<td></td>
		</tr>
		<tr>
			<td>Jewlers Skrews</td>
			<td>Stoelting</td>
			<td>51457</td>
			<td></td>
		</tr>
		<tr>
			<td>Microwire Array</td>
			<td>Microprobes</td>
			<td>Custom (Flexible)</td>
			<td></td>
		</tr>
		<tr>
			<td>Ground Wire</td>
			<td>Omnetics</td>
			<td>Custom Plug</td>
			<td></td>
		</tr>
		<tr>
			<td>Dental Acrylic</td>
			<td>Fisher (BAS)</td>
			<td>50-854-402</td>
			<td></td>
		</tr>
		<tr>
			<td>Absorbable Sutures</td>
			<td>Fisher (Ethicon)</td>
			<td>NC0258473</td>
			<td></td>
		</tr>
		<tr>
			<td>Puralube (Opthalamic Ointment/Lubricant)</td>
			<td>Fisher (Henry Schein)</td>
			<td>008897</td>
			<td></td>
		</tr>
	</tbody>
</table>
<table>
	<tbody>
		<tr>
			<td>Table 2. List of Surgical Instruments</td>
		</tr>
		<tr>
			<td>2x Microforceps</td>
			<td>George Tiemann &#38; Co.</td>
			<td>#160-57</td>
			<td>Multi-use (e.g. clearing debris in skull window)</td>
		</tr>
		<tr>
			<td>2x Forceps</td>
			<td>George Tiemann &#38; Co.</td>
			<td>#160-93</td>
			<td>Multi-use (e.g. tying sutures)</td>
		</tr>
		<tr>
			<td>6x Hemostats</td>
			<td>George Tiemann &#38; Co.</td>
			<td>#105-1125</td>
			<td>Clamp and open incision</td>
		</tr>
		<tr>
			<td>1x Small scissors</td>
			<td>George Tiemann &#38; Co.</td>
			<td>#105-411</td>
			<td>Cut sutures after tying</td>
		</tr>
		<tr>
			<td>1x Tissue forceps</td>
			<td>George Tiemann &#38; Co.</td>
			<td>#105-222</td>
			<td>Holding tissue while suturing</td>
		</tr>
		<tr>
			<td>1x Needle holder</td>
			<td>George Tiemann &#38; Co.</td>
			<td>#105-1259</td>
			<td>Holding suture needle</td>
		</tr>
		<tr>
			<td>1x Scalpel holder (with #11 blade)</td>
			<td>George Tiemann &#38; Co.</td>
			<td>#105-80 (w/ #105-71 blade)</td>
			<td>Making skull incision</td>
		</tr>
		<tr>
			<td>1x # 22 Scalpel blade</td>
			<td>George Tiemann &#38; Co.</td>
			<td># 160-381</td>
			<td>Shaving scalp</td>
		</tr>
		<tr>
			<td>1x Surgical Spatula</td>
			<td>George Tiemann &#38; Co.</td>
			<td>#160-718</td>
			<td>Scraping skull to clear tissue on skull</td>
		</tr>
		<tr>
			<td>Machine/Jewelers Screws</td>
			<td>Various</td>
			<td>N/A</td>
			<td>0/80 x 1/8&#8221;</td>
		</tr>
	</tbody>
</table>
<table>
	<tbody>
		<tr>
			<td>Table 3. List of Equipment for Recording Electrophysiological Signals</td>
		</tr>
		<tr>
			<td>Microwire Array &#38; Connector</td>
			<td>Micro Probe, Inc. (Gaithersburg, MD)</td>
			<td>&#160;N/A</td>
			<td>Cranially implanted in target recording region. Arrays are customized based on desired wire spacing, length, etc.</td>
		</tr>
		<tr>
			<td>(Part No. Based on array characteristics)</td>
		</tr>
		<tr>
			<td>Unity-Gain Harness/Headstage</td>
			<td>M.B. Turnkey Designs (Hillsborough, NJ)</td>
			<td>Proj 1200</td>
			<td>Initial amplification of neural signal; allows for propagation of small neural signals.</td>
		</tr>
		<tr>
			<td>Commutator (&#38; Optional Fluid Swivel)</td>
			<td>Plastics One, Inc. (Roanoke, VA)</td>
			<td>SL18C</td>
			<td>Allows animals to freely rotate while propagating electrical signal to preamp</td>
		</tr>
		<tr>
			<td>Pre-Amplifier</td>
			<td>M.B. Turnkey Designs (Hillsborough, NJ)</td>
			<td>Proj 1198</td>
			<td>Differentially amplifies neural signals against a reference electrode.</td>
		</tr>
		<tr>
			<td>Filter &#38; Amplifier</td>
			<td>M.B. Turnkey Designs (Hillsborough, NJ)</td>
			<td>Proj 1199</td>
			<td>Band-pass filters and further amplifies the differentially amplified signal.</td>
		</tr>
		<tr>
			<td>Acquisition Computer</td>
			<td>EnGen (Phoenix, AZ)</td>
			<td>N/A (Custom Build)</td>
			<td>Runs software and hardware for behavioral and neural data acquisition.</td>
		</tr>
		<tr>
			<td>A/D Card&#160;</td>
			<td>Data Translation (Marlboro, MA)</td>
			<td>DT-3010</td>
			<td>Digitizes neural signals for computer sampling.</td>
		</tr>
		<tr>
			<td>Digital I/O Card</td>
			<td>Measurement Computing (Norton, MA)</td>
			<td>PCI CTR-05</td>
			<td>Acquires behavioral inputs and outputs</td>
		</tr>
	</tbody>
</table>

a procedure for implanting organized arrays of microwires for single-unit recordings in awake, behaving animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

A list of Equipment used by this laboratory for recording electrophysiological signals can be found in Table 3. Following recovery from surgery, single-units are recorded by plugging a unity-gain headstage into the implanted connector. This headstage is connected via a cable to a commutator, which is capable of free rotation without breaks in the electrophysiological recording through the use of electrical slip rings. The commutator allows subjects to freely move while recording during behavior, which is...

Watch this Scientific Journal Video about A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals at JoVE.com

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

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JoVE Journal

Neuroscience

13.2K Views.  Rutgers, the State University of New Jersey. Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges. Methods for performing this technique and the equipment necessary are described. Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

Video: A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

Electrode Fabrication and Implantation in Aplysia californica for Multi-channel Neural and Muscular Recordings in Intact, Freely Behaving Animals

Recording from key nerves and muscles of Aplysia during feeding behavior allows us to study the patterns of neural control in an intact animal. Simultaneously recording from multiple nerves and muscles gives us precise information about the timing of neural activity. Previous recording methods have worked for two electrodes, but the study of additional nerves or muscles required combining and averaging the recordings of multiple animals, which made it difficult to determine fine details of timing and phasing, because of variability from response to response, and from animal to animal. Implanting four individual electrodes has a very low success rate due to the formation of adhesions that prevent animals from performing normal feeding movements. We developed a new method of electrode fabrication that reduces the bulk of the electrodes inside the animal allowing for normal feeding movements. Using a combination of glues to attach the electrodes results in a more reliable insulation of the electrode which lasts longer, making it possible to record for periods as long as a week. The fabrication technique that we describe could be extended to incorporate several additional electrodes, and would be applicable to vertebrate animals.

Electrode Fabrication and Implantation in Aplysia californica for Multi-channel Neural and Muscular Recordings in Intact, Freely Behaving Animals

A technique is described for implanting four in vivo electrodes to monitor the neuromuscular control of feeding behavior in Aplysia californica.

Recording from key nerves and muscles of Aplysia during feeding behavior allows us to study the patterns of neural control in an intact animal.  ...

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

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

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

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

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

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 ...

Design and Assembly of an Ultra-light Motorized Microdrive for Chronic Neural Recordings in Small Animals

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of neural circuits underlying a variety of natural behaviors, including navigation1 , decision making 2,3, and the generation of complex motor sequences4,5,6. Advances in precision machining has allowed for the fabrication of light-weight devices suitable for chronic recordings in small animals, such as mice and songbirds. The ability to adjust the electrode position with small remotely controlled motors has further increased the recording yield in various behavioral contexts by reducing animal handling.6,7
 Here we describe a protocol to build an ultra-light motorized microdrive for long-term chronic recordings in small animals. Our design evolved from an earlier published version7, and has been adapted for ease-of use and cost-effectiveness to be more practical and accessible to a wide array of researchers. This proven design 8,9,10,11 allows for fine, remote positioning of electrodes over a range of ~ 5 mm and weighs less than 750 mg when fully assembled. We present the complete protocol for how to build and assemble these drives, including 3D CAD drawings for all custom microdrive components.

The design, fabrication and assembly of an ultra-light motorized microdrive is described. The device provides a cost-effective and easy-to-use solution for chronic recordings of single units in small behaving animals.

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of ...

Simultaneous Long-term Recordings at Two Neuronal Processing Stages in Behaving Honeybees

In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent and divergent anatomical connections including feed forward and feedback wiring. Furthermore, information of the same origin is partially sent via parallel pathways to different and sometimes into the same brain areas. To understand the evolutionary benefits as well as the computational advantages of these wiring strategies and especially their temporal dependencies on each other, it is necessary to have simultaneous access to single neurons of different tracts or neuropiles in the same preparation at high temporal resolution. Here we concentrate on honeybees by demonstrating a unique extracellular long term access to record multi unit activity at two subsequent neuropiles1, the antennal lobe (AL), the first olfactory processing stage and the mushroom body (MB), a higher order integration center involved in learning and memory formation, or two parallel neuronal tracts2 connecting the AL with the MB.&#160;The latter was chosen as an example and will be described in full. In the supporting video the construction and permanent insertion of flexible multi&#160;channel wire electrodes is demonstrated. Pairwise differential amplification of the micro&#160;wire electrode channels drastically reduces the noise and verifies that the source of the signal is closely related to the position of the electrode tip. The mechanical flexibility of the used wire electrodes allows stable invasive long&#160;term recordings over many hours up to days, which is a clear advantage compared to conventional extra&#160;and intracellular in vivo recording techniques.

Simultaneous extracellular long term recordings from two different brain neuropiles or two different anatomical tracts were established in honeybees. These recordings allow the investigation of temporal aspects of neuronal processing across different brain areas at the single neuron as well as at the ensemble level in a behaving animal.

In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

Investigating Object Representations in the Macaque Dorsal Visual Stream Using Single-unit Recordings

Previous studies have shown that neurons in parieto-frontal areas of the macaque brain can be highly selective for real-world objects, disparity-defined curved surfaces, and images of real-world objects (with and without disparity) in a similar manner as described in the ventral visual stream. In addition, parieto-frontal areas are believed to convert visual object information into appropriate motor outputs, such as the pre-shaping of the hand during grasping. To better characterize object selectivity in the cortical network involved in visuomotor transformations, we provide a battery of tests intended to analyze the visual object selectivity of neurons in parieto-frontal regions.

A detailed protocol to analyze object selectivity of parieto-frontal neurons involved in visuomotor transformations is presented.

Previous studies have shown that neurons in parieto-frontal areas of the macaque brain can be highly selective for real-world objects, disparity-defined ...

Real-Time fMRI Brain Mapping in Animals

Dynamic fMRI responses vary largely according to the physiological conditions of animals either under anesthesia or in awake states. We developed a real-time fMRI platform to guide experimenters to monitor fMRI responses instantaneously during acquisition, which can be used to modify the physiology of animals to achieve desired hemodynamic responses in animal brains. The real-time fMRI set-up is based on a 14.1T preclinical MRI system, enabling the real-time mapping of dynamic fMRI responses in the primary forepaw somatosensory cortex (FP-S1) of anesthetized rats. Instead of a retrospective analysis to investigate confounding sources leading to the variability of fMRI signals, the real-time fMRI platform provides a more effective scheme to identify dynamic fMRI responses using customized macro-functions and a common neuroimage analysis software in the MRI system. Also, it provides immediate troubleshooting feasibility and a real-time biofeedback stimulation paradigm for brain functional studies in animals.

Animal brain functional mapping can benefit from the real-time functional magnetic resonance imaging (fMRI) experimental set-up. Using the latest software implemented in the animal MRI system, we established a real-time monitoring platform for small animal fMRI.

Dynamic fMRI responses vary largely according to the physiological conditions of animals either under anesthesia or in awake states. We developed a ...

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit functions in systems neuroscience. However, high absorbance and scattering of light in mammalian tissue limit intravital imaging mostly to superficial brain regions, leaving deep-brain areas, such as the hippocampus, out of reach for optical microscopy. In this video, we show the preparation and implantation of the custom-made imaging window to enable chronic in vivo imaging of the dorsal hippocampal CA1 region in head-fixed behaving mice. The custom-made window is supplemented with an infusion cannula that allows targeted delivery of viral vectors and drugs to the imaging area. By combining this preparation with wide-field imaging, we performed a long-term recording of neuronal activity using a fluorescent calcium indicator from large subsets of neurons in behaving mice over several weeks. We also demonstrated the applicability of this preparation for voltage imaging with single-spike resolution. High-performance genetically encoded indicators of neuronal activity and scientific CMOS cameras allowed the recurrent visualization of subcellular morphological details of single neurons at high temporal resolution. We also discuss the advantages and potential limitations of the described method and its compatibility with other imaging techniques.

This article demonstrates the preparation of a custom-made imaging window supplemented with infusion cannula and its implantation onto the CA1 region of the hippocampus in mice.

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit ...

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

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

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